Medicine, Faculty of
Medicine, Department of
DSpace
UBCV
Sun, Guobin
Doctor of Philosophy - PhD
University of British Columbia
Focal adhesions (FA) form interfaces between the extracellular matrix and the cytoskeleton to regulate cell responses such as cell proliferation, survival, and migration. The functions and dynamic interactions of many of the ~180 molecules in this integrin-initiated FA complex network are far from fully understood. Among these is the receptor-like protein tyrosine phosphatase PTPα. In addition to the integrin-proximal action of PTPα to catalyze activation of Src family kinases, subsequent phosphorylation of PTPα at its C-terminal Tyr789 site is essential and acts in an unknown manner to promote cell spreading and migration. We used reconstitution assays in PTPα-null cells to identify and distinguish integrin signaling events that were PTPα-dependent and PTPα-Tyr789-dependent. Results show that PTPα-Tyr789 mediates the localization of PTPα and the scaffolding protein Cas to FAs where Cas interacts with and gets phosphorylated by Src to initiate Cas/Crk-Rac/Cdc42-PAK signaling. Linking these events, this study identifies the Cas-binding protein BCAR3 as a molecular connector of PTPα and Cas, with phosphoTyr789-PTPα interacting with the BCAR3 SH2 domain to recruit BCAR3-Cas to newly forming adhesion sites. These findings identify phospho-Tyr789-PTPα as the first cellular ligand for the SH2 domain of BCAR3, and reveal a role of PTPα in integrin-induced adhesion complex assembly that enables Src-mediated activation of the pivotal function of Cas in cell migration. Furthermore I extended this study into a cancer cell model by showing that the disruption of this PTPα-BCAR3-Cas-Src signaling module inhibits the invasive motility in a rhabdomyosarcoma cell line. In summary, my results in this thesis elucidate the molecular mechanism underlying the PTPα-Tyr789-dependent cell spreading and migration, and support that the PTPα-BCAR3-Cas complex is a critical regulator of cell migration as well as cancer cell invasion and metastasis.
THE ROLE OF PROTEIN TYROSINE PHOSPHATASE ALPHA TYROSINE 789 PHOSPHORYLATION IN INTEGRIN SIGNALING by Guobin Sun B.Sc., Nankai University, 1999 M.Sc., Simon Fraser University, 2006 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Experimental Medicine) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) January 2013 © Guobin Sun, 2013 ii Abstract Focal adhesions (FA) form interfaces between the extracellular matrix and the cytoskeleton to regulate cell responses such as cell proliferation, survival, and migration. The functions and dynamic interactions of many of the ~180 molecules in this integrin-initiated FA complex network are far from fully understood. Among these is the receptor-like protein tyrosine phosphatase PTPα. In addition to the integrin-proximal action of PTPα to catalyze activation of Src family kinases, subsequent phosphorylation of PTPα at its C-terminal Tyr789 site is essential and acts in an unknown manner to promote cell spreading and migration. We used reconstitution assays in PTPα-null cells to identify and distinguish integrin signaling events that were PTPα-dependent and PTPα-Tyr789-dependent. Results show that PTPα-Tyr789 mediates the localization of PTPα and the scaffolding protein Cas to FAs where Cas interacts with and gets phosphorylated by Src to initiate Cas/Crk-Rac/Cdc42- PAK signaling. Linking these events, this study identifies the Cas-binding protein BCAR3 as a molecular connector of PTPα and Cas, with phosphoTyr789-PTPα interacting with the BCAR3 SH2 domain to recruit BCAR3-Cas to newly forming adhesion sites. These findings identify phospho-Tyr789-PTPα as the first cellular ligand for the SH2 domain of BCAR3, and reveal a role of PTPα in integrin-induced adhesion complex assembly that enables Src- mediated activation of the pivotal function of Cas in cell migration. Furthermore I extended this study into a cancer cell model by showing that the disruption of this PTPα-BCAR3-Cas- Src signaling module inhibits the invasive motility in a rhabdomyosarcoma cell line. In summary, my results in this thesis elucidate the molecular mechanism underlying the PTPα- Tyr789-dependent cell spreading and migration, and support that the PTPα-BCAR3-Cas complex is a critical regulator of cell migration as well as cancer cell invasion and metastasis. iii Preface The research project presented in this dissertation was originally developed in Dr. Catherine J. Pallen's laboratory. I have participated in experimental design under Dr. Pallen's supervision and conducted all the experiments presented in this thesis. I was also responsible for data collection and analysis, as well as preparation of manuscripts for peer-reviewed journal publication. Publication resulting from this research: 1. Sun G, Cheng SY, Chen M, Lim CJ, Pallen CJ. (2012) Protein Tyrosine Phosphatase α Phosphotyrosyl-789 Binds BCAR3 To Position Cas for Activation at Integrin-Mediated Focal Adhesions. Mol. Cell Biol. 2012 Sep;32(18):3776-89. Epub 2012 Jul 16. Chapter 2,3, and 4 of my thesis contain data that were published in this manuscript. iv Table of Contents Abstract……. ........................................................................................................................... ii Preface……. ............................................................................................................................ iii Table of Contents ................................................................................................................... iv List of Figures ......................................................................................................................... ix List of Abbreviations ............................................................................................................. xi Acknowledgements ............................................................................................................... xv Chapter 1: Introduction ....................................................................................................... 1 1.1 The role of cell migration in mammalian biology .................................................... 1 1.2 Cell motility is driven by cytoskeleton rearrangement............................................ 3 1.2.1 The cytoskeleton and its rearrangement .............................................................. 3 1.2.2 Cyoskeletal rearrangement is controlled by actin binding proteins .................... 4 1.2.3 Polarization: preparation for the move ................................................................ 7 1.2.4 Protrusions: Actin polymerization as the engine ............................................... 10 1.3 Integrins………………………………….. ............................................................... 14 1.3.1 Integrin structure................................................................................................ 15 1.3.2 Integrin activation .............................................................................................. 15 1.4 Focal adhesions .......................................................................................................... 17 1.4.1 Focal adhesion assembly ................................................................................... 18 1.4.2 Contraction: driving the cell body forward ....................................................... 21 1.4.3 Detachment: tail retraction and focal adhesion disassembly ............................. 23 v 1.5 Tyrosine kinases link integrins to signaling pathways that control the cytoskeleton................................................................................................................ 24 1.5.1 Central kinases in integrin signaling: FAK and Src .......................................... 25 Focal adhesion kinase (FAK) ............................................................................ 25 Src ...................................................................................................................... 27 1.5.2 Key downstream effectors: Cas and paxillin ..................................................... 28 p130Cas ............................................................................................................. 29 Paxillin ............................................................................................................... 30 1.5.3 The integrin signaling adapter BCAR3, a member of the novel SH2 domain- containing protein (NSP) family........................................................................ 32 1.6 PTPs control tyrosine kinase pathways in both negative and positive manners . 33 1.7 PTPα plays an important role in integrin signaling............................................... 38 1.7.1 Structure of PTPα .............................................................................................. 39 1.7.2 PTPα in integrin signaling ................................................................................. 42 1.8 Rationale and hypothesis .......................................................................................... 45 Chapter 2: Materials and Methods ................................................................................... 52 2.1 Cell lines and cell culture.......................................................................................... 52 2.2 Fibronectin stimulation ............................................................................................ 52 2.3 Antibodies and immunological detection reagents ................................................ 53 2.4 BCAR3 expression constructs .................................................................................. 53 2.5 PTPα and BCAR3 depletion, and re-expression .................................................... 54 2.6 Cell lysis, immunoprecipitation, and immunoblot analysis .................................. 55 2.7 Membrane translocation assay ................................................................................ 56 2.8 Focal adhesion enrichment assay............................................................................. 56 vi 2.9 Pulldown assays ......................................................................................................... 57 2.10 Far-Western blotting ................................................................................................ 57 2.11 Cell migration assays ................................................................................................ 58 2.12 Matrigel transwell invasion assays .......................................................................... 58 2.13 Statistical analyses..................................................................................................... 59 Chapter 3: PTPα Tyr789 Phosphorylation Regulates Integrin-induced Cell Spreading and Migration through a Cas/Crk-Rac/Cdc42 Signaling Axis .................... 60 3.1 Rationale .................................................................................................................... 60 3.2 Rac1/Cdc42 signaling is regulated by PTPα Tyr789 ............................................. 61 3.2.1 Roles of PTPα and PTPα Tyr789 in Rac1/Cdc42 activation............................. 62 3.2.2 Roles of PTPα and PTPα Tyr789 in PAK activation ........................................ 65 3.3 PTPα Tyr789 is necessary for efficient integrin -induced Cas tyrosine phosphorylation and Cas-Crk association .............................................................. 67 3.4 Cas and Src association is impaired in cells expressing mutant Y789F PTPα .... 70 3.5 PTPα-Tyr789 is required for integrin-induced focal adhesion localization of PTPα and Cas but not Src ........................................................................................ 71 3.6 Discussion ................................................................................................................... 73 Chapter 4: Cas Complexes with PTPα via its Binding Protein BCAR3 to Regulate Cell Migration .......................................................................................................... 87 4.1 Rationale .................................................................................................................... 87 4.2 BCAR3 knockdown cells have a similar phenotype to mutant Y789F PTPα- expressing cells .......................................................................................................... 88 4.3 Integrin-induced FA localization of the Cas binding protein BCAR3 requires PTPα Tyr789 ............................................................................................................. 90 vii 4.4 PhosphoTyr789 of PTPα directly interacts with the SH2 domain of BCAR3 ..... 91 4.4.1 PTPα associates with Cas and Src in an integrin- and BCAR3- dependent manner ............................................................................................................... 91 4.4.2 PTPα Tyr789 phosphorylation is required for PTPα-BCAR3 interaction ........ 93 4.4.3 BCAR3-SH2 domain is required for the PTPα-phosphoTyr789-BCAR3 interaction .......................................................................................................... 94 4.5 The binding of BCAR3 and Grb2 to PTPα phosphoTyr789 is mutually exclusive ........................................................................................................................... 95 4.6 The SH2 and GEF domains of BCAR3 are respectively required for the physical linkage of PTPα with Cas/Src ......................................................................... 95 4.7 Discussion ................................................................................................................... 97 Chapter 5: Disruption of PTPα-BCAR3-Cas Complex Formation Inhibits the Migration and Invasion of RD Rhabdomyosarcoma Cells ........................ 111 5.1 Rationale .................................................................................................................. 111 5.2 Expression of PTPα, BCAR3, and Cas in cancer cell lines ................................. 112 5.3 PTPα, BCAR3, and Cas form a complex in RD cells........................................... 114 5.4 RD cell migration and invasion are dependent on PTPα and PTPα Tyr789..... 115 5.5 BCAR3 is required for RD cell migration and invasion...................................... 117 5.5.1 BCAR3 silencing reduces RD cell invasion .................................................... 118 5.5.2 BCAR3 SH2 mutant and GEF mutant inhibit RD cell invasion ..................... 118 5.6 Discussion ................................................................................................................. 119 Chapter 6: Summary and Future Directions ................................................................. 129 6.1 The novel PTPα-BCAR3-Cas-Src signaling module in integrin signaling ........ 129 viii 6.2 PTPα-BCAR3-Cas-Src signaling in promoting cancer cell invasive motility ... 133 6.3 The PTPα phosphoTyr789 binding proteins ........................................................ 136 6.4 Conclusion................................................................................................................ 139 Bibliography ........................................................................................................................ 143 Appendix…… ...................................................................................................................... 179 ix List of Figures Figure 1.1 Schematic diagram of cell morphology and substratum interactions during cell migration ............................................................................................................ 46 Figure 1.2 Diagram of a filopodium and lamellipodium .................................................... 47 Figure 1.3 The activation cycle of Rho family small GTPases .......................................... 48 Figure 1.4 Domain structures of FAK and Src ................................................................... 49 Figure 1.5 Domain structures of the scaffolding proteins Cas and paxillin ........................ 50 Figure 1.6 Domain structure of PTPα ................................................................................. 51 Figure 3.1 Lack of PTPα Tyr789 impairs integrin-induced membrane association of Rac, Cdc42 and PAK. ................................................................................................ 77 Figure 3.2 Lack of PTPα Tyr789 impairs integrin-induced activation of Rac, Cdc42 and PAK. .................................................................................................................. 78 Figure 3.3 Integrin-induced Src activation is not dependent on PTPα Tyr789 phosphorylation. ................................................................................................ 79 Figure 3.4 Integrin-induced FAK activation is independent of PTPα Tyr789.................... 80 Figure 3.5 PTPα Tyr789 is required for integrin-induced Cas phosphorylation. ............... 81 Figure 3.6 PTPα Tyr789 is required for integrin-induced Cas phosphorylation. ............... 82 Figure 3. 7 PTPα Tyr789 is required for integrin-induced Cas-Crk association. ................ 83 Figure 3.8 Integrin-stimulated Cas-Src association requires PTPα-Tyr789. ...................... 84 Figure 3.9 PTPα Tyr789 directs PTPα to integrin-mediated focal adhesion-enriched fractions. ............................................................................................................ 85 Figure 3.10 PTPα-Tyr789 directs Cas, and Crk, but not Src, to integrin-mediated focal adhesion-enriched fractions. .............................................................................. 86 Figure 4.1 BCAR3 promotes Src activation, PTPα Tyr789 phosphorylation, and downstream signaling. ..................................................................................... 101 Figure 4.2 BCAR3 promotes PTPα and Cas FA localization. .......................................... 102 Figure 4.3 PTPα Tyr789 regulates integrin-induced BCAR3 localization to focal adhesion- enriched fractions. ........................................................................................... 103 x Figure 4.4 BCAR3 is required for the formation of a PTPα-BCAR3-Cas-Src complex and BCAR3-Cas association is independent of PTPα. ........................................... 104 Figure 4.5 The formation of PTPα-BCAR3-Cas-Src complex is regulated by integrin. .. 105 Figure 4.6 PTPα Tyr789 phosphorylation is required for the formation of a PTPα-BCAR3- Cas-Src complex. ............................................................................................. 106 Figure 4.7 BCAR3-SH2 domain is required for BCAR3 to associate with phosphoTyr789 of PTPα. ........................................................................................................... 107 Figure 4.8 PTPα phosphoTyr789 directly interacts with the BCAR3-SH2 domain. ........ 108 Figure 4.9 BCAR3 and Grb2 binding to phosphoTyr789 PTPα is mutually exclusive. ... 109 Figure 4.10 BCAR3-SH2 and -GEF domains are required for BCAR3 to associate with and link phosphoTyr789 of PTPα and Cas. ............................................................ 110 Figure 5.1 Expression of PTPα, BCAR3, and Cas in cancer cell lines ............................. 123 Figure 5.2 PTPα, BCAR3, and Cas form a complex in RD cells that is BCAR3-dependent. ......................................................................................................................... 124 Figure 5.3 RD cell migration is dependent on PTPα and PTPα Tyr789. .......................... 125 Figure 5.4 PTPα and PTPα Tyr789 are required for the invasion of RD cells. ................ 126 Figure 5.5 BCAR3 is required for RD cell invasion. ........................................................ 127 Figure 5.6 An intact BCAR3 SH2 domain or GEF domain is required for RD cell invasion. ......................................................................................................................... 128 Figure 6.1 Schematic diagram illustrating the dual roles of PTPα in integrin-mediated cell migration .......................................................................................................... 140 Figure 6.2 A schematic diagram of the mechanism by which phosphoTyr789-PTPα coordinates Cas positioning and phosphorylation in FAs. .............................. 141 Figure 6.3 Alignment of the homologous region in PTPα and EphB2. ............................ 142 Appendix Figure 1 Mutation of PTPα Tyr782 does not prevent the association of PTPα with BCAR3. ........................................................................................................... 179 xi List of Abbreviations ADF actin depolymerizing factor APC adenomatous polyposis coli BSA bovine serum albumin Cas Crk-associated substrate CCH Cas family C-terminal homology domain Cdc42 cell division cycle 42 DMEM Dulbecco’s Modified Eagle Medium DIP Dia-interacting protein Dok1 docking protein 1 DSP dual specific protein phosphatase ECD extracellular domain ECM extracellular matrix EDTA ethylenediaminetetraacetic acid EGF epidermal growth factor EGFR epidermal growth factor receptor ELMO engulfment and motility 1 ERK extracellular signal related kinase EyA eyes absent FA focal adhesion FB fibrillar adhesion FC focal complex xii FAK focal adhesion kinase FAT focal adhesion targeting domain FBS fetal bovine serum FERM protein 4.1, ezrin, radixin, moesin domain FH1 formin homology domain 1 FH2 formin homology domain 2 FN fibronectin FRAP fluorescence recovery after photobleaching FRET Förster (fluorescence) resonance energy transfer GAP GTPase activating protein GDI GDP dissociation inhibitor GEF guanine nucleotide exchange factor GPCR G protein-coupled receptor GST glutathione S transferase HRP horse radish peroxidase IGF1R insulin-like growth factor receptor 1 ILK integrin-linked kinase JMY junction-mediating and regulatory protein LCA Leukocyte common antigen LD leucine-rich domain LMPTP low molecular weight PTP LRP LCA-related protein MAPK mitogen-activated protein kinase xiii MHC myosin heavy chain MKP MAPK phosphatase MLC myosin light chain MLCK myosin light chain kinase MRCK myotonic dystrophy kinase-related Cdc42-binding kinase MT microtubule MTOC microtubule-organizing center NA nascent adhesion NMDA N-methyl-D-aspartate NP-40 Nonidet P-40 PAK p21-activated kinase PBD p21-binding domain PBM PDZ binding motif PBS phosphate-buffered saline PI3K phosphoinositide 3 kinase PIN1 peptidyl-prolyl cis/trans isomerase 1 PIP2 phosphatidylinositol-4,5-triphosphate PIP3 phosphatidylinositol-3,4,5-triphosphate PIX PAK-interacting exchange factor PKL paxillin kinase linker PTB phosphotyrosine binding PTEN phosphatase and tensin homolog PTK protein tyrosine kinase xiv PTP protein tyrosine phosphatase PTPα protein tyrosine phosphatase alpha PVDF polyvinylidene difluoride Rac Ras-related C3 botulinum toxin substrate RIAM Rap1-GTP-interacting adapter molecule RIF Rho in filopodia ROCK Rho-associated protein kinase R-PTK Receptor protein tyrosine kinase SBD Src binding domain SD substrate domain SDS sodium dodecyl sulfate SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis SFK Src family kinase SH2 Src homology 2 SH3 Src homology 3 Sos son of sevenless SYF cell mouse cells deficient in Src, Yes and Fyn THD talin head domain TRD talin rod domain WASH WASP-interacting SH3 protein WASP Wiskott-Aldrich syndrome protein WAVE WASP-family verprolin-homologous protein WT wildtype xv Acknowledgements I would like to express my deep and sincere thanks to my supervisor, Dr. Catherine Pallen, who has been patient and understanding, and is always there whenever I need guidance. Her support has made the completion of this thesis work possible. I am grateful to thank my committee members, Dr. Vincent Duronio, Dr. Michael Cox, and Dr. Michael Gold for their invaluable suggestions and careful revisions of this thesis. I warmly thank all the Pallen lab members who have made my time in the lab an enjoyable experience, and special thanks to our lab manager, Dr. Jing Wang, who has helped me in every possible way. I would lastly like to thank my family for their understanding and constant support during the down times. 1 Chapter 1: Introduction 1.1 The role of cell migration in mammalian biology Cell migration is an essential cellular process that is involved in embryogenesis, tissue development, neuron network formation, immune surveillance, and wound healing (Gilbert et al. 2010). It plays key roles from conception to death. Starting from a fertilized zygote, a human embryo grows to form a group of cells in a hollow ball-like structure termed a blastocyst, which migrates out of the fallopian tube and implants into the uterus. Once the blastocyst attaches to the inner layer of the uterine epithelium, the blastocyst outer layer cells migrate into the wall of uterus to form a placenta and stimulate endothelial cells in the uterus to initiate angiogenesis. The placenta also secretes hormones such as estrogen, progesterone, and human chorionic gonadotropin, informing the mother’s body that she is pregnant. During gastrulation, cells migrate from the surface of the embryo to the inside and form three germ layers: ectoderm, mesoderm and endoderm, which later will migrate to specific sites and different into various tissues for organ formation in the fetus. During the development of the nervous system, the extension of axons and dendrites is driven by migration of the growth cone, a key process for synapse formation (Sanes et al. 2006; O'Toole et al. 2008). Later in adulthood, cell migration is important for tissue renewal involving mucosa, skin, and bone (Ridley et al. 2003). Upon skin injury, the clotting cascade is activated to stop bleeding. Neutrophils are quickly attracted to the broken tissue to clear cell debri and bacteria by phagocytosis and stimulate the secretion of growth factors to surrounding tissue. After sensing the chemoattractant, monocytes in the blood stream transmigrate through the endothelial cell layer and migrate to the wounded site. Monocytes then mature into 2 macrophages to prepare for tissue repair by debridement. The neutrophils and macrophages usually migrate through wounded tissue to phagocytize bacteria or floating cell debris. Once the inflammation stage is approaching the end, endothelial cells are recruited for angiogenesis and fibroblasts come on site to reconstruct the extracellular matrix (ECM) by depositing collagen and fibronectin (FN) to the wound. Finally epithelial cell sheets migrate to propel the edges of the wound towards each other until the wound closes (Werner et al. 2003; Muller et al. 2012). Usually cell migration is precisely regulated to maintain the proper functioning of the body in response to stimulation such as chemical cues, ECM orientation, and physical forces. Defective cell migration can result in severe pathological conditions such as chronic inflammatory disease and cancer metastasis (Ridley et al. 2003; Eble et al. 2006; Sero et al. 2012). Metastasis is a hallmark of cancer (Hanahan et al. 2000,2011), which although formerly considered to be the last step of cancer progression, increasing evidence indicates that cell dissemination actually happens at a very early stage (Klein 2008). The genome of cancer cells is very unstable, resulting in continuous accumulation of genetic alterations even during the process of metastasis (Knudson 2001; Croce 2008). When cancer cells migrate from a primary site to a remote anatomical site, the cells break through the basement membrane to migrate into the underlying substratum, dissociate from the primary tumor mass, and transmigrate into the blood or lymph vessels, a process termed intravasation. If these disseminated cells survive in the circulation, they will travel through the bloodstream and extravasate to a new site. Usually most of the cancer cells cannot go this far. However, some successfully survive and eventually find a niche at a remote site to initiate a new colony of cancer cells, called a metastatic tumor (Talmadge et al. 2010; Peinado et al. 2011). Thus, 3 understanding the molecular mechanisms governing cell migration is the key to unlocking the enigma of novel effective cancer therapeutics. 1.2 Cell motility is driven by cytoskeleton rearrangement Based on more than 40 years of research on cell migration, a general cell migration cycle can be briefly summarized in five key steps (Fig. 1.1). Following either chemical or mechanical signals, a cell establishes polarized structures along the axis of movement, and generates membrane protrusions termed filopodia and lamellipodia at the leading edge. These protrusions are stabilized and anchored to the ECM or adjacent cells by adhesions. Stress fiber contraction then generates force to pull the cell body forward. Finally the trailing edge of the cell detaches from the substrate (Lauffenburger et al. 1996; Huttenlocher et al. 2011; Rottner et al. 2011). These steps require extensive remodeling of the cellular actin cytoskeleton and continuous adhesion alterations to achieve directional cell migration (Bugyi et al. 2010; Nagano et al. 2011; Rottner et al. 2011). 1.2.1 The cytoskeleton and its rearrangement The cytoskeleton is a network of protein polymers in the cytoplasm that not only provides morphological and mechanical support to a cell, but also controls the internal organization of a cell to facilitate many specific functions (Sobue 2011; Vignaud et al. 2012). Actin filaments, microtubules, and intermediate filaments are three main components of the cytoskeleton of prokaryotic and eukaryotic cells (Frixione 2000; Shih et al. 2006). Microtubules resist compressive forces and intermediate filaments provide strength to 4 withstand stretching. Actin filaments are intrinsically very unstable and turn over rapidly in the cell. Based on the nature of actin filaments, many studies have demonstrated that reorganization of the actin cytoskeleton is an essential process that drives cell motility (Alberts et al. 2007; Sobue 2011). Actin monomers (globular actin, G-actin) can associate with ATP and slowly hydrolyze ATP into ADP with their intrinsic ATPase activity. The interaction of G-actin with ATP or ADP changes the conformation of the actin monomers. ATP-bound G-actins tend to polymerize into actin filaments (F-actins), by contrast, ADP-G-actins prefer to dissociate from the filaments. Actin filaments are inherent polarized polymers, which have fast-growing ends termed barbed ends and slow-growing ends termed pointed ends. If we take the ATP/ADP-binding into consideration, the barbed end of an actin filament usually has more elevated ATP-binding that facilitates polymerization, whereas the ADP-bound pointed end tends to depolymerize. This process is termed the ‘treadmilling’ of an actin filament. In fact actin turnover occurs in the cell at all times, polymerizing at the barbed end from the membrane and disassembling at the pointed end in the cytoplasm (Alberts et al. 2007). 1.2.2 Cyoskeletal rearrangement is controlled by actin binding proteins The assembly of actin filaments rarely occurs spontaneously. F-actin dynamics are mediated by several groups of actin-binding proteins: nucleating proteins, sequestering proteins, crosslinking proteins, severing proteins, and capping proteins. In brief, nucleating proteins (formin, spire, Arp2/3 complex) promote actin polymerization; sequestering proteins (profilin) can specifically bind to actin monomers (G-actins); crosslinking proteins (fimbrin, filamin, and fascin) hold actin filaments in bundles; severing proteins (cofilin) break and 5 promote depolymerization of actin filaments; and capping proteins can bind to the barbed ends of actin filaments to stop elongation. Formin Formins are a family of formin homology 2 (FH2) domain-containing proteins, which initiate the elongation of linear actin filaments. Formin homodimers use their FH2 domains to associate with actin filaments and promote actin polymerization at the barbed end. During the elongation process, formins can remain attached to the polymerizing barbed end of actin filaments and antagonize capping proteins, functioning as an assembly motor to promote continuous nucleation (Faix et al. 2006; Pollard 2007). Usually formins also contain a formin homology 1 (FH1) domain for recruitment of profilin-ATP-G-actin to the barbed end of actin filament (Goley et al. 2006). Profilin is a sequestering actin-binding protein, which prefers ATP-G-actin binding and can also recharge actin by replacing ADP with ATP. It can increase the rate of formin-catalyzed nucleation by delivering the ATP-G-actin monomers to formins (Kovar 2006; Romero et al. 2007). Formins can be activated by Rho family small GTPase RhoA. The C-terminal FH2 domain of formin mDia1 interacts with its N-terminal region leading to an intra-molecular inhibitory conformation, and RhoA binding to the N terminus can relieve this auto-inhibition and activate mDia1 (Li F et al. 2005; Otomo et al. 2005; Nezami et al. 2006; Goode et al. 2007). Formins are inhibited in vitro by Dia- interacting protein/WASP-interacting SH3 protein (DIP/WASH), tropomyosin, and Bud14p (Eisenmann et al. 2007; Chesarone et al. 2009; Skau et al. 2009). 6 The Arp2/3 complex Upon binding to the side of an existing actin filament, the Arp2/3 complex initiates actin filament branching by functioning as a new nucleation core. The new actin filament polymerizes with actin monomers from the Arp2/3 complex, resulting in a new actin filament branching at a 70° angle. In this way, Arp2/3 complexes generate dendritic actin filament arrays at the leading edge of a migrating cell, pushing the membrane forward to form a brush-like protrusion termed a lamellipodium (Pollard et al. 2000; Pollard et al. 2003). The Arp2/3 (actin-related protein) complex consists of 7 subunits: an Arp2 and Arp3 core stabilized by a five-subunit complex (ARPC1-5) (Pollard 2007). The Arp2/3 complex can be activated by binding to nucleation promoting factors (NPFs), which have two well-studied members: WASP (Wiskott-Aldrich syndrome protein) and Scar/WAVE (suppressor of cyclic AMP repressor/ WASP-family verprolin-homologous protein). WASP and WAVE can also be activated by binding of Rho family small GTPases (Rac and Cdc42), phospholipids (PI(4,5)P2), and SH3 domain-containing proteins (Nck and Grb2) (Goley et al. 2006). Spire The Spire family of actin-binding proteins share a common property; they all contain WASP homology 2 (WH2) domains, including Spire, cordon-bleu (Cobl), adenomatous polyposis coli (APC), and junction-mediating and regulatory protein (JMY). They use their WH2 domains to gather an actin nucleus to initiate linear actin polymerization (Qualmann et al. 2009). Furthermore, Spire is also able to sever existing actin filaments and mediate new polymerization at the barbed end (Bosch et al. 2007). 7 Cofilin The major role of the actin depolymerizing factor (ADF) cofilin in cytoskeleton reorganization is to sever and depolymerize actin filaments, and to regulate intracellular actin pools (Tsai et al. 2012). Cofilin binds ADP-G-actin with higher affinity than ATP-G-actin and hydrolyzes ATP in actin filaments, exactly opposite to the action of profilin (Carlier et al. 1997). Active Rho GTPases induce Rho-associated protein kinase (ROCK)-mediated phosphorylation and activation of LIM kinase LIMK (Kobayashi et al. 2007). LIMK in turn phosphorylates the cofilin Ser3 residue and inhibits cofilin-mediated actin filament depolymerization (Bernard 2007). Cofilin phosphoSer3 can be dephosphorylated by slingshot phosphatase and protein phosphatase 1 (PP1) to promote F-actin depolymerization (Takuma et al. 1996). Cofilin can also be activated by the phospholipid PIP2, a product of the phosphatase PTEN (Bernstein et al. 2010). In summary, the above actin-binding proteins function as machinery that can remodel the actin cytoskeleton in response to signaling from upstream regulators, such as phospholipids and Rho family small GTPases. 1.2.3 Polarization: preparation for the move The concept of cell polarization is based on the observation that some key structures localize asymmetrically in different compartments in migrating cells. Morphologically, a migrating cell can be divided into three regions along the axis of movement: the leading edge contains thinner actin filaments and smaller adhesions, whereas the central and rear regions have thicker actin filament bundles and larger stable adhesions (Ridley et al. 2003) (Fig. 1.2). 8 Molecularly, active Cdc42 can be detected at the leading edges of migrating cells at the edge of a wound in a cultured cell monolayer (Nalbant et al. 2004). Another report showed that not only active Cdc42 but also its guanine nucleotide exchange factor (GEF), βPIX, accumulates at the leading edge of the astrocytes during wound-induced migration (Osmani et al. 2010). Active Rac is also enriched at the leading edge as shown by FRET experiments (Kraynov et al. 2000; Donaldson et al. 2009), as are key actin cytoskeleton controllers such as the Arp2/3 complex and formin (Lai et al. 2008; Campellone et al. 2010; Suraneni et al. 2012). Using GTPase biosensors, it was shown that RhoA is activated at the cell leading edge, indicating that RhoA is also involved in the initial events of protrusion, whereas the activation of Cdc42 and Rac1 is delayed and localized behind the front of the leading edge, (Machacek et al. 2009). Rho family small GTPases have been linked to the regulation of actin cytoskeleton reorganization in many aspects, and three of these family members, Rac, Cdc42, and RhoA, are the best studied (Ridley 2006; Mattila et al. 2008). GTP binding induces the activation of Rho-GTPases and hydrolysis of GTP to GDP by their intrinsic GTPase activity switches them off. The activation of Rho-GTPases is positively regulated in vivo by specific guanine nucleotide exchange factors (GEFs), and negatively regulated by GTPase-activating proteins (GAPs) and GDP dissociation inhibitors (GDIs) (Rittinger et al. 1997; Scheffzek et al. 2000; Pertz et al. 2006) (Fig. 1.3). In brief, the GEFs promote GDP to GTP exchange to recharge the RhoGTPases with GTP (Rossman et al. 2005; Cote et al. 2007), whereas the GAPs enhance the intrinsic GTPase activity of Rho-GTPases to accelerate GTP hydrolysis (Tcherkezian et al. 2007). The GDIs preferably bind to the C- terminal isoprenyl groups of GDP-bound Rho-GTPases and sequester them in the cytoplasm. In this way, GDIs prevent the translocation of Rho-GTPases to the plasma membrane where 9 the Rho-GTPases interact with effector proteins (DerMardirossian et al. 2005). Recent reports also show that the phosphorylation of Rho GTPases plays a negative role in their activation by promoting their dissociation from the membrane and association with GDIs (Boulter et al. 2012). Interestingly, PI3K, the kinase that synthesizes the phospholipid PI(3,4,5)P3, has also been found to be concentrated at the leading edge of cells, while the phosphatase PTEN, which is responsible for dephosphorylation of PI(3,4,5)P3 to PI(4,5)P2, is restricted to the side and rear of the cell, suggesting that these phospholipids play multiple roles in spatially regulating cytoskeleton reorganization (Devreotes et al. 2003; Merlot et al. 2003). Another key polarized structure in the cell is the microtubule-organizing center (MTOC), which is usually reoriented in front of the nucleus and after the leading edge in slow migrating cells (Ridley et al. 2003). This MTOC orientation has been used as a marker for cell polarization, and can be clearly recognized to distinguish the leading edge towards the direction and the axis of movement (Friedl et al. 2010; Vicente-Manzanares et al. 2011). The polarization of cells has been proposed to be maintained through microtubule-mediated vesicular trafficking (Ridley et al. 2003; Ridley 2011). Cdc42 regulates the orientation of MTOC through adenomatous polyposis coli (APC)-mediated capture of microtubule growing plus ends at the leading edge, facilitating microtubule-mediated vesicle trafficking to generate polarity (Ridley et al. 2003; Sugioka et al. 2012). Currently polarization is believed to be a prerequisite for directional cell migration, which allows the local activation of different molecules, such as phospholipids and Rho family small GTPases in the front or back of a migrating cell. These molecules in turn activate their effectors to drive the actin machinery for cytoskeletal remodeling (Ridley et al. 2003; Ridley 10 2011). Cdc42 is activated at the leading edge of a migrating cell to promote filopodium and lamellipodium formation, since either the depletion of Cdc42 or the global activation of Cdc42 impairs directional cell migration (Etienne-Manneville et al. 2002; Itoh et al. 2002; Srinivasan et al. 2003). Positive feedback loops at the leading edge also contribute to the generation of intracellular asymmetry, such as Rac and Cdc42 activation at the leading edge and Rho activation at the trailing edge of the cell (Ridley 2011). The phospholipid PI(3,4,5)P3, a PI3K product at the leading edge, has been shown to activate RacGEFs, such as p-Rex1, which induce the activation of Rac (Welch et al. 2003; Wertheimer et al. 2012). In turn, Rac activation promotes PI3K membrane translocation and activation, further enhancing Rac activation at the leading edge (Yang HW et al. 2012). The engagement of integrins with their ECM ligand induces focal adhesion (FA) formation and FA formation promotes Rac activation at the leading edge, which in turn promotes integrin clustering and the maturation of FAs (Harburger et al. 2009). Activated Rac also promotes the formation of more cell protrusions and FAs, which further induce Rac activation, keeping Rac constitutively active at the leading edge of the cell (Ren et al. 2000; Ridley 2011). 1.2.4 Protrusions: Actin polymerization as the engine The first step in cell migration is the extension of protrusions at the leading edge. The two main protrusive structures are known as lamellipodia and filopodia. Lamellipodia are sheet- like protrusions, 0.1~0.2μm thick and supported by underlying branched actin networks (Mattila et al. 2008) (Fig. 1.2A). Lamellipodia localize at the front of the leading edge, supported by structures termed lamella, which provide substratum adhesion and facilitate 11 myosin II-mediated contractility. The C-terminal acidic domains of WASP/WAVE proteins interact with and activate the Arp2/3 complex, promoting Arp2/3 complex binding to the side of an existing actin filament to initiate the nucleation of a new branch. The Arp2/3 complex will hold the pointed end of the new actin filament on the old filament, and formin or Spire may promote the elongation of the new filament with ATP-G-actin monomers delivered by profilin. Meanwhile, capping proteins stop the elongation process to prevent forming a long linear actin filament (Pollard et al. 2003; Mejillano et al. 2004). The phosphorylation of cortactin, which can be catalyzed by active Src or FAK (Kowalski et al. 2005; Tomar et al. 2012), promotes interaction of cortactin with the Arp2/3 complex to stabilize novel Y-shape branches (Weaver et al. 2001; Ammer et al. 2008). JMY also stimulates the Arp2/3 complex to generate more branches to form a branched actin filament network (Firat-Karalar et al. 2011; Rottner et al. 2011). Filamin A and α-actinin can then crosslink the branches to further stabilize the actin array (Esue et al. 2009; Kasza et al. 2009). This array of actin filaments pushes the membrane forward, resulting in the formation of a lamellipodium. The Rho family GTPases Rac1 and Cdc42 are central regulators of lamellipodia formation (Rottner et al. 2011). Using a photoactivatable Rac1 and live cell microscopy, it was shown that the local activation of Rac1 directly induced the generation of a localized lamellipodium in MEFs and HeLa cells (Wu YI et al. 2009). Activated Rac-GTP induces the activation of the WASP family Verprolin-homologous protein (WAVE), which utilizes its WCA domain to activate the Arp2/3 complex (Miki et al. 2000; Takenawa et al. 2007; Chen Z et al. 2010). A Rac-GTP effector, PAK, promotes the delivery of WAVE to the membrane at the lamellipodia through WAVE-PAK-kinesin complex formation (Takahashi et al. 2009). PAK also phosphorylates and activates LIM kinase, which inhibits cofilin-mediated severing and 12 depolymerization of actin filaments (Bernard 2007; Manetti 2012). On the other hand, Cdc42-GTP activates WASP to induce Arp2/3 complex activation and promote formin activation to initiate linear polymerization at the barbed ends (Stradal et al. 2006). Filopodia, also called microspikes, are finger-like membrane protrusive structures containing parallel bundles of actin filaments, which usually have diameters between 0.1- 0.3μm (Mattila et al. 2008) (Fig. 1.2B). Filopodia have been proposed to be the exploratory protrusions that sense the surroundings during cell migration and transduce the extracellular signal into the cell (Yang C et al. 2011). The formation of filopodia starts from either uncapped existing actin filaments or nucleation directly from monomers (Svitkina et al. 2003; Yang C et al. 2007). Formin Dia2 (diaphanous-related formin 2) and Ena/VASP (enabled/vasodilator-stimulated phosphoprotein) proteins attach to the barbed end of actin filaments and initiate elongation (Steffen et al. 2006; Gupton et al. 2007). The novel actin filaments are converged to bundles by Myosin-X binding (Tokuo et al. 2007). Once the novel filopodial actin bundles start to push the membrane, insulin receptor substrate p53 (IRSp53), an I-BAR domain containing protein, deforms the membrane at the site to facilitate the formation of a membrane protrusion (Suetsugu et al. 2006). Myosin-X-mediated protein transportation delivers fascin and integrins to the novel filopodium (Zhang H et al. 2004; Watanabe TM et al. 2010). Fascin crosslinks actin filaments into stiff bundles, while the formin Dia2 continuously promotes the actin polymerization at the tip of novel filopodium (Machesky et al. 2010; Yamashiro 2012). Microinjection of Cdc42 has been reported to induce filopodia in Swiss 3T3 fibroblasts (Paterson et al. 1990; Ridley et al. 1992). After twenty years of research, we know that Cdc42 is enriched at the leading edge (Nalbant et al. 2004; Osmani et al. 2010), and can 13 activate formin Dia2 to initiate the formation of filopodia (Ahmad et al. 2010). Cdc42 also forms a complex with IRSp53 and Ena/VASP family member MENA and induces their local activation at the tip of the filopodia to protect elongating actin filaments from capping (Zhao et al. 2011). Surprisingly, depletion of Cdc42 does not affect the formation of filopodia and even has no significant influence on the speed of directed migration in fibroblastoid cells. Overexpression of a dominant-negative mutant of Cdc42 inhibits protrusion formation towards the wound and directed cell migration, suggesting that Cdc42 is not the only GTPase that can induce filopodia formation (Czuchra et al. 2005). Another study showed that the overexpression of Rho in filopodia (RIF) also enhances the formin Dia-mediated formation of filopodia in NIH3T3 cells and N1E115 mouse neuroblastoma cells, indicating an overlapping role of Cdc42 and RIF in regulating filopodia formation (Pellegrin et al. 2005; Goh et al. 2011). Once membrane protrusions have formed, they have two fates: 1) if their surface receptors cannot attach to the substratum, the protrusions retract to merge with the membrane, forming a structure termed a membrane ruffle; 2) if their surface receptors engage with ECM ligands, membrane protrusions will be stabilized by adhesions and evolve into more stable structures, such as lamellae. The extracellular matrix (ECM) is composed of numerous insoluble fibrous proteins and polysaccharides secreted locally by epithelial and stromal cells. It provides physical support to neighboring cells within a tissue, segregates tissues from one another, and serves as a reservoir for many growth factors that regulate intercellular communications (Ozbek et al. 2010; Hynes et al. 2012). While attaching to the ECM, cells use their surface receptors in the protrusions to sense the environment such as the presence of soluble cues and the adhesive states of the ECM surface (Hynes 2002; Humphries et al. 2006). There are two 14 specific types of cell-to-ECM adhesions: 1) adhesomes which link the actin cytoskeleton to the ECM (Zaidel-Bar et al. 2010); 2) hemidesmosomes, connecting the cellular intermediate filaments to the ECM (Litjens et al. 2006). Both types of cell-to-ECM adhesions are mediated by a major family of cell surface receptors known as integrins. 1.3 Integrins Integrins are cell surface receptor proteins that play essential roles in mediating cell-to- ECM interactions. All integrins are non-covalently linked dimers containing one α subunit and one β subunit (Humphries et al. 2006; Luo et al. 2007). So far 18 α subunits, 8 β subunits, and a total of 24 different integrins have been identified in vertebrates (Hynes 2002; Zaidel-Bar et al. 2010; Hu P et al. 2013). Most integrins can interact with a wide range of ECM components, while the same ligand may also be recognized by multiple integrins (Humphries et al. 2006; Kim et al. 2011). The knockout of different integrins in mice has been shown to induce distinct phenotypical defects, suggesting that each integrin may have a specific function and a great number of integrins can carry out precise, cell type-specific regulation of a wide range of cell behaviors in response to different extracellular conditions (Cheresh et al. 2002; Raymond et al. 2005; Foubert et al. 2011). Integrins are ubiquitously expressed, and cells from different tissues have their own specific integrin expression profiles related to their developmental stage, specific functions, and environments (Barczyk et al. 2010). Some integrins, such as α1β1 and α2β1, are associated with inhibition of tumorigenesis (Desgrosellier et al. 2010; Rathinam et al. 2010), whereas others, such as αvβ3 and α9β1, are implicated in aggressive and metastatic cancers (Spangenberg et al. 2006; Gupta et al. 2012). Switching cell surface integrin molecules is a common trick used by 15 cancer cells to invade and metastasize during malignant progression (Hanahan et al. 2000,2011). 1.3.1 Integrin structure Integrin α and β subunits are type I transmembrane proteins, characterized by large N- terminal extracellular domains (ECD) and short cytoplasmic domains at the C-terminus, with the exception that the integrin β4 subunit also has a large cytoplasmic domain (Suzuki et al. 1990; Hu P et al. 2013). Both the α and β subunits of the integrins are involved in ligand binding specificity (Humphries et al. 2006), and they are usually associated with each other in a bent confirmation on the cell surface, representing a low affinity “resting” state (Xiong et al. 2001; Zhu et al. 2008; Xie et al. 2010). 1.3.2 Integrin activation Upon stimulation by ligand binding or intracellular signals, the cytoplasmic tails of the integrin α and β subunits separate from each other, disrupting the association between their transmembrane domains. This causes the ECD of integrins to switch to the extended or upright conformation, which displays a high affinity towards the ligands (Luo et al. 2007; Campbell et al. 2011). It has been shown that crosslinking the integrin α2bβ3 transmembrane domains by a disulfide bond results in abolishment of integrin activation and defective cell spreading (Zhu et al. 2007). The initial cytoplasmic separation of integrin α and β subunits can be induced by the binding of an important integrin-activating protein known as talin to the β subunit of 16 integrins. Talin is a 270 kD cytoplasmic actin-binding protein with an 50 kD N-terminal talin head domain (THD) followed by a 220 kD talin rod domain (TRD). The head domain of talin contains a phospholipid-binding domain and a phosphotyrosine-binding (PTB) domain. Through this PTB domain, talin interacts with an integrin β subunit membrane-proximal NPxY motif, an essential step for integrin activation (Calderwood et al. 2002). It has been shown that the association of the THD with the integrin β3 subunit cytoplasmic tail is sufficient to change integrin α2β3 to its extended conformation and induce integrin activation in phospholipid nanodiscs (Ye et al. 2010). The rod domain of talin consists of multiple binding regions for proteins such as actin and vinculin, providing the talin-mediated linkage of integrin to the cytoskeleton (Zhang X et al. 2008; Wang P et al. 2011). Talin also has an intramolecular autoinhibitory domain in its TRD, which covers its own THD and prevents it from interacting with integrins, thus inhibiting integrin activation and clustering (Goksoy et al. 2008; Goult et al. 2009). The interaction of PI(4,5)P2 with the THD can recruit talin to the membrane and break this autoinhibitory conformation, resulting in talin-mediated activation and clustering of integrins on the cell surface (Saltel et al. 2009). Once a cell senses the cue for migration via its surface receptors such as GPCRs during chemotaxis, it induces the activation of the small GTPase Rap1, which recruits talin to the integrins (Watanabe N et al. 2008; Lee HS et al. 2009). Active Rap1-GTP forms a complex with talin through its effector Rap1-GTP-interacting adapter molecule (RIAM). The membrane-targeting domain of Rap1 directs the membrane translocation of the Rap1-RIAM- talin complex, delivering talin proximal to the integrins. Talin then binds to the integrin β subunit cytoplasmic NPxY motif to activate the integrin (Watanabe N et al. 2008; Lee HS et al. 2009). Activated integrins engage their ligands with high affinity and form clusters to 17 initiate adhesion formation. This process can be regulated by many talin binding partners, such as kindlin, docking protein 1 (Dok1), and filamin. Kindlin is proposed to have a synergistic effect with the THD to promote integrin activation since co-expression of kindlin- 1/2 together with the THD showed a higher activation of integrin α2bβ3 than that achieved by expression of the THD alone (Ma et al. 2008; Harburger et al. 2009). A recent report showed that talin-mediated neutrophil integrin activation is defective in the absence of kindlin-3, which confirmed that kindlin is required for the full activation of integrin with optimal affinity (Lefort et al. 2012). Dok1 and filamin are negative regulators of integrin affinity which compete with talin to interact with the integrin β subunit cytoplasmic tail to attenuate integrin activation (Kiema et al. 2006; Wegener et al. 2007). 1.4 Focal adhesions Integrin signaling is bi-directional. On the one hand, the “inside-out” signaling promotes integrin activation and physically links the cytoskeleton to the ECM (Miranti et al. 2002 2013). On the other hand, ligand-engaged integrins also function as cell surface sensors to transduce extracellular environmental signals to intracellular pathways, which is termed “outside-in” signaling (Hynes 2002; Huttenlocher et al. 2011; Shen B et al. 2012). Since integrins lack enzymatic activity, “outside-in” signaling has to rely on the recruitment of other proteins and the formation of signaling structures called focal adhesions (FAs) to facilitate subsequent signal transduction to downstream pathways, such as the mitogen- activated protein kinase (MAPK) pathway, the phosphoinositide-3 kinase (PI3K)-AKT pathway, and the Rho GTPase-mediated actin cytoskeleton pathway to control cell proliferation, survival, and motility (Ginsberg et al. 2005; Harburger et al. 2009). 18 FAs refer to large dynamic protein complexes (>180 components) on the cytoplasmic side of the cell membrane. These contain numerous different proteins, such as kinases, phosphatases, small GTPases, cytoskeletal proteins, proteases and scaffolding proteins (Burridge et al. 1988; Zaidel-Bar 2009; Hu P et al. 2013). The engagement and clustering of integrins are essential for FA formation, and the cytoplasmic tails of integrins are known to interact with many signaling and structural proteins, such as paxillin and talin. Paxillin and talin can further recruit other proteins to the FAs directly or indirectly. These recruited proteins include focal adhesion kinase (FAK), Src, integrin-linked kinase (ILK), vinculin, and Crk-associated substrate (Cas), which help to transduce the integrin-initiated chemical and mechanical signals into a cell as well as physically link integrins to the cytoskeleton (Zaidel-Bar et al. 2007; Harburger et al. 2009). Each FA complex is in a state of constant flux, since proteins are recruited to the complex and dissociate from the complex continually. The exact mechanism of this FA formation is largely unclear, but protein-protein interaction is definitely an essential process in this event. At the last count, more than 742 direct interactions have been reported between FA components. These interactions can be regulated by local interaction affinity, conformation change, phosphorylation states, and the GTP loading of small GTPases, making up a very complex signaling network (Harburger et al. 2009; Zaidel-Bar et al. 2010). 1.4.1 Focal adhesion assembly If we look at FA formation in more detail, it can be further divided into four stages based on the predominant structures formed in each stage and their size: 1) nascent adhesion (NA), 2) focal complex (FC), 3) focal adhesion (FA), and 4) fibrillar adhesion (FB) (Vicente- 19 Manzanares et al. 2011). Upon ligand binding or chemotaxis, integrins at the front of the lamellipodia or at the tip of the filopodia switch to their high affinity confirmation, bind to their specific ligands and form clusters on the cell undersurface (Miyamoto et al. 1995; Ballestrem et al. 2001; Roca-Cusachs et al. 2009). Integrin clustering initiates the recruitment of a wide range of proteins to form small dot-like shaped NAs at the leading edge of a migrating cell. The components of NAs include most of the adhesome proteins, including integrin, talin, vinculin, focal adhesion kinase (FAK), and Crk-associated substrate (Cas) (Choi D et al. 2008; Vicente-Manzanares et al. 2011). Interestingly, at the NA stage, FAK, Src, paxillin, and Cas are already highly phosphorylated, indicating that the outside-in integrin signaling begins very early in adhesion formation (Nayal et al. 2006; Vicente- Manzanares et al. 2011). NAs are very dynamic and usually assemble and disassemble very quickly with a turnover rate of less than 1 minute (Zaidel-Bar et al. 2003; Nayal et al. 2006; Oser et al. 2009). Actin polymerization promotes NA initiation (Alexandrova et al. 2008; Vicente-Manzanares et al. 2011), and the formation of NAs stabilizes the local membrane protrusions and prevents them from retracting like membrane ruffles, which in turn facilitates the formation of new protrusions (Brown CM et al. 2006; Hu K et al. 2007; Alexandrova et al. 2008). As the leading edge moves forward, NAs turn into intermediate structures termed focal complexes (FC), which are larger than NAs and localize close to the lamellum. Although most FCs go through disassembly and turnover, some FCs eventually mature into large oval- shaped structures known as focal adhesions (FA) (Geiger et al. 2011; Vicente-Manzanares et al. 2011). FAs are located in the lamella and associate with actin bundles that are cross- linked by α-actinin and non-muscle myosin II (NMII) (Choi CK et al. 2008). Since FAs are 20 major anchorage sites of stress fibers for intercellular force generation, the further elongation and cross-linking of actin filaments into large bundles leads to the formation of even larger adhesions called fibrillar adhesions (FB). FBs are very stable with a turnover rate of several hours, and are located in the center, side and rear of the migrating cells (Zaidel-Bar et al. 2004; Geiger et al. 2011). Mature FAs and FBs are characterized by largely reduced phosphotyrosine and recruitment of tensin and zyxin to the adhesion sites (Zamir et al. 2001; Zaidel-Bar et al. 2004; Vicente-Manzanares et al. 2009). In rat embryo fibroblasts, integrin α5β1 becomes the dominant integrin in FBs, instead of integrin αvβ3 in early-stage FAs (Zamir et al. 1999). During FA maturation, myosin II-generated contraction is a key factor that determines the fate of FAs. Those FAs that cannot provide enough anchorage force from integrin-bound substratum will quickly go through disassociation and induce retraction of protrusions (Vicente-Manzanares et al. 2009; Gardel et al. 2010). The multi-layer ultrastructure of FAs has been revealed on the nanometer scale by interferometric photoactivated localization microscopy and cryo-electron tomography (Kanchanawong et al. 2010; Patla et al. 2010). The distribution of FA components into three spatial and functional layers has been proposed: a membrane-proximal integrin signaling layer (FAK and paxillin), a middle force transduction layer (talin and vinculin), and an uppermost actin regulatory layer (VASP, zyxin, α-actinin) (Kanchanawong et al. 2010). The molecular composition of FAs is highly dynamic and diverse. An immunofluorescent study in the fish keratinocyte showed that NAs at the front of the lamellipodium only contain integrin β1 and talin and the more mature FAs behind the lamellipodium front possess vinculin. Those located at the rear region of the lamellipodium further recruit α-actinin and FAK (Lee J et al. 1997). Expression of GFP fusion proteins in CHO cells showed that 21 paxillin is recruited to NAs earlier than α-actinin and integrin α5 (Laukaitis et al. 2001). Similar experiments in porcine aortic endothelial (PAE) cells revealed that only β3-integrin, talin, paxillin and low levels of vinculin and FAK were present in NAs, whereas zyxin and tensin co-localized with more mature FAs (Zaidel-Bar et al. 2003). The above studies suggest that FA assembly is a sequential process: Once the membrane at the leading edges of lamellipodia is in contact with the substratum, integrin activation and clustering initiate NA formation by recruiting actin binding molecules (talin and vinculin) and signaling molecules (FAK and Src) to adhesion sites, followed by the recruitment of other signaling proteins such as paxillin and Cas. The latter in turn interact with their downstream effectors to build up mature focal adhesions (Zaidel-Bar et al. 2004; Geiger et al. 2011). Since the composition of adhesion structures at each stage is very similar, in this thesis I will use the general term focal adhesion (FA) to represent integrin-induced cell-to-ECM adhesions, in accordance with most published articles. 1.4.2 Contraction: driving the cell body forward Once the cell is anchored to the ECM through FA formation, myosin-generated contraction pulls the cell body and the nucleus forward over the FAs. This process is coordinated by the formation of force-generating stress fibers and anchorage-providing mature FAs. Stress fibers consist of actin filaments and non-muscle myosin II. Actin filaments provide scaffolding support for the latter like railway tracks and non-muscle myosin II generates moving force like a locomotive (Vicente-Manzanares et al. 2009). 22 Myosin II complexes are composed of two 230 kD heavy chains (MHCs) that form the back bone of the complex, two regulatory light chains (R-MLCs) that regulate complex activation, and two essential light chains (E-MLCs) that stabilize the structure of the complex. Each MHC has 3 major domains: a globular head domain, central neck domain, and a long rod domain at the C-terminus. The head domain interacts with actin filaments and has ATP hydrolyzing activity. Using the energy from ATP, the head domain walks on the actin filament to generate contractile force. The flexible neck domain, which is also called the lever arm, interacts with MLCs and links the head domain with the rod domain for mechanical force transduction. The rod domain mediates the interactions between the MHCs, which include the association of two MHC monomers and the tail-to-tail interactions between myosin II complexes. The latter interactions link actin filament-bound myosin II complexes to form stress fibers for contractile force generation (Takashima 2009; Sweeney et al. 2010). Myosin II complexes can be activated by reversible phosphorylation of the R-MLCs at Thr18, Ser19, or both to promote stress fiber formation and ATP hydrolysis (Ikebe et al. 1986; Hirata et al. 2009). Many kinases have been demonstrated to phosphorylate the R- MLCs, such as myosin light chain kinase (MLCK), Rho-associated kinase (ROCK), and myotonic dystrophy kinase-related Cdc42-binding kinase (MRCK), which are further regulated by other upstream signaling molecules (Matsumura 2005; Tan et al. 2008; Vicente- Manzanares et al. 2009). RhoA activates Rho-associated kinase (ROCK), which can directly phosphorylate and activate MLCK. Activated MLCK phosphorylates the R-MLCs at Thr18 and Ser19 to activate the myosin II complex. Meanwhile, active ROCK also inhibits the dephosphorylation of R-MLCs by suppressing myosin phosphatase PP1 activity. Both events 23 induce intracellular tension from contractility of the stress fibers (Ikebe et al. 1986; Hirata et al. 2009). Since the ends of actin filaments at the leading edge are anchored to the ECM through integrin-mediated FAs, the contraction of these stress fibers pulls the cell body towards the leading edge. Thus, the lamella area that is right behind the lamellipodia has been shown to have the strongest pulling force in the cell (Vicente-Manzanares et al. 2009). 1.4.3 Detachment: tail retraction and focal adhesion disassembly While the cell body moves forward, the rear of the cell has to detach from the substratum to complete directional migration. The importance of microtubules (MTs) in FA turnover has been established in studies using the small molecule drug nocodazole, which interferes with MT polymerization, causing MT cytoskeleton breakdown (Nagano et al. 2011). The addition of nocodazole into the culture media disrupts the MT cytoskeleton and inhibits FA disassembly, resulting in stabilized cell-to-ECM adhesions, enhanced cell spreading and reduced cell migration. The withdrawal of nocodazole from the media induces the reconstruction of MT cytoskeletal networks and rescues cell migration defects by the initiation of FA disassembly (Ezratty et al. 2005; Ezratty et al. 2009). Nocodazole treatment also induces increased tyrosine phosphorylation of FA proteins, which is quickly reduced upon the removal of nocodazole from the media, suggesting that MT targeting suppresses the signal transduction within FAs. Several phosphatases, including PTP-PEST, PTP1B, and SHP2, have been proposed to localize to FAs via MTs to dephosphorylate FAK, Src, and actinin (Yu et al. 1998; Zhang Z et al. 2006; de Oliveira et al. 2009; Zheng Y et al. 2011). Local enrichment of integrin-binding proteins such as Dok1 and filamin, displaces talin from its interaction with the β subunit tails of integrins to deactivate integrins at the trailing edge 24 (Millon-Fremillon et al. 2008; Harburger et al. 2009). After integrins are released from their ECM ligands, a MT-binding GTPase known as dynamin initiates the endocytosis of integrins and other surface receptors for recycling (White et al. 2007; Ezratty et al. 2009). Several irreversible mechanisms have also been proposed to release the cell from the ECM at the trailing edge. Myosin II-mediated contraction can induce the activation of stretch-activated calcium channels. Ca 2+ influx through the activated channel promotes Ca 2+ - dependent protease calpain to cleave the integrins, talin, paxillin, and FAK (Glading et al. 2002; Franco et al. 2004; Perrin et al. 2006; Flevaris et al. 2007). Cleaved FA components are subjected to ubiquitin-mediated protein degradation (Schmidt et al. 2005). Sometimes the contractile force generated by myosin II can be strong enough to disrupt the linkage between the integrin and the cytoskeleton, leaving a track of integrins on the substratum behind the migrating cell (Ridley et al. 2003). 1.5 Tyrosine kinases link integrins to signaling pathways that control the cytoskeleton Upon integrin activation and clustering, FA components assemble around integrins at the cytoplasmic tail of the integrins, which not only physically connect the integrins to the cytoskeleton, but also initiate the “outside-in” integrin signaling through multiple signaling proteins to control cell behaviors. The recruitment of two important kinases to FAs, focal adhesion kinase (FAK) and Src, have been identified as the earliest outside-in signaling events (Sieg et al. 1999; Ginsberg et al. 2005; Mitra et al. 2005; Sheetz et al. 2006). 25 1.5.1 Central kinases in integrin signaling: FAK and Src Focal adhesion kinase (FAK) FAK is a 125 kD non-receptor protein tyrosine kinase (PTK) that was identified at FAs (Hanks et al. 1992; Schaller et al. 1992). It contains an N-terminal FERM (protein 4.1, ezrin, radixin, and moesin homology) domain, a central kinase domain, a proline-rich region, and a FA targeting domain (FAT) at the C-terminus (Giancotti et al. 1999; Cohen et al. 2005) (Fig. 1.4). The FERM domain mediates the membrane association of FAK and participates in an autoinhibitory conformation of FAK. An interaction between the FERM domain and the kinase domain covers the catalytic site of FAK, preventing the interaction of FAK with its substrates (Lietha et al. 2007). This inhibitory interaction can be disrupted by the binding of phospholipid PI(4,5)P2 to the FAK FERM domain at the membrane (Zhang X et al. 2008). The proline-rich region can be a docking site for the binding of SH3 domain-containing proteins such as Cas (Polte et al. 1997). The FAT domain is responsible for FAK FA localization and interacts with both talin and paxillin (Cooley et al. 2000; Schaller 2010). FAK-null or dominant-negative FAK mutant (FRNK)-expressing fibroblasts are round- shaped with increased numbers of small FAs, and display impaired directional cell migration (Ilic et al. 1995; Owen et al. 1999; Sieg et al. 1999). Later studies showed that the disassembly of FAs and tail detachment are defective at the trailing edge in FAK -/- and siRNA-mediated FAK knockdown fibroblasts (Ren et al. 2000). The formation of nascent FAs is normal at the cell periphery in FAK-null fibroblasts, suggesting that novel FA formation is independent of FAK and its kinase activity. However, these new FAs fail to mature into larger stress fiber-linked adhesions and arrest at the cell periphery, unable to move on to complete the FA assembly and disassembly cycle. Re-introduction of functional 26 FAK into FAK -/- cells rescues the defective FA maturation and turnover dynamics, indicating that FAK plays an essential role in the tail retraction of a migrating cell through FA turnover (Sieg et al. 1999; Izaguirre et al. 2001; Iwanicki et al. 2008; Lim Y et al. 2008). FAK has also been proposed to positively regulate cell protrusions at the leading edge, since FAK- depleted cells fail to form lamellipodia at the edge of a wound (Tilghman et al. 2005). The recruitment of FAK to clustered integrins through its FAT domain is one of the earliest signaling events during FA assembly (Miyamoto et al. 1995; Kirchner et al. 2003). FAK at this stage is still in its autoinhibitory conformation, and interacts with Arp3 to promote Arp2/3 complex-mediated lamellipodia formation at the leading edge (Serrels et al. 2007). The force applied to FAs induces FAK auto-phosphorylation at Tyr397, creating a binding site for the SH2 domain of the Src family kinases Src and Fyn (Tomar et al. 2009). Protein tyrosine phosphatase (PTP)-mediated Tyr527 dephosphorylation (Zeng et al. 2003) and the association of FAK phosphoTyr397 with the Src SH2 domain disrupt the Src autoinhibitory conformation by displacing the interaction between the Src SH2 domain and the Src C-terminal phosphoTyr527. Once Src switches to its extended conformation, auto- phosphorylation at Tyr416 of Src synergistically promotes the activation of Src (Schaller et al. 1994; Xu et al. 1999). Activated Src then phosphorylates FAK at Tyr576/577 residues in its kinase activation loop to induce maximal activation of FAK and formation of an activated FAK-Src kinase complex (Schlaepfer et al. 1996; Tomar et al. 2009). This complex subsequently phosphorylates FAK-bound scaffolding proteins such as Cas and paxillin, and their tyrosine phosphorylation leads to the recruitment of further downstream signaling molecules to FAs, resulting in the activation of Rho family small GTPases via multiple pathways to promote FA maturation, stress fiber formation, and eventually cell migration. 27 Besides cell migration, FAK can also mediate the integrin outside-in signaling-induced MAPK and AKT activation to regulate cell survival and proliferation (Hsia et al. 2003). Src- dependent phosphorylation of FAK Tyr925 recruits the adapter protein Grb2, which not only recruits dynamin to FAs to induce integrin internalization and recycling (Ezratty et al. 2005), but also promotes the SOS-Ras-Raf signaling complex-mediated MAPK signaling cascade activation (Schlaepfer et al. 1994; Wang JG et al. 2005). FAK phosphoTyr397 also directly interacts with the regulatory p85 subunit of PI3K to activate the AKT signaling cascade (Mitra et al. 2005; Wu HJ et al. 2010). Src The non-receptor PTK Src is also called cellular Src (c-Src) because it was identified as the cellular counterpart of the Rous sarcoma virus transforming product known as v-Src (Parker et al. 1981). Src is the prototype member of to a kinase family named after it, which encompasses 10 other members including Fyn and Yes. Structurally, Src family kinases (SFKs) share similar domain arrangements: an N-terminal Src homology 3 (SH3) domain, a Src homology 2 (SH2) domain, and a kinase domain at the C-terminus (Fig. 1.4). Functionally their kinase activities are regulated by a common mechanism. In its inactive state, Src is phosphorylated at its C-terminal Tyr527 (inhibitory site), which interacts with its SH2 domain to fold the Src molecule into a hairpin-shape, blocking the access to its kinase domain. The interaction between the Src SH3 domain and the linker region between the SH2 and the kinase domains further contributes to the formation of this autoinhibitory conformation. During Src activation, disruption of the above two intramolecular interactions and dephosphorylation of the inhibitory Tyr527 residue release Src from its inactive, closed 28 conformation, facilitating the auto-phosphorylation of Tyr416 (a key tyrosine in the kinase activation loop) to achieve full activation of Src (Xu et al. 1999; Boggon et al. 2004; Alvarado et al. 2010). Src is ubiquitously expressed in mammalian cells. Overexpression of Src in chicken embryonic fibroblasts results in defective actin filament crosslinking and fewer focal adhesions, suggesting a function for Src in integrin-induced cell migration (Westhoff et al. 2004). The compensatory effects of Src family members made the study of the biological functions of SFKs very difficult since no major defects are observed in Src -/- fibroblasts. The generation of Src, Fyn, and Yes triple knockout (SYF) MEFs eventually revealed crucial defects in the absence of these SFKs (Klinghoffer et al. 1999). SYF fibroblasts exhibit severely impaired FA disassembly with reduced FAK phosphorylation, similar to cells treated with the specific SFK inhibitor PP2 or cells overexpressing a kinase-dead mutant of Src (Webb et al. 2004; Brunton et al. 2005; Yeo et al. 2006). Taken together, this evidence indicates that the cooperation of FAK and SFKs links integrin ligation to multiple downstream signaling events through the phosphorylation of downstream effectors such as Cas and paxillin. 1.5.2 Key downstream effectors: Cas and paxillin After integrin-stimulated FAK-Src complex activation, several key scaffolding proteins are recruited to FAs, behaving as molecular hubs for intracellular signaling pathways and organizing complex signaling networks (Defilippi et al. 2006; Wickstrom et al. 2009; Tikhmyanova et al. 2010). 29 p130Cas The scaffolding protein Cas is a 130 kD protein ubiquitously expressed in all vertebrates (Sakai et al. 1994). Cas knockout mice are embryonic lethal, and Cas -/- MEFs exhibit disorganized actin stress fibers and defective cell migration, suggesting that Cas is involved in the regulation of cell motility (Cary et al. 1998; Honda et al. 1998; Klemke et al. 1998). Cas contains multiple protein-protein interaction domains, including an N-terminal SH3 domain, a substrate domain (SD), a Src binding domain (SBD), and a Cas C-terminal homology (CCH) domain (Fig. 1.5). Many binding partners of Cas have been identified in FAs, such as FAK, PTP1B, PTP-PEST, C3G, and Dock180, all of which can bind to the Cas SH3 domain (Burnham et al. 1996; Hsia et al. 2003); Crk and Nck, which interact with phosphotyrosine sites in the SD domain of Cas (Burnham et al. 1996; Schlaepfer et al. 1997); Src, which directly binds to the Src binding domain of Cas (Sakai et al. 1994; Nakamoto et al. 1996); and PI3K, Ajuba and BCAR3, which interact with the CCH domain (Gotoh et al. 2000; Li E et al. 2000; Pratt et al. 2005). Upon integrin stimulation Cas is highly tyrosine phosphorylated, especially in its SD. This contains fifteen YxxP motifs, ten of which can be efficiently phosphorylated (Songyang et al. 1993; Sakai et al. 1994). FAK catalytic activity is not required for SD phosphorylation, but the interaction of FAK with the Src SH2 domain is necessary to recruit Src to phosphorylate the SD of FAK-associated Cas. Thus Src is the major kinase that catalyzes the phosphorylation of Cas during integrin signaling (Ruest et al. 2001). There are two known mechanisms of Cas SD tyrosine phosphorylation: 1) using FAK as a platform, the Cas SH3 domain binds to FAK via FAK poly-proline motifs, and FAK phosphoTyr397 forms a docking site for the SH2 domain of Src, such that Src is recruited to phosphorylate Cas in the 30 SD; and 2) the SH3 domain of Src binds directly to the Cas SBD and Src phosphorylates Cas in the SD. After activation by tyrosine phosphorylation, the Cas SD recruits downstream effectors to FAs, notably the SH2/SH3 adaptors Crk and Nck that can activate the GEFs Dock180 and C3G. Dock180 can promote Rac activation at the leading edge to promote protrusions, whereas C3G can activate another small GTPase, Rap1, to further promote integrin activation and clustering, resulting in persistent directional cell migration (Gotoh et al. 1995; Bouton et al. 2001; Ohba et al. 2001; Chodniewicz et al. 2004; Defilippi et al. 2006; Delorme-Walker et al. 2011). The phosphorylation of the Cas SD has been detected predominantly at FAs and at the cell periphery in nascent integrin adhesion sites, suggesting the activation of Cas may depend on its localization to FAs (Fonseca et al. 2004; Ballestrem et al. 2006). Two domains of Cas mediate its targeting to adhesion complexes: 1) the N-terminal SH3 domain of Cas associates with FAK to recruit Cas to focal adhesions; 2) the CCH domain of Cas is also necessary for its focal adhesion localization via an unknown mechanism. Both domains are required for optimal FA recruitment and tyrosine phosphorylation of Cas (Nakamoto et al. 1997; Harte et al. 2000; Donato et al. 2010). Paxillin Paxillin is a 68 kD multi-domain FA protein that functions as a molecular platform to facilitate integrin signaling (Brown MC et al. 2004; Harburger et al. 2009). Paxillin-null mouse embryos display abnormal development of the heart and somites at E7.5-E8.5 and die at E9.5 days, suggesting that paxillin is essential for embryonic development (Hagel et al. 2002; Brown MC et al. 2004). Paxillin -/- MEFs exhibit defective lamellipodia maintenance 31 and focal adhesion turnover, leading to delayed cell spreading and reduced motility (Webb et al. 2004). The N-terminal region of paxillin contains five leucine-rich motifs (LD) that interact with the actin-binding protein vinculin and the signaling protein FAK (Turner et al. 2001; Hoellerer et al. 2003; Deakin et al. 2008) (Fig. 1.5). The LD4 motif of paxillin, via the paxillin kinase linker (PKL), recruits the Cdc42/Rac guanine nucleotide exchange factor PIX, PAK, and the SH2-SH3 adapter protein Nck to FAs. The formation of paxillin-PKL-PIX- PAK-NCK complex further facilitates the activation of Rac and Cdc42 to promote FA formation and cell migration (Turner et al. 1999; Brown MC et al. 2004). Paxillin also contains an N-terminal poly-proline region, which interacts with the Src SH3 domain (Webb et al. 2004). Four Lim domains are present at the C-terminus of paxillin, and have been reported to associate with the protein tyrosine phosphatase PTP-PEST to promote FA disassembly (Brown MC et al. 2004). In addition, paxillin binds directly to the cytoplasmic tails of α4, β1, and β3 integrin to mediate integrin-stimulated cell migration (Liu et al. 1999; Chen LM et al. 2000). The numerous serine, threonine, and tyrosine phosphorylation sites in paxillin either regulate the conformation of paxillin or function as binding sites for SH2/PTB domain-containing proteins (Deakin et al. 2008). As a substrate of Src (Glenney et al. 1989), integrin-simulated phosphorylation of paxillin Tyr31 and Tyr118 creates SH2 domain binding sites for the adapter protein Crk (Schaller et al. 1995) and p120RasGAP (Tsubouchi et al. 2002), which can lead to Rac activation and promotes lamellipodia formation (Kiyokawa et al. 1998). Serine phosphorylation of paxillin promotes the formation of the paxillin-GIT-PIX-PAK complex to promote FA assembly (Roach et al. 1998; Nayal et al. 2006). Through the above interactions, paxillin functions as a molecular platform, which not 32 only recruits its binding proteins to FAs but also transmits integrin-initiated signaling to downstream effectors to promote cell migration (Brown MC et al. 2004; Deakin et al. 2008). 1.5.3 The integrin signaling adapter BCAR3, a member of the novel SH2 domain-containing protein (NSP) family BCAR3 was first identified as a 95KD protein in a screen for genes responsible for conferring anti-estrogen resistance in breast cancer cells (van Agthoven et al. 1998). Later studies showed that BCAR3 can enhance Src activity and increase Cas tyrosine phosphorylation to promote Cas downstream signaling and cancer properties such as invasion, as well as confer anti-estrogen resistance (Riggins et al. 2003; Schuh et al. 2010). BCAR3 contains an N-terminal SH2 domain and a C-terminal guanine nucleotide exchange factor (GEF)-like domain. The BCAR3 C-terminal region has a 3D structure similar to the CDC25-like domain in SOS, a well-known Ras GEF in the MAPK pathway. Overexpression of murine BCAR3 in COS7 cells induces the activation of Ral, Rap, and R- Ras, which suggested that it functions as a GEF (Gotoh et al. 2000). However, the same group later observed that BCAR3 had no GEF activity on RalA, Rap1A, R-Ras, and H-Ras as assessed by pulldown assays in B lymphoid cells (Cai et al. 2003). Recent structural analysis revealed that the BCAR3 GEF domain lacks a key structure required for function as a GEF and is structurally more suited for Cas binding (Mace et al. 2011). The BCAR3 C- terminal region is required for BCAR3-Cas interaction and downstream integrin signaling (Cai et al. 1999; Riggins et al. 2003). BCAR3-enhanced migration is abolished when the BCAR3 GEF domain responsible for Cas binding is deleted or mutated (Riggins et al. 2003; Vanden Borre et al. 2011). The BCAR3 N-terminal SH2 domain is also required for 33 downstream integrin signaling, since the deletion or point mutation of the BCAR3 SH2 domain failed to induce Rac1 activation and confer anti-estrogen resistance in breast cancer cells (Felekkis et al. 2005; Kowalski et al. 2005). However, the specificity of the BCAR3 SH2 domain is still unknown, and no intracellular ligand has been identified yet (Vanden Borre et al. 2011). BCAR3 (also known as NSP2, Shep2, or AND-34) belongs to a novel SH2 domain- containing protein (NSP) family, and all NSPs share similar domain arrangement (van Agthoven et al. 1998; Cai et al. 1999). Overexpression of NSP3 (also called Shep1 or Chat) in 293T cells induces significant activation of Rap1 through the Cas-Crk-C3G signaling pathway (Sakakibara et al. 2002). Overexpression of NSP1 induces Cas tyrosine phosphorylation and Rac/Cdc42 activation in MCF7 cells (Near et al. 2007; Makkinje et al. 2009), suggesting that NSP1 and NSP3 can also regulate integrin signaling. Later studies further showed that EphB2-induced Tyr635 phosphorylation in the NSP3 GEF domain disrupts the NSP3-Cas association and impairs cell migration (Dail et al. 2004), and NSP3- null cells exhibit impaired Src activation, and reduced Cas-Crk association and Cas tyrosine phosphorylation (Roselli et al. 2010). In summary, all three members of this NSP family are Cas-binding proteins and promote integrin-stimulated cell migration (Lu et al. 1999; Sakakibara et al. 2000; Makkinje et al. 2009). 1.6 PTPs control tyrosine kinase pathways in both negative and positive manners As the protein tyrosine kinases (PTKs) add phosphate groups from ATP to tyrosyl residues in proteins, the dephosphorylation that removes phosphate groups from tyrosine residues by hydrolysis is catalyzed by protein tyrosine phosphatases (PTPs). So far about 108 34 PTP genes have been identified in the human genome, and this number is comparable to the number of PTK genes, suggesting that PTPs are as equally important as PTKs in modulating cell signaling systems (Alonso et al. 2004; Tonks 2006; Kapp et al. 2007; Sacco et al. 2012). The coordinated and balanced actions of PTKs and PTPs are therefore critical for proper cellular functions such as cell survival, proliferation, and migration (Larsen et al. 2003). PTKs and PTPs are important regulators of many physiological processes, and their exact roles are cell type- and signal-dependent. Early indications of PTP functions in cell migration came from observations that phosphate analogue vanadate (a PTP inhibitor) treated fibroblasts exhibited delayed adhesion and spreading on a fibronectin (FN) coated surface (Edwards et al. 1991), and microinjection of PTPs into cells induced the disassembly of focal adhesions and breakdown of stress fibers (Schneider et al. 1998; Larsen et al. 2003). Indeed some PTPs, such as PTEN, are negative regulators of cell motility (Yamada et al. 2001; Sze et al. 2011). The famous tumor suppressor PTEN (phosphatase and tensin homology on chromosome 10) is a dual-specific phosphatase that contains the PTP catalytic signature motif and has the ability to dephosphorylate both phosphotyrosyl proteins and lipids (Myers et al. 1997; Myers et al. 1998). As a lipid phosphatase, PTEN dephosphorylates the 3 position of phosphatidyl- inositol-3,4,5-trisphosphate (PIP3) to phosphatidylinositol-4,5-bisphosphate (PIP2), whereas PI3K catalyzes the reverse process by adding a phosphate group to PI(4,5)P2 at the 3 position (Maehama et al. 1998; Yamada et al. 2001; Sze et al. 2011). PI3K and PTEN are major proteins for establishing and maintaining cell polarity that is crucial for directional migration. PTEN knockout mice are early embryonic lethal at E7.5 with severe central nervous system defects (Di Cristofano et al. 1998). Overexpression of PTEN in cells results in decreased cell 35 migration in a wound healing assay (Tamura et al. 1998), whereas PTEN -/- fibroblasts and PTEN-depleted cells exhibit increased cell migration, which is induced by hyperactivation of Rac and Cdc42 (Liliental et al. 2000). Some PTPs are positive regulators of cell motility, such as PTP1B and SHP2 (Larsen et al. 2003). PTP1B (PTPN1) is a non-receptor PTP and PTP1B knockout mice have no obvious disease phenotype and normal fetal viability (Elchebly et al. 1999). PTP1B-null fibroblasts and fibroblasts expressing catalytically inactive mutant PTP1B exhibit impaired cell spreading and migration upon FN stimulation (Arregui et al. 1998; Cheng et al. 2001). These cells also display reduced FAK and Src activation and hypophosphorylation of the downstream effector Cas, suggesting that PTP1B may regulate FAK-Src-Cas signaling to promote cell migration. SHP2 (PTPN11) is another ubiquitously expressed intracellular PTP that contains two SH2 domains. Besides playing important roles in MAPK signaling, SHP2 also acts on multiple targets to regulate integrin signaling and cell migration (Neel et al. 1997; Zheng H et al. 2009). Homozygous SHP2 mutant mice die around day E8.5-E10.5 and have similar abnormalities to FAK -/- and FN -/- embryos (Ilic et al. 1995; Schaller 1996; Yu et al. 1998). Cells expressing the loss-of-function mutant SHP2 Δ46-110 or dominant-negative mutant (SHP2-C/S) SHP2 display enhanced formation of FAs and stress fibers, resulting in decreased FN-stimulated cell migration (Yu et al. 1998; Kodama et al. 2000), whereas expression of a dominant-active form of SHP2 promotes the formation of protrusions through inhibition of RhoA at the leading edge, indicating that SHP2 is involved in the positive regulation of cell migration (Kodama et al. 2001; Lacalle et al. 2002). Some PTPs can regulate cell migration both positively and negatively. PTP-PEST (PTPN12) is a ubiquitously expressed cytoplasmic PTP that contains proline-glutamic acid- 36 serine-threonine (PEST)-rich sequences (Yang Q et al. 1993; Cote et al. 1998). PTP-PEST knockout mice exhibit embryonic lethality at E10.5 with severe defects in blood vessel and liver development (Sirois et al. 2006). Analyses of embryonic fibroblasts from PTP-PEST- deficient mice showed that PTP-PEST has two major roles in integrin-induced cell migration: 1) inhibiting membrane protrusions and cell spreading, and 2) promoting FA turnover and tail retraction (Larsen et al. 2003; Hendriks et al. 2012). PTP-PEST possesses an N-terminal PTP domain and a C-terminal substrate-interacting region, which is involved in direct binding and dephosphorylation of scaffolding proteins including Cas and paxillin (Garton et al. 1996; Shen Y et al. 1998; Angers-Loustau et al. 1999). Cas was initially identified as a substrate of PTP-PEST by expression of a substrate-trapping mutant of PTP-PEST in HeLa and COS cells. Later, the same group showed that the overexpression of PTP-PEST in rat fibroblasts impaired Cas phosphotyrosine-dependent downstream signaling and cell migration (Garton et al. 1996; Garton et al. 1999). Interestingly, the PTP-PEST-deficient fibroblasts display more protrusions and faster spreading, while FN-stimulated cell migration remains defective (Angers-Loustau et al. 1999; Jamieson et al. 2005). PTP-PEST-null cells have an increased number of and bigger FAs that impede tail detachment, whereas PTP- PEST overexpressing cells induce fewer and smaller FAs that are incapable of providing sufficient anchorage for stress fiber contraction, resulting in defective motility in both situations (Larsen et al. 2003). Molecular studies of PTP-PEST -/- fibroblasts revealed that the increased protrusions are due to enhanced tyrosine phosphorylation of Cas, paxillin, and FAK and hyperactivation of their downstream effector Rac1 (Shen Y et al. 1998; Angers- Loustau et al. 1999; Jamieson et al. 2005). PTP-PEST disrupts the paxillin-PKL-PIX complex through direct dephosphorylation of paxillin and PKL (Schoenwaelder et al. 2000; 37 West et al. 2001), leading to inhibition of Rac-mediated membrane protrusions (Jamieson et al. 2005). Another report showed that PTP-PEST can directly dephosphorylate and inhibit 1) the Rac GEF Vav2 to reduce Rac-mediated membrane protrusions at the leading edge and 2) p190RhoGAP activation to promote RhoA-mediated tail retraction at the cell rear, suggesting that precisely regulated local activation of PTP-PEST is essential for proper cell migration (Sastry et al. 2006). PTP-PEST also associates with and dephosphorylates FAK Tyr397 through a peptidyl-prolyl cis/trans isomerase PIN1-mediated mechanism in rat fibroblasts (Zheng Y et al. 2009; Zheng Y et al. 2011), and overexpression of PTP-PEST in colon cancer Caco-2 cells induces dephosphorylation of Src Tyr416 and inhibits Src activation (Mathew et al. 2008). These results indicate that PTP-PEST can target multiple signaling proteins to control cell motility, and the local activation of PTP-PEST activity must be precisely regulated to maintain an equilibrium of forward protrusion and formation of new FAs at the leading edge with the disassembly of FAs and detachment from the substratum at the trailing edge of a cell. PTPα is another ubiquitously expressed receptor tyrosine phosphatase that functions as a positive regulator of cell migration (Pallen 2003). Studies of PTPα overexpression and knockout systems revealed that PTPα is an important activator of SFKs, such as Src and Fyn, through dephosphorylation of their inhibitory C-terminal phosphotyrosine. When studying PTPα-induced transformation in rat embryonic fibroblasts, Zheng et al. (1992) observed increased Src kinase activity and reduced tyrosine phosphorylation at its C-terminus tyrosine residue (Tyr527), and further confirmation that PTPα functions as a positive regulator of SFKs was obtained from studies of PTPα-null mice (Ponniah et al. 1999; Su et al. 1999). Since SFKs are important in promoting cell migration, the above studies suggest that PTPα 38 also plays a role in this process (Westhoff et al. 2004). The detailed mechanisms of PTPα in regulating cell motility are discussed in the next section. In summary, the concerted activities of PTKs and PTPs in integrin signaling are essential for proper cell migration. 1.7 PTPα plays an important role in integrin signaling PTPα was discovered in 1990 by using the phosphatase domain of CD45 (also known as leukocyte common antigen, LCA) to screen mouse and human brain cDNA libraries from mouse and human, and mapped to human chromosome 20 and mouse chromosome 2 (Jirik et al. 1990; Kaplan R et al. 1990; Krueger et al. 1990; Matthews et al. 1990; Sap et al. 1990). This phosphatase is ubiquitously expressed in the body but is most abundant in the brain and kidney (Matthews et al. 1990; Sap et al. 1990). Since multiple groups cloned PTPα around the same period, various names were given to this protein, including receptor protein tyrosine phosphatase alpha (PTPα), receptor-linked protein-tyrosine-phosphatase alpha (RPTPα), and LCA-related peptide (LRP) (Jirik et al. 1990; Kaplan R et al. 1990; Sap et al. 1990). According to the nomenclature of the genes encoding PTP superfamily members (Alonso et al. 2004), PTPα is encoded by the gene PTPRA and the protein is also sometime referred to as PTPRA. For consistency, the name “PTPα” will be used in this thesis. When studying PTPα-induced transformation in rat embryonic fibroblasts, Zheng et al. (1992) observed increased Src kinase activity and reduced tyrosine phosphorylation at its C- terminus (Tyr527), and another group showed that exogenous PTPα expression induces the same effects in P19 embryonal carcinoma cells (den Hertog et al. 1993). In contrast, PTPα- null cells from PTPα-deficient mice display decreased Src kinase activity and enhanced 39 tyrosine phosphorylation at Tyr527 (Ponniah et al. 1999; Su et al. 1999). An experiment using antisense oligo nucleotides to inhibit endogenous PTPα expression showed that 85% down-regulation of PTPα expression led to 80% reduction of Src activity in 3T3-L1 adipocytes (Arnott et al. 1999). The above studies demonstrated that SFKs are major substrates of PTPα, suggesting that PTPα may function as a positive regulator of SFKs to control cell behaviors, such as cell survival, proliferation, and migration. 1.7.1 Structure of PTPα PTPα contains an extracellular domain (ECD), a transmembrane region and two tandem phosphatase domains (D1 and D2) (Jirik et al. 1990; Kaplan R et al. 1990; Krueger et al. 1990; Matthews et al. 1990; Sap et al. 1990) (Fig. 1.6). The ECD of PTPα is short (~133 residues in the isoform4) and heavily N- and O-glycosylated compared to other receptor PTPs. Although the predicted molecular weight is only ~90 kD based on its primary sequence, the highly glycosylated PTPα runs at ~130 kD on SDS-PAGE (Sap et al. 1990; Daum et al. 1994; Tonks 2006). Contactin, a neural cell adhesion molecule, is so far the only identified extracellular interacting partner of PTPα, and the cis-interaction of these two receptors suggests PTPα may play a role in neural signaling (Zeng et al. 1999). Four alternatively spliced isoforms of human PTPα (uniprot entry P18433) have been identified that are 99% identical and differ slightly in their ECD regions. Overexpression of the most studied isoform (isoform4) in NIH3T3 cells induces anchorage-independent proliferation and colony formation in soft agarose, whereas overexpression of isoform3 lacks the ability to promote these effects, suggesting that variation in the ECD of PTPα may affect its 40 transforming ability (Tremper-Wells et al. 2010). To be consistent with most publications, all annotation of PTPα amino acid (aa) residues will refer to the 793 aa isoform4. PTPα is a type I transmembrane protein that has a single transmembrane domain (aa 134- 156). Two serine residues in the juxtamembrane region can be phosphorylated by PKC and CaMKIIα to enhance PTPα phosphatase activity (den Hertog et al. 1995; Bodrikov et al. 2008). It has been proposed that phosphoSer180/204 are required for PTPα-mediated Src activation in NIH3T3 cells during mitosis (Zheng XM et al. 2002). 12-O-tetradecanoyl- phorbol-13-acetate (TPA) stimulation induces PKC-dependent phosphorylation of PTPα Ser180/204 in NIH3T3 cells (Tracy et al. 1995). This TPA-stimulated serine phosphorylation of PTPα at Ser180/Ser204 increases the PTPα phosphatase activity 2~3 fold, and leads to Src Tyr527 dephosphorylation and activation in A7r5 rat vascular smooth muscle cells (Brandt et al. 2003). The PKC and PTPα-mediated Src activation has been shown to activate the Ras/Sos complex and downstream MAPK cascade in A431 cells and C2C12 skeletal muscle cells (Stetak et al. 2001; Buitrago et al. 2011). However, a recent study proposed that the phosphorylation of PTPα Ser204 actually inhibits Src activation, since mitosis induces PP2A-mediated dephosphorylation of PTPα Ser204 to facilitate PTPα-Src interaction and promote Src activation (Vacaru et al. 2010). The membrane proximal PTPα-D1 domain carries almost all of its phosphatase activity towards 1) phosphotyrosyl peptides in vitro and 2) Src, its substrate in vivo (Wang Y et al. 1991; den Hertog et al. 1993; Wu L et al. 1997). All PTPα isoforms show similar catalytic activity towards the small molecule substrate para-nitrophenyl phosphate (pNPP) and c-Src (Kapp et al. 2007). 41 The membrane distal PTPα-D2 has only ~10% intrinsically catalytic activity towards pNPP (Wang Y et al. 1991). Several studies showed that point mutations at two key residues in the PTPα D2 domain, V555Y in the phosphotyrosine recognition loop (KNRY motif) and E690D in the WPD loop, can actually restore its phosphatase activity towards pNPP and phosphotyrosine-containing peptides to a level similar to that of the D1 domain (Lim KL et al. 1998; Buist et al. 1999). Not being very active catalytically, PTPα-D2 has been proposed to have multiple important regulatory functions, such as stabilizing the PTPα-D1 domain in the correct orientation in solution (Nam et al. 1999; Nam et al. 2005), participating in dimerization-mediated inhibition of PTPα activity (Blanchetot et al. 2000), functioning as a redox sensor through the PTPα Cys723 residue to regulate PTPα activity (Blanchetot et al. 2002; van der Wijk et al. 2003), and affecting the substrate binding specificity of PTPα-D1 (Wang Y et al. 1991; Vacaru et al. 2010). Soon after its discovery, PTPα was found to be constitutively tyrosine phosphorylated at Tyr789 in NIH3T3 cells (den Hertog et al. 1994; Su et al. 1994). This Tyr789 residue is the only phosphotyrosine site identified in PTPα to date, and this site is located four residues away from the C-terminal amino acid (den Hertog et al. 1994). The phosphorylation of PTPα at Tyr789 can be regulated by multiple pathways. Activated growth factor receptors and integrin-stimulated SFKs have been shown to induce phosphorylation of the Tyr789 residue (Chen M et al. 2006; Chen SC et al. 2009). Oxidative stress can negatively regulate PTPα Tyr789 phosphorylation in fibroblasts (Hao et al. 2006). Although PTPα has been proposed to trans-auto-dephosphorylate itself (den Hertog et al. 1994), the exact mechanism of PTPα dephosphorylation is still unknown. 42 An unphosphorylatable mutant of PTPα (Y789F) displays similar phosphatase activity towards the synthetic peptide Raytide or myelin basic protein to wildtype PTPα (Zheng XM et al. 2000). Expression of mutant PTPα-Y789F rescues the defective Src Tyr527 dephosphorylation and activation observed in PTPα-null MEFs (Chen M et al. 2006), suggesting that Tyr789 does not affect the catalytic activity of PTPα and PTPα-mediated Src activation. As PTPα is a transmembrane receptor-like protein, the phosphoTyr789 residue has the potential to function as a docking site for the recruitment of other signaling proteins to the membrane. The ubiquitously expressed adapter protein Grb2 was identified as a binding partner of PTPα through the interaction of the SH2 domain of Grb2 with phosphoTyr789 of PTPα (den Hertog et al. 1994; Su et al. 1994). Grb2 plays a significant role in growth factor signaling, where it functions to recruit Son of sevenless (Sos), a Ras GEF (guanine nucleotide exchange factor), to numerous growth factor receptors to promote Ras-Raf-MEK-Erk (MAPK) pathway activation (Giubellino et al. 2012). However, PTPα- bound Grb2 does not interact with Sos (den Hertog et al. 1996), leaving the function of the PTPα-Grb2 complex a mystery. 1.7.2 PTPα in integrin signaling Overexpression of PTPα in A431 epidermoid carcinoma cells induces increased cell- substratum adhesion together with enhanced Src activation, suggesting a role of PTPα in mediating cell adhesion (Harder et al. 1998). PTPα-null MEFs show delayed integrin- stimulated cell spreading and migration compared to wildtype MEFs (Su et al. 1999; Zeng et al. 2003). Molecular investigations revealed that integrin-induced dephosphorylation of Src/Fyn inhibitory phosphotyrosine and the activation of these SFKs were impaired in PTPα- 43 null MEFs (Ponniah et al. 1999; Su et al. 1999). Wildtype MEFs treated with the specific SFK inhibitor PP2 also display retarded cell spreading on fibronectin (FN), a defect that is morphologically similar to that of PTPα-/- MEFs (Zeng et al. 2003). Some of the downstream effectors of Src/Fyn, such as paxillin and Cas, also exhibit reduced tyrosine phosphorylation in integrin-stimulated PTPα-/- MEFs (Su et al. 1999; Chen M et al. 2006). These results demonstrate that PTPα plays a positive role in early integrin signaling by dephosphorylating and activating Src/Fyn. The integrin-induced tyrosine phosphorylation of FAK is also defective in PTPα-null MEFs (Su et al. 1999). Integrin activation induces FAK auto-phosphorylation at Tyr397 and promotes SFK-mediated FAK Tyr576/577 phosphorylation for optimal activation of FAK (Schaller et al. 1994). A later study showed that the defective phosphorylation of FAK Tyr397 in PTPα-/- MEFs is due to impaired Src/Fyn activation and can be rescued by the re- introduction of wildtype PTPα (Zeng et al. 2003). Collectively, the above results indicate that PTPα is important in early integrin signaling by promoting activation of the central signaling molecules Src and FAK that are critical early regulators of cell spreading and migration. In 2006, our group discovered that the phosphorylation of PTPα Tyr789 is positively regulated by integrin signaling in a Src/Fyn/Yes- and FAK-dependent manner (Chen M et al. 2006). Using an adenovirus-mediated PTPα reconstitution system, wildtype, catalytically inactive (C433S/C723S), or unphosphorylatable (Y789F) PTPα was re-introduced into PTPα-/- MEFs. PTPα-null cells exhibit impaired Src/Fyn and FAK activation, delayed cell spreading, and defective cell migration. Wildtype PTPα rescued all these defects, whereas catalytically inactive PTPα did not rescue any of them. Interestingly, the Y789F mutant of PTPα was found to effectively catalyze Src/Fyn and FAK activation, suggesting that PTPα 44 Tyr789 phosphorylation is not required for these early integrin-stimulated PTPα-dependent events. In accord with this finding, another study investigating PTPα actions in mitosis showed that the interaction of PTPα with its substrate Src is independent of either this Tyr789 residue of PTPα or the SH2 domain of Src, confirming that PTPα Tyr789 is not required for the PTPα-mediated Src activation (Vacaru et al. 2010). However, in contrast to the non-requirement for PTPα Tyr789 in integrin-induced Src/Fyn activation, our lab also found that unphosphorylatable Y789F PTPα failed to restore defective cell spreading, focal adhesion formation, and cell migration, suggesting that PTPα phosphoTyr789 is necessary for signaling downstream of Src-FAK (Chen M et al. 2006). These findings distinguished two roles of PTPα in integrin signaling: 1) PTPα-mediated SFK activation that requires PTPα catalytic activity but is independent of Tyr789 phosphorylation, and 2) PTPα phosphoTyr789-mediated Src/FAK downstream signaling events that promote cell spreading and migration (Chen M et al. 2006). PTPα has been shown to translocate to FAs when nerve growth cones are plated on laminin (Helmke et al. 1998), and this FA localization of PTPα was proposed to be dependent on its phosphorylation at Tyr789 (Lammers et al. 2000). PTPα has been demonstrated to co-localize with integrin αv subunit upon integrin activation, and a complex of integrin αv and PTPα can be co-immunoprecipitated after cross-linking, suggesting PTPα Tyr789 may play a role in nascent FA complex formation (von Wichert et al. 2003). In summary, these studies demonstrate that PTPα Tyr789 plays an unknown but key role in promoting integrin-induced cytoskeletal reorganization events in cell spreading and migration. Identification of the detailed signaling mechanism involving PTPα 45 phosphoTyr789 and its role(s) in integrin-induced focal adhesion formation and cell migration is the main focus of my thesis study. 1.8 Rationale and hypothesis The studies described above reveal a role for PTPα Tyr789 phosphorylation in integrin signaling. Since PTPα Tyr789 phosphorylation is required for integrin-induced cell spreading and migration, I hypothesize that phosphoTyr789 of PTPα mediates important signaling events to promote integrin-regulated cell motility. The first aim of this study was to identify the immediate signaling targets/effectors of PTPα phosphoTyr789, and the second aim was to investigate the detailed molecular mechanism(s) employed by PTPα phosphoTyr789 to regulate integrin signaling to promote cell migration. 46 E A D B C F Figure 1.1 Schematic diagram of cell migration cycle Following signal cues, (A) a cell establishes polarized structures along the axis of movement and is ready to move. (B) The cell generates membrane protrusion termed filopodium and lamellipodium at the leading edge. (C) The membrane protrusion retracts to form a membrane ruffle. (D) The protrusion is stabilized and anchored to substratum by nascent focal adhesions (FAs). (E) Stress fiber contraction generates the force to pull the cell body forward. (F) FA disassembly and tail detachment from the substratum at the trailing edge. (adapted from Nagano et al. 2011) nucleus focal adhesions substratum ruffle protrusion migrating direction 47 Figure 1.2 Diagram of a filopodium and lamellipodium Two main protrusive structures are the lamellipodium and filopodium, found at the leading edge of a migrating cell. (A) A filopodium is a finger-like protrusive structure containing parallel bundles of actin filaments. (B) the lamellipodium is a sheet-like protrusion, that is supported by an underlying branched actin filament array. A. filopodium B. lamellipodium nucleus leading edge lamella trailing edge Figure 1.3 The activation cycle of Rho family small GTPases Rho family GTPases switch between an inactive GDP-bound form and an active GTP-bound form. This process is regulated by GEFs, GAPs, and GDIs. Membrane Rho GDP PP GDIs GAPs P GEFs Cytoplasm GDIs 48 49 Src SH3 Kinase Tyr416 SH2 Tyr527myristoylation KinaseFERM Tyr397 Tyr577 Tyr576 Tyr925 Pro‐rich Pro‐rich FAK Figure 1.4 Domain structures of FAK and Src Non-receptor protein tyrosine kinase FAK contains an N-terminal FERM (protein 4.1,ezrin, radixin, and moesin homology) domain, a central kinase domain, a proline- rich region, and a FA targeting domain (FAT) at C-terminus. Src consists of three domains, an N-terminal SH3 domain, a central SH2 domain and a tyrosine kinase domain also known as SH1 domain. Src associates with the cytoplasmic face of the plasma membrane through its N-myristoylation. All phosphorylation sites are labeled to their position. FAT 50 Paxillin 1 2 3 4 5 Lim2 Lim3 Lim4Lim1 Tyr31 Tyr118 Acetylation Pro‐rich Leu‐rich domain Cas Tyr410 Substrate Domain (SD)SH3 SBD Tyr387 Tyr362 Tyr327 Tyr292 Tyr287 Tyr269 Tyr267 Tyr249 Tyr234 Tyr224 Tyr222 Tyr139 Tyr134 Tyr134 Figure 1.5 Domain structures of scaffolding proteins Cas and paxillin Cas contains multiple protein-protein interaction domains, an N-terminal SH3 domain, a substrate domain (SD), and a Src binding domain (SBD) at its C-terminus. The N-terminal region of paxillin possesses five leucine-rich motifs, a proline-rich SH3 domain-binding site, and four LIM domains at its C-terminus, which can mediate interactions with many other proteins. All phosphorylation sites are labeled to their position. 51 S ECD T Catalytic domain (D1) Catalytic domain (D2) 1 19 133 156 232 492 524 782 Extracellular domain Cytoplasmic domain Ser180 Ser204 Tyr789 Figure 1.6 Domain structure of PTPα PTPα contains a signal peptide (S), an extracellular domain (ECD), a transmembrane region (T), two tandem phosphatase domains (D1 and D2). Two serine phosphorylation sites (Ser180 and Ser204) and one tyrosine phosphorylation site (Tyr789) are indicated. 52 Chapter 2: Materials and Methods 2.1 Cell lines and cell culture Wild type (PTPα+/+) and PTPα-/- mouse embryonic fibroblasts (MEFs) have been described previously (Zeng et al. 2003). The RD rhabdomyosarcoma cell line was from ATCC (CCL-136). All cells were grown in uncoated culture-grade dishes (adherent cells) in Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum and penicillin/streptomycin. 2.2 Fibronectin stimulation Prior to timed stimulation of integrin signaling, the cells were starved in medium containing 0.5% fetal bovine serum for 18 h. They were then detached from the dish using 0.05% trypsin-EDTA (Invitrogen). The trypsinization was stopped with 0.5 mg/ml soybean trypsin inhibitor (Invitrogen) in DMEM, and the cells were washed twice with serum-free medium. After centrifugation, the cells were re-suspended in serum-free medium containing 0.1% bovine serum albumin and maintained at 37 °C for 1 h (suspension, or 0 min). Suspended cells were plated onto fibronectin-coated plates (10 5 cells/ml) and incubated at 37°C for 15 min before harvesting. The fibronectin-coated dishes were prepared ahead of time by incubating dishes containing 10 μg/ml fibronectin (Chemicon International, Inc.) in phosphate-buffered saline overnight at 4 °C. Before use, the dishes were washed twice with PBS and then incubated with serum-free DMEM at 37 °C for 1 h. 53 2.3 Antibodies and immunological detection reagents PTPα and PTPα-phosphoTyr789 antibodies have been described previously (Chen M et al. 2006). Antibodies to Cas, Crk, FAK, Rac1, and Cdc42 were from BD Transduction Laboratories. Antibody to Src (v-Src) was from Calbiochem. Antibody to BCAR3 was from Bethyl Laboratories, Inc. Antibodies to PAK, RhoGDI, and GST were from Santa Cruz Biotechnology, Inc. Anti-VSVG and -actin antibodies were from Cell Signaling. Anti- phosphotyrosine antibody (4G10) was from Upstate Biotechnology. Phosphorylation site- specific antibodies to Cas Tyr410 and PAK Ser144 were from Cell Signaling and those to Src Tyr416 and Tyr527 were from Biosource. 2.4 BCAR3 expression constructs The plasmid pCMV6-AC-BCAR3 containing full-length human BCAR3 cDNA ( OriGene Technologies, Inc., Rockville, Maryland USA) was used as a template in a PCR reaction (primers: F1, 5’-ACTGAAGCTTATGGCTGCAGGAAAATT-3’ and R1, 5’- AAAAAAAG CGGCCGCTCAAAGCTCTGCCTG -3’) to generate full-length wild-type BCAR3 cDNA with a 5’ HindIII site and a 3’ NotI site. This PCR product was cloned into pXJ40-FLAG vector that had been digested with HindIII and NotI to generate the expression plasmid pXJ40-FLAG-BCAR3 (WT). Site-directed mutagenesis using overlap extension PCR of the pCMV6-AC-BCAR3 template was performed to generate full-length BCAR3 cDNA with mutations in the codon for Arg177 (changing the expressed amino acid to Lys) in the SH2 domain. Separate PCR reactions were performed to produce two pieces of the complete BCAR3 cDNA that overlapped in sequence around the mutated region, using the primer pairs F1 (above) with R2 (5’-GGACAGAGAGTCTTTAACTAGGAAGTC-3’, mutated bases 54 underlined), and F2 (5’-GACTTCCTAGTTAAAGACTCTCTGTCC-3’, mutated bases underlined) with R1 (above). These PCR products were gel-purified and mixed to serve as templates in another PCR reaction using primers F1 and R1 to generate full-length mutant BCAR3 cDNA. This was cloned into HindIII- and NotI-digested pXJ40-FLAG to produce the expression plasmid pXJ40-FLAG-BCAR3 (R177K). The same procedure was used to generate the plasmid pXJ40-FLAG-BCAR3 (R478A) encoding full-length BCAR3 cDNA with mutations in the codon for Arg478 (changing the expressed amino acid to Ala) in the GEF domain. The primer pairs used for first round PCR reactions were F1 and R3 (5’- GGCCATGAATGCCGCTGTTGCCAA-3’, mutated bases underlined), and F3 (5’- TTGGCAACAGCGGCATTCATGGCC-3’, mutated bases underlined) and R1, followed by second round PCR on the mixed templates using primer pairs F1 and R1. The pXJ40- mCherry-BCAR3 plasmids expressing WT BCAR3 or BCAR3 with a mutant SH2 domain (R177K) were generated by EcoRI/Hind III excision of the FLAG tag from the appropriate BCAR3-expressing plasmids, and its replacement with PCR-generated mCherry sequence prepared from another plasmid template using appropriate primers with added EcoRI and HindIII sequences. All wild-type and mutant BCAR3 cDNA insert sequences and their in- frame cloning with the FLAG or mCherry tags were verified by sequencing. 2.5 PTPα and BCAR3 depletion, and re-expression The generation and use of adenovirus to express WT PTPα and Y789F PTPα in PTPα-/- MEFs was previously described (Chen M et al. 2006). The plasmids pXJ41-neo-VSVG- PTPα (WT) (Bhandari et al. 1998), pXJ41-neo-VSVG-PTPα (Y789F) (Chen M et al. 2006), and BCAR3-expressing plasmids were transfected into cells using Lipofectamine™ LTX 55 reagent (Invitrogen). Infected and transfected cells were cultured for 24 h prior to further experimentation. Small interfering RNA (siRNA) targeting murine BCAR3, human BCAR3, or human PTPα (ON-TARGETplus SMARTpool) and control non-targeting siRNA (ON- TARGETplus Non-Targeting Pool) were from Dharmacon. MEFs were transfected with 100nM BCAR3 siRNA (murine), and RD cells were transfected with 50nM BCAR3 siRNA or 20nM PTPα siRNA. Lipofectamine™ RNAiMAX reagent from Invitrogen was used for transfection. Cells were cultured for 48 h prior to further experimentation. The siRNA-resistant PTPα expression system was constructed by Hoa Le (Pallen Lab, unpublished data), which includes siRNAs and siRNA-resistant PTPα plasmids. Two siRNAs targeting the ECD of human PTPα (ON-TARGETplus, catalog numbers: J-004519- 06-0005 and J-004519-08-0005) and control non-targeting siRNA (ON-TARGETplus Non- Targeting Pool) were from Dharmacon. The siRNA targeting regions of human PTPα in plasmids pXJ41-neo-WT-hPTPα-mECD and pXJ41-neo-Y789F-hPTPα-mECD were replaced by two fragments from murine PTPα cDNA that contain silent mutations in these regions, leading to the above two human PTPα siRNAs no longer inhibit plasmid PTPα expression in siRNA-transfected cells. RD cells were co-transfected with 20nM of each siRNA and siRNA-resistant PTPα plasmids using Lipofectamine™ RNAiMAX reagent from Invitrogen. Cells were cultured for 48 h prior to further experimentation. 2.6 Cell lysis, immunoprecipitation, and immunoblot analysis Cells were harvested in lysis buffer (50 mM Tris pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Nonidet P-40, 2 mM Na3VO4, 1 mM phenylmethylsulfonyl fluoride (PMSF), 10 μg/ml 56 leupeptin, and 10 μg/ml aprotinin) and processed for immunoprecipitation and immunoblotting experiments as described (Zeng et al. 2003). 2.7 Membrane translocation assay The preparation of cell membrane and cytosolic fractions was carried out as described (Wegener et al. 2007). The PTPα+/+ and PTPα-/- MEFs were placed on fibronectin-coated dishes for 15 min as described above, and then the dishes were chilled on ice. The cells were washed twice with ice-cold PBS and lysed with hypotonic lysis buffer (10mM Tris-HCl pH7.5, 1.5mM MgCl2, 5mM NaCl, 1mM NaF, 0.2mM Na3VO4). Cells were then scraped from the dishes and after 20s sonication the lysate was centrifuged at 16,000×g at 4ºC for 15 min. The supernatant was transferred to a fresh tube, and this fraction is referred to as the cytosolic fraction (C), and the insoluble pellets were dissolved in 2X SDS Sample Buffer (membrane fraction, M). Fractionation was verified by probing for IGF1Rβ (membrane protein) and RhoGDIα (cytosolic protein). 2.8 Focal adhesion enrichment assay The preparation of focal adhesion-enriched (insoluble) and detergent-soluble (soluble) cell fractions was carried out as described (Kaplan KB et al. 1994; Sakai et al. 1994). The cells were placed on fibronectin-coated dishes for 15 min as described above, and then the dishes were chilled on ice. The cells were washed twice with ice-cold PBS and lysed with cytoskeleton stabilizing buffer (CSB) (0.3 M sucrose, 0.5% Triton X-100, 10 mM Pipes pH 6.8, 100 mM KCl, 1 mM CaCl2, 2.5 mM MgCl2, 1 mM Na3VO4, and protease inhibitors; 1 mM PMSF, 10 μg/ml leupeptin, 10 μg/ml aprotinin) for 5 min, and collected into tubes. This 57 fraction is referred to as the detergent-soluble or soluble fraction. The dishes were rinsed briefly (<5s) twice with CSB, and RIPA buffer (50 mM Hepes pH 7.4, 1% sodium deoxycholate, 1% Triton X-100, 0.1% SDS, 150 mM NaCl, 1 mM EDTA, 1 mM Na3VO4 and protease inhibitors as described above) was added to the dishes and they were kept for 5 min on ice. The dishes were scraped and the resulting solution was centrifuged at 16,000×g at 4ºC for 15 min to clear cell debris. The supernatant is referred to as the insoluble or focal adhesion-enriched fraction. Following Western blotting of the fractions and densitometric quantification, the % insoluble protein representing focal adhesion-localized protein was calculated as . 2.9 Pulldown assays Rac1 and Cdc42 activities were measured by GST-PBD (PAK-binding domain) pull-down assays performed with 100 μg cell lysate using a Rac1 Activation Kit from Stressgen. To assess BCAR3-SH2 domain interactions, cell lysates were mixed with recombinant GST- BCAR3-SH2 (Signosis, Inc.) and GST-Bind Resin (Novagen) for 18 h at 4 o C. The resin was collected, washed three times with cell lysis buffer, and resolved by SDS-PAGE and analyzed by immunoblotting. 2.10 Far-Western blotting Anti-VSVG-PTPα (WT and Y789F) immunoprecipitates were resolved by SDS-PAGE and transferred to a PVDF membrane. The membrane was probed with GST-BCAR3-SH2 domain fusion protein at 4°C. After overnight incubation, the membrane was washed three 58 times with PBST (PBS containing 0.1% Tween-20), followed by the addition of HRP- conjugated anti-GST antibody at room temperature for 2 h. After washing three times in PBST, signal was detected using ECL Plus™ (Amersham). 2.11 Cell migration assays Wound healing experiments were carried using the BioFlux200 system. Adherent cells were detached by trypsin and re-plated as a monolayer of cells into each FN-coated channel of a BioFlux 24-well plate (Catalog#910-0050, Fluxion Biosciences) at 2 Dyn/cm 2 using the BioFlux200 system. After 24 hours incubation at 37°C, cells were removed from half of the channel (~200μm) by 0.25% trypsin-EDTA at 10 Dyn/cm2 using the BioFlux200 system and the plate was incubated at 37ᵒC for another 24 hours to allow cells move to the wound. Cells were stained with 1 nM Calcein AM (Sigma) in DMEM at 37ᵒC for 15 min and imaged using the Olympus IX81system equipped with a CoolSnap HQ2 CCD camera. 2.12 Matrigel transwell invasion assays Cell invasion experiments were carried in BD BioCoat™ BD Matrigel™ Invasion Chambers (Cat#354480). The lower chambers were pre-incubated with DMEM containing 10% serum for 30min. Adherent cells were detached from culture dishes by trypsin and 6×10 4 cells in DMEM were seeded into the top chambers. After incubating at 37°C for 24 hours, the Matrigel and cells on the top of the membrane in each top chamber were removed using cotton swabs. The cells on the bottom of the membrane were stained with 1 nM Calcein AM (Sigma) at 37°C for 15min, and imaged using an Olympus FV10i microscope. 59 The numbers of invaded cells in each chamber were counted by ImageJ software and confirmed by manual counting. 2.13 Statistical analyses Densitometric quantification of immunoblots from at least three independent experiments was statistically analyzed using the Student’s t-test. 60 Chapter 3: PTPα Tyr789 Phosphorylation Regulates Integrin- induced Cell Spreading and Migration through a Cas/Crk-Rac/Cdc42 Signaling Axis 3.1 Rationale PTPα Tyr789 is phosphorylated in cells growing on culture plates (den Hertog et al. 1994), and this is reversibly regulated by integrin signaling (Chen et al. 2006). The regulation and function of PTPα in integrin signaling have been a main research focus in our laboratory. As reported, the integrin-stimulated phosphorylation of PTPα Tyr789 is catalyzed by Src family kinases (SFKs) and focal adhesion kinase (FAK), and requires an intact actin cytoskeleton (Chen et al. 2006). Functional characterization of this phosphorylation event showed that it was not required for PTPα to dephosphorylate Src Tyr527 and activate Src, since the defective integrin-induced Src Tyr527 dephosphorylation and Tyr416 auto- phosphorylation (Src activation events) in PTPα-/- cells were fully rescued by the re- expression of mutant Y789F PTPα. Likewise, integrin-induced Src-dependent tyrosine phosphorylation of FAK and paxillin were not altered by PTPα Y789F mutation. Surprisingly, despite Src and FAK activation proceeding normally in PTPα-/- cells re- expressing Y789F mutant PTPα, immunofluorescent staining for F-actin and vinculin still showed reduced integrin-induced stress fiber assembly and focal adhesion formation when compared to wildtype MEFs (PTPα+/+) or PTPα-/- cells re-expressing wildtype (WT) PTPα (Chen et al. 2006). Similarly, re-expression of PTPα Y789F mutant was unable to completely rescue the transient delay in cell spreading and the reduced rate of cell migration observed in PTPα-/- MEFs (Su et al. 1999; Zeng et al. 2003), which implicates a role for PTPα Tyr789 61 phosphorylation in these integrin regulated events. However, the signaling events that are transduced by PTPα phospho-Tyr789 are still unclear. In the present study, I examined the signaling pathways that are crucial for focal adhesion formation, stress fiber formation, and cell spreading in order to identify the molecules and signaling pathways with activation dependent on the Tyr789 phosphorylation of PTPα. To this purpose, I investigated and identified integrin signaling events that are defective in PTPα-/- MEFs, and determined which of these could be rescued by adenovirus-mediated expression of wild-type (WT) PTPα or an unphosphorylatable mutant (Y789F) form of PTPα. In this way, I aimed to pinpoint the PTPα Tyr789 phosphorylation dependent events. 3.2 Rac1/Cdc42 signaling is regulated by PTPα Tyr789 Cell migration is a complex multi-step process that incorporates a) cell spreading, b) focal adhesion formation, c) formation of stress fibers and contraction, d) cell-body displacement, and e) detachment of the cell rear from the ECM. These events require the coordination of multiple signal transduction pathways and numerous signaling molecules (Ridley et al. 2003). Cell adhesion and spreading are initial steps in cell migration, and are driven by the formation of lamellipodia and filopodia at the cell periphery that promote the outwards extension of the cell membrane towards the substratum. This process can be divided into two steps, which include the formation of cell protrusions (filopodia and lamellipodia) and focal complexes that anchor the leading edge of the cell to the ECM. Both steps are regulated by Rho family GTPases, including Rac1 and Cdc42. Rac1 is the key mediator of the formation of lamellipodia (Ridley and Hall 1992). Inhibiting Rac1 action by introducing dominant- negative Rac1 S17N can impair lamellipodia formation and cell spreading on FN (Price et al. 62 1998). Similarly, Cdc42 has been shown to induce filopodia in response to integrin stimulation (Kozma et al. 1995; Nobes and Hall 1995). Two major defects observed in PTPα- /- MEFs and in PTPα-/- MEFs re-expressing mutant PTPα Y789F are reduced focal adhesion formation and impaired cell spreading on FN (Zeng et al. 2003; Chen et al. 2006). Thus, I hypothesized that PTPα Tyr789 phosphorylation may regulate the activation of Rac1 and/or Cdc42 to control integrin-induced cytoskeleton re-arrangement. I first examined the activation of Rac1 and Cdc42 upon FN stimulation of MEFs. 3.2.1 Roles of PTPα and PTPα Tyr789 in Rac1/Cdc42 activation Like all GTPases, Rac1 and Cdc42 have two states: the GTP-bound active form and the GDP-bound inactive form. The GTPases hydrolyze GTP to GDP with the help of GTPase activating proteins (GAPs), while GDP to GTP exchange is catalyzed by guanine nucleotide exchange factors (GEFs) (Hall 1998). Other regulators of small GTPases are guanosine nucleotide dissociation inhibitors (GDIs), which function to sequester GTPases in the cytoplasm away from their sites of action at cell membranes (Burridge and Wennerberg 2004). It has been reported that PTPα is required for integrin-induced Rac1 activation and Rac1 re-localization to plasma membranes (Herrera Abreu et al. 2008). To confirm the role of PTPα in Rac1 and Cdc42 membrane translocation, PTPα+/+ and PTPα-/- MEFs were trypsinized, maintained in suspension for 1 hour, and re-plated on FN coated dishes for 15 minutes. Membrane and cytosolic fractions were then prepared as described (refer to Materials and Methods Section 2.7), resolved by SDS-PAGE electrophoresis, and probed for a membrane binding protein ‘IGFR1β’ and a cytosolic protein ‘RhoGDIα’ to validate the 63 fractionation as well as the loading amounts. This clearly showed that there was almost no contamination between the two fractions, since virtually all IGFR1β was present in the membrane fraction and RhoGDIα was almost exclusively present in the cytosolic fraction (Fig. 3.1A, bottom panels). Furthermore, equal sample loading was confirmed for fractions from the two cell types (RhoGDIα: 0.98±0.05 in PTPα-/- MEFs compared to that in wild-type MEFs; IGFR1β: 1.09±0.08 in PTPα-/- MEFs compared to that in wild-type MEFs) (Fig. 3.1B). Consistent with Abreu et al. (Herrera Abreu et al. 2008), I found that after stimulating integrin signaling with FN for 15 min there was less Rac1 (54.2±17.4%) and Cdc42 (39.3±16.2%) in the membrane fraction of integrin-stimulated PTPα-/- MEFs compared to wild-type MEFs (Fig. 3.1A and C). No significant differences in the cytosolic localization of Rac1 and Cdc42 were detected between wild-type and PTPα-null cells (Fig. 3.1C, bottom graphs). This confirmed that PTPα is required for the optimal membrane localization of Rac1 and Cdc42 that promotes their activation. To confirm that Rac1 and Cdc42 signaling was accordingly affected by the impaired membrane localization of these GTPases, the PTPα- dependent membrane localization of the downstream Rac1/Cdc42 effector, p21-activated kinase (PAK), was examined. PAK re-localization is likewise required for its signaling function (Lu et al. 1997). Indeed, the PTPα-/- MEFs contained less membrane-associated PAK (55.2±23.5% compared to wild-type MEFs) (Fig. 3.1A and C), indicating that PTPα regulates formation of the Rac1/Cdc42-PAK signaling complex at the membrane. The above membrane translocation assay reveals a critical event mediated by PTPα in activating Rac1 and Cdc42 signaling. To investigate Rac1 and Cdc42 activation status directly, I employed an in vitro method of determining the cellular activation status of Rho GTPases (Ren et al. 1999) that has been extensively used to determine the activities of small 64 GTPases for more than a decade. In brief, cell lysates are subjected to pulldown assays involving glutathione beads conjugated to a GST-fusion protein comprising the GTPase binding domain of the relevant Rho GTPase effector protein (PAK binding domain for Rac1 and Cdc42), which specifically recognizes the active GTP-bound form of the Rho GTPase. Therefore, the GTP-bound Rho GTPase is precipitated by the GST-fusion protein-coupled glutathione beads in the pull-down assay. The amount of precipitated Rho GTPase, representing the active form of the GTPase (GTP-bound form), is visualized by immunoblotting to provide a quantitative indication of the cellular activation status of the Rho GTPase of interest. PTPα+/+ and PTPα-/- cells were kept in suspension for 1 hour, and then plated on FN coated plates for 15 minutes. In PBD (PAK binding domain, that recognizes GTP-bound Rac1 and Cdc42) pulldown assays, reduced amounts of GTP-bound Rac1 and Cdc42 were isolated from FN-stimulated PTPα-/- cells compared to the amounts of active Rac1 and Cdc42 detected in PTPα+/+ MEFs (Fig. 3.2A). Since similar amounts of Rac1 and Cdc42 protein were present in PTPα+/+ and PTPα-/- cell lysates (Fig. 3.2A), the differences observed in the amounts of active Rac1 and Cdc42 activity were not due to down-regulated protein expression or to unequal amounts of lysate used for pulldown. To determine the role of PTPα Tyr789 phosphorylation in Rac1 and Cdc42 activation, wildtype (WT) and mutant Y789F PTPα were re-introduced into PTPα-/- cells via adenovirus infection. Expression of exogenous PTPα at physiological levels was confirmed by immunoblotting the cell lysates for PTPα (Fig. 3.2A, bottom panels). As expected, upon integrin stimulation WT PTPα rescued the defective activation of Rac1 and Cdc42 to a level comparable to that of PTPα+/+ cells (Fig 3.2A). However, re-introduction of unphosphorylatable Y789F PTPα into PTPα-/- 65 cells did not rescue the defective activation of Rac1 and Cdc42. This indicated that PTPα Tyr789 phosphorylation regulates the GTP-loading and activation of Rac1 and Cdc42. 3.2.2 Roles of PTPα and PTPα Tyr789 in PAK activation PAK, a protein serine/threonine kinase, is one of the main downstream effectors of Rac1/Cdc42. Microinjection of constitutively active mutant PAK into cells can induce the formation of lamellipodia and membrane ruffles (Sells et al. 1997), and the introduction of active PAK can partially rescue the defective cytoskeleton re-arrangement observed in Rac1 knockout MEFs (Guo et al. 2006). Upon FN stimulation, active Rac1/Cdc42 can bind to PAK, disrupting the inhibitory interaction between the PAK kinase domain and its auto- inhibitory domain. This conformational change leads to autophosphorylation at Ser144 and subsequent kinase activation (Manser et al. 1994; Martin et al. 1995; Knaus et al. 1998; Chong et al. 2001). PAK activation status can thus be assayed by immunoblotting for phospho-Ser144 PAK. To confirm the functional consequence of the defective Rac1/Cdc42 activation in Y789F PTPα expressing cells, I examined the role of phosphoTyr789 PTPα in integrin-induced PAK activation. As shown in Fig. 3.2B, immunoblotting with PAK antibody revealed the presence of two PAK isoforms, the 68 kD PAK1 and the 62 kD PAK2. Immunoblotting with the phosphosite- specific PAK Ser144 antibody detected mainly the 68 kD PAK1 with very minimal signal detected for the 62 kD PAK2 (Fig. 3.2B). This is due to either a lower sensitivity of the phosphosite-specific antibody towards the PAK2 isoform or to much lower PAK2 phosphorylation compared to PAK1 upon integrin activation. The quantitative analysis thus only included the PAK1 isoform. PTPα+/+ and PTPα-/- fibroblasts growing on plastic dishes 66 were trypsinized and maintained in suspension for 1 hour, followed by plating on FN to stimulate integrin signaling for 15 minutes. As shown in Fig. 3.2B, immuno-blotting with PAK phospho-Ser144 antibody revealed almost undetectable PAK Ser144 phosphorylation in suspended PTPα+/+ and PTPα-/- cells. After plating the cells on FN for 15 min, PAK Ser144 phosphorylation was greatly induced. Although both PTPα+/+ and PTPα-/- cells exhibited a similar trend of increased PAK phosphorylation during integrin stimulation, densitometric analysis revealed a significant ~50% reduction in PAK Ser144 phosphorylation level in PTPα-/- cells at 15 min on FN compared to that in FN-stimulated PTPα+/+ cells (106±23% for PTPα+/+ cells and 59±13% for PTPα-/- cells) (Fig. 3.2C). These results indicated that integrin-induced PAK Ser144 phosphorylation is impaired in PTPα-/- cells. To investigate the role of PTPα Tyr789 in integrin-induced PAK activation, WT or mutant Y789F PTPα were re-introduced into PTPα-/- MEFs using adenovirus. Integrin-induced PAK Ser144 phosphorylation in PTPα-/- MEFs re-expressing WT PTPα was rescued to ~90%, but in MEFs re-expressing Y789F PTPα the phosphorylation of PAK was only restored to ~65% (91±1% for PTPα-/-MEFs re-expressing WT PTPα; 66±15% for PTPα-/- cells re-expressing Y789F PTPα) (Fig. 3.2B and C). These findings revealed that PTPα Tyr789 indeed regulates the full activation of PAK in integrin signaling. In summary, the above results show that PTPα Tyr789 regulates the integrin-induced activation of the small GTPases Rac1 and Cdc42 and one of their downstream effectors, PAK. To determine if Rac1 and Cdc42 activation was directly regulated by PTPα Tyr789, or if the regulatory action of PTPα Tyr789 was exerted on an activator of the Rac1/Cdc42 signaling, I next investigated upstream signaling events. 67 3.3 PTPα Tyr789 is necessary for efficient integrin-induced Cas tyrosine phosphorylation and Cas-Crk association After cell attachment to the ECM activates integrin heterodimers, one of the earliest signaling events is FAK activation (Sieg et al. 1999). Early low level auto-phosphorylation of FAK at Tyr397 forms a docking site for Src, resulting in the full activation of the FAK-Src complex. Previously work in our lab demonstrated that mutant Y789F PTPα catalyzes integrin-induced Src and FAK activation as effectively as WT PTPα (Chen et al. 2006), and I confirmed these results in FN-stimulated MEFs (Figs. 3.3 and 3.4). This active FAK-Src complex phosphorylates two important scaffolding proteins, p130Cas and paxillin. Cas is an key regulator of cell migration (Cary et al. 1998; Klemke et al. 1998), and is ubiquitously expressed in all vertebrates (Sakai et al. 1994). The Cas knockout mouse is embryonic lethal and Cas -/- MEFs show disorganized actin stress fibers and defective cell migration (Honda et al. 1998). Cas tyrosine phosphorylation, especially at multiple sites in its substrate domain (SD), generates binding sites for adapter proteins such as Crk and Nck. In turn Crk can recruit a complex of Dock180/ELMO to activate Rac. Paxillin is a 68 kDa multi-domain focal adhesion protein that functions as a molecular scaffold to facilitate integrin signaling (Brown and Turner 2004; Playford and Schaller 2004). It is essential for embryonic development and plays an important role in the regulation of cell attachment, spreading and motility (Hagel et al. 2002; Brown and Turner 2004). The amino terminal region of paxillin contains five leucine-rich domains (LD), (Brown and Cooper 1996; Brown et al. 1998). The LD4 motif of paxillin, via the paxillin kinase linker (PKL), binds a complex that includes the Cdc42/Rac guanine nucleotide exchange factor PIX, PAK, and the SH2- SH3 adapter protein Nck, to activate Rac1/Cdc42 and PAK signaling (Turner et al. 1999; 68 Brown and Turner 2004). Previous data showed that FN-stimulated paxillin phosphorylation is independent of PTPα Tyr789 (Chen et al. 2006), thus there remained the possibility that Cas signaling may be regulated by phosphoTyr789-PTPα. I therefore investigated if PTPα Tyr789 was required for the tyrosine phosphorylation of Cas that is required to promote Cas signaling. The integrin-induced tyrosine phosphorylation of Cas is impaired in MEFs lacking PTPα (Su et al. 1999; Kostic et al. 2007), as I confirmed (46±14% in PTPα-/- MEFs compared to wild-type MEFs) (Fig 3.5, left panels). This has been attributed to the reduced activity of the PTPα-activated Src family kinases (Src and Fyn) that phosphorylate Cas. Expressing WT PTPα in PTPα-/- MEFs restored the defective Cas tyrosine phosphorylation when suspended cells were placed on dishes coated with the integrin ligand fibronectin (FN) (Fig. 3.5, right panels). However, a considerably reduced tyrosine phosphorylation of Cas in PTPα-/- MEFs was detected in PTPα-/- MEFs expressing Y789F PTPα (44±3% in Y789F PTPα cells compared to WT PTPα expressing MEFs) (Fig. 3.5, right panels), indicating a distinct Tyr789-dependent action of PTPα. Integrin-stimulated tyrosine phosphorylation of Cas occurs at multiple sites, particularly within a central region known as the substrate domain (SD). The SD is characterized by fifteen YxxP motifs, ten of which can be efficiently phosphorylated by Src (Songyang et al. 1993; Sakai et al. 1994). FAK catalytic activity is not required for SD phosphorylation, but FAK interacts with the Src SH2 domain to recruit Src to phosphorylate the SD of FAK- associated p130Cas (Ruest et al. 2001). The phosphorylation of the Cas SD has been detected predominantly at FAs and at the cell periphery in nascent integrin adhesion sites (Fonseca et al. 2004; Ballestrem et al. 2006). After activation by tyrosine phosphorylation, the Cas SD is 69 able to recruit downstream effectors, notably the SH2/SH3 adaptors Crk and Nck that act to promote protrusion of the leading edge plasma membrane by activation of Rac1/Cdc42-PAK signaling (Bouton et al. 2001; Chodniewicz and Klemke 2004; Defilippi et al. 2006; Rivera et al. 2006). To confirm that the reduced overall Cas phosphorylation described above included tyrosine sites in the SD, I determined whether the phosphorylation of Cas at a specific tyrosine residue within the SD, Tyr410, is regulated by PTPα Tyr789. PTPα+/+ and PTPα-/- cells were kept in suspension for 1 hour, and plated on FN coated plates for 15 minutes. Cell lysates were probed for phosphoTyr410 Cas, Cas, and PTPα. I observed decreased phosphorylation of Cas Tyr410 in PTPα-/- cells (53±16%) upon FN stimulation, compared to PTPα+/+ MEFs (Fig. 3.6, left panels). Wildtype (WT) and Y789F PTPα were re-introduced into PTPα-/- cells by adenovirus infection and the expression of the exogenous PTPα to an equivalent physiological level was confirmed by immunoblotting the cell lysate for PTPα (Fig. 3.6A, bottom panels). As expected, WT PTPα rescued the defective FN-induced phosphorylation of Cas Tyr410, but this remained defective in cells expressing Y789F PTPα (60±21%) (Fig. 3.6, right panels). To confirm the functional consequence of the defective Cas phosphorylation in Y789F PTPα expressing cells, I also examined the roles of PTPα and PTPα Tyr789 in regulating Cas and Crk association. In accord with the impaired tyrosine phosphorylation of Cas in PTPα-/- and Y789F PTPα-expressing MEFs, the association of Cas and Crk was reduced in these cells, as detected by reciprocal co-immunoprecipitation and probing of Cas and Crk (Fig. 3.7). 70 Together, these results indicated that PTPα Tyr789 regulates integrin-mediated Cas SD phosphorylation and Crk recruitment, thus promoting Rac1/Cdc42 signaling and eventually cell migration. 3.4 Cas and Src association is impaired in cells expressing mutant Y789F PTPα There are two known mechanisms of Cas SD tyrosine phosphorylation: 1) the Cas SH3 domain binds to FAK via FAK poly-proline motifs, FAK phosphoTyr397 forms a docking site for the SH2 domain of Src, and Src phosphorylates Cas in the SD using FAK as a platform; and 2) the Src SH3 domain binds directly to Cas at the Src binding domain and Src phosphorylates Cas in the SD. It is Src that functions as a major kinase to catalyze the phosphorylation of Cas and FAK kinase activity is not required (Ruest et al. 2001). Since Y789F PTPα is as efficient as WT PTPα in activating Src in integrin-stimulated MEFs (Fig. 3.3) (Chen et al. 2006), the defective Cas phosphorylation in Y789F PTPα expressing cells is clearly not due to defective kinase activity of Src. I therefore investigated whether the reduced Cas tyrosine phosphorylation in Y789F PTPα expressing cells was due to the impaired association of Cas and Src. Cas immunoprecipitates from wild-type and PTPα-/- MEFs, and from PTPα-/- MEFs re-expressing WT and Y789F PTPα, were probed for associated Src. Integrin stimulation induced the association of Src with Cas in wild-type MEFs but not in PTPα-/- MEFs (36±10% in PTPα-/- MEFs compared to wild-type) (Fig. 3.8, left panels). Likewise increased Src-Cas association was induced by integrin stimulation in PTPα-/- cells expressing WT but not mutant Y789F PTPα (67±1% in PTPα-/- MEFs expressing Y789F PTPα compared to WT PTPα expressing MEFs) (Fig. 3.8, right panels). 71 These results indicate that Tyr789 of PTPα is necessary for the Cas-Src association that enables Src to phosphorylate Cas and promote downstream signaling. This is the most upstream PTPα Tyr789-dependent signaling event that was identified post-FAK-Src complex activation in integrin signaling. Interestingly, my findings (Fig.3.8B, left graph) suggest that there is a basal level (36%) of Cas-Src association in the absence of PTPα. I propose that the unphosphorylatable Y789F PTPα can rescue about another one-third of Cas-Src association (Fig. 3.8B, right graph) due to restored phosphatase activity of mutant Y789F PTPα promoting Src activation, with the remaining approximately one-third of Cas-Src association being dependent on phospho-Tyr789 of PTPα. This suggests that PTPα Tyr789 phosphorylation is equally important as its phosphatase activity in regulating cell migration. 3.5 PTPα-Tyr789 is required for integrin-induced focal adhesion localization of PTPα and Cas but not Src Integrin engagement induces the formation of intracellular multi-protein complexes at the integrin-substrate adhesion sites, known as focal adhesions (FAs). The components in these huge complexes have so far been identified to include one hundred and eighty proteins (including PTPα, Src, and Cas), four lipids, and one ion (calcium) (Zaidel-Bar et al. 2007; Zaidel-Bar and Geiger). However, the detailed mechanisms by which this complex forms upon integrin activation are still unclear. It has been shown that PTPα, Src, Cas can localize to FAs upon integrin stimulation (Petch et al. ; Vuori and Ruoslahti 1995; Lammers et al. 2000; Donato et al. 2010), and Cas moves to nascent FAs where it is phosphorylated by Src (Fonseca et al. 2004; Ballestrem et al. 2006). The defective Cas-Src association in mutant Y789F PTPα-expressing cells suggests that in the absence of PTPα-Tyr789 phosphorylation, 72 Cas and Src might not co-localize properly in newly forming FAs. I therefore investigated whether Cas recruitment to FAs is dependent on PTPα, and on PTPα-Tyr789, using a biochemical FA enrichment assay. Immunofluorescent visualization of PTPα indicates that PTPα-Tyr789 directs the localization of PTPα to adhesion sites or FAs (Lammers et al. 2000), raising the possibility that Cas or Src recruitment to FAs might be regulated by phospho-Tyr789 PTPα-dependent recruitment to these structures. I determined the integrin-induced redistribution of endogenous PTPα and expressed WT and Y789F PTPα by fractionating suspended or integrin-stimulated cells into soluble and FA-enriched fractions (Kaplan et al. 1994; Sakai et al. 1994) (refer to Materials and Methods Section 2.8). In wild-type MEFs, most of the endogenous PTPα was located in the soluble fraction (88±6.3% of total PTPα) of suspended cells that lack integrin-mediated substrate attachment and signaling. Plating these cells on FN induced PTPα Tyr789 phosphorylation and the re-localization of a majority of PTPα (77±1% of total PTPα) to the insoluble FA-enriched fraction (Fig. 3.9, left panels, first four lanes). In PTPα-/- cells expressing introduced WT PTPα, a similar integrin-stimulated phosphorylation and re-localization of WT PTPα was detected (74±8% of total PTPα in insoluble fraction) (Fig. 3.9, right panels, first four lanes). However, the integrin-induced re-localization of expressed mutant Y789F PTPα to the FA-enriched fraction was severely impaired (only 27±6% of total PTPα in insoluble fraction) (Fig. 3.9, right panel, last four lanes). Examination of the localization of Cas, Crk, and Src in the four cell types above revealed that in wild-type MEFs, populations of all these proteins moved to FA-enriched fractions upon integrin stimulation. Strikingly, the re-localization of Cas and Crk, but not Src, was PTPα-dependent, as this was significantly reduced in PTPα-/- MEFs (Fig. 3.10A left panels, 73 first four lanes). Indeed only 8±3% of total Cas was found in the insoluble fraction of PTPα-/- MEFs compared to 21±3% in PTPα+/+ MEFs, whereas 83±1% of total Src was present in insoluble fraction of PTPα-/- MEFs that was equivalent to the 83±7% found in this fraction of PTPα+/+ MEFs) (Fig. 3.10B and C, left graphs). Expression of WT PTPα in the PTPα-null cells restored Cas and Crk re-localization to the FA-enriched fraction, whereas expressed Y789F PTPα was much less effective (Cas: 17±10% of total Cas in the insoluble fraction of PTPα-/- MEFs compared to 37±6% in PTPα+/+ MEFs; Src: 88±2% of total PTPα in insoluble fraction in PTPα-/- MEFs compared to 86±5% in PTPα+/+ MEFs) (Fig. 3.10A-C, right panels and graphs). These results indicate that integrin-stimulated PTPα phosphorylation at Tyr789 promotes the recruitment of PTPα and Cas to FAs, while the movement of Src to these structures is PTPα-independent. 3.6 Discussion Integrin-induced phosphorylation of PTPα at Tyr789 is required for cell spreading and cell migration (Chen et al. 2006). Through reconstituting WT PTPα or unphosphorylatable mutant Y789F PTPα expression in PTPα-/- MEFs and evaluating their abilities to rescue integrin signaling defects observed in PTPα -/- cells, I identified the tyrosine phosphorylation of Cas SD as the most integrin-proximal signaling event that is dependent on Tyr789 of PTPα. I also discovered that impaired Cas SD phosphorylation is caused by the defective association of Src and Cas in Y789F PTPα-expressing MEFs. Biochemical localization studies demonstrated that only WT but not Y789F PTPα moved into FA-enriched fractions upon FN stimulation, consistent with the previously reported Tyr789-dependent immunofluorescent re-localization of PTPα at FAs (Lammers et al. 2000). Interestingly, the 74 reduced presence of Y789F PTPα in FAs was also associated with the impaired FA localization of Cas, but the localization of Src to FAs is independent of PTPα and its phosphorylation. This indicated that the reduced association of Src and Cas in Y789F PTPα- expressing PTPα-/- MEFs is due to defective Cas FA translocation in these cells. Together, these results indicate that PTPα phosphoTyr789 has a key role in recruiting not only PTPα, but also Cas, to nascent FA complexes where Cas is in proximity to Src, thus promoting Src phosphorylation of the Cas SD to transmit the signal to downstream effectors. My findings demonstrate a new level of functional interdependency between PTPα and Src in integrin signaling as not only does PTPα dephosphorylate and activate Src, but Src- mediated phosphorylation of PTPα (Chen et al. 2006) promotes the accessibility of Cas as a substrate for Src. The dephosphorylation of Src at Tyr527 and its activation are impaired in PTPα-/- MEFs, and in MEFs expressing mutant catalytically inactive PTPα Src is not activated and PTPα Tyr789 phosphorylation is defective (Ponniah et al. 1999; Su et al. 1999; Chen et al. 2006), indicative of PTPα-Src-PTPα signaling. Furthermore, the expression of Y789F PTPα in PTPα-/- MEFs rescues the Src activation (Chen et al. 2006) (Fig. 3.3), but still cannot restore the downstream Cas signaling (Figs. 3.1, 3.2, 3.5-3.8) and defective spreading and migration phenotypes (Chen et al. 2006), which suggesting the Src-catalyzed phosphorylation of PTPα Tyr789 is one of the key events after Src activation in integrin- stimulated cell migration. Upon integrin activation, wildtype PTPα and Cas co-localize to FA in a Tyr789-dependent manner, raising the possibility that PTPα phospho-Tyr789 associates with Cas to form a protein complex that translocates to newly formed FAs. However, the mechanism by which these proteins might interact is unknown. 75 PTPα has a very short extracellular domain, a transmembrane domain, two catalytic domains (D1 and D2), and a short C-terminal tail containing Tyr789. Structurally PTPα does not have any obvious protein-protein interacting domain, such as an SH3 domain-binding poly-proline region, which could associate with Cas. In fact, Cas has been reported by one group as a substrate that can be dephosphorylated by PTPα, but Cas can only be co- precipitated with the phosphatase-dead PTPα D1 domain (Buist et al. 2000). This suggests that wildtype PTPα likely does not interact with Cas in a stable manner, but rather form a transient enzyme-substrate complex. There are two types of domains that can interact with phosphotyrosine motifs in proteins: the PTB domain and the SH2 domain. So far, only two SH2 domain-containing proteins, Src and Grb2, have been reported to interact with PTPα phospho-Tyr789 in integrin signaling (den Hertog et al. 1994; Zheng et al. 2000). My results clearly showed a reduced Cas-Src interaction and defective Cas FA co-localization in mutant Y789F PTPα expressing MEFs upon integrin-stimulation, suggesting Src is not the molecular linker between PTPα and Cas. So far, an association between Grb2 and Cas has not been reported to occur during integrin signaling. When interacting with PTPα phosphoTyr789, Grb2 is not bound to the Ras/MAPK activator, SOS and inhibits the phosphatase activity of PTPα (den Hertog et al. 1996). This negative role of Grb2 indicates that it is not likely to be involved in mediating the direct interaction of PTPα with Cas to promote cell migration. The exact function of PTPα phosphoTyr789-bound Grb2 still requires further investigation. In contrast, Cas, as a scaffolding protein, has many binding partners that have been identified in FAs, such as FAK, PTP-1B, PTP-PEST, C3G, and Dock180 that can bind to Cas SH3 domain (Burnham et al. 1996; Hsia et al. 2003); Crk and Nck that interact with phosphotyrosine sites in the SD domain of Cas (Burnham et al. 1996; Schlaepfer et al. 1997); 76 Src that directly binds to the Src binding domain in Cas (Sakai et al. 1994; Nakamoto et al. 1996); and PI3K, Ajuba and BCAR3 that interact with the Cas C-terminal domain (Gotoh et al. 2000; Li et al. 2000; Pratt et al. 2005). However, Cas does not possess SH2 or PTB domains that could directly interact with PTPα phosphoTyr789, so I postulated that an intermediary protein or protein complex connects PTPα phosphoTyr789 and Cas to facilitate their FA localization. In the next chapter, I describe approaches to identify the protein that involves in PTPα Tyr789-mediated Cas FA localization. AFigure 3.1 Lack of PTPα Tyr789 impairs integrin-induced membrane association of Rac, Cdc42 and PAK. Wild-type (+/+) and PTPα-/- (-/-) MEFs were placed on FN-coated dishes for 15 min. (A) Cell membrane and cytosolic fractions were prepared and probed for the indicated proteins. The distributions of the membrane-associated protein IGF1Rβ and the cytosolic protein RhoGDIα were probed to validate the fractionation. Results from 3 experiments were quantified by densitometry, and (B) the relative amounts of IGF1Rβ in the membrane fraction and of RhoGDIα in the cytosol fraction of wild- type (+/+) and PTPα-/- (-/-) MEFs are shown in the graphs as mean ± S.D. (C) The relative amounts of Rac, Cdc42, and PAK in the both membrane and cytosolic fractions, per unit IGF1Rβ or RhoGDIα, respectively, are shown in the graphs as mean ± S.D. Asterisks indicate significant differences (*, p=0.01; **, p=0.003; ***, p=0.03). IGF1Rβ RhoGDIα Rac1 Cdc42 PAK PTPα: M C +/+ M C ‐/‐ +/+ ‐/‐M em br an e p ro te in am ou nt / I GF 1R β * 1.0 0.2 0.4 0.6 0.8 ** Rac1 Cdc42 PAK +/+ ‐/‐ +/+ ‐/‐ 0.2 0.4 0.6 0.8 1.0 1.2 1.4 Cy to so lic pr ot ei n am ou nt / R ho GD Iα +/+ ‐/‐ +/+ ‐/‐ +/+ ‐/‐ *** PTPα: PTPα: IGF1Rβ +/+ ‐/‐ 1.0 0.8 0.6 0.4 0.2 1.2 M em br an e a m ou nt PTPα: +/+ ‐/‐ RhoGDIα 1.0 0.8 0.6 0.4 0.2 1.2 Cy to so lic am ou nt B C 77 BFigure 3.2 Lack of PTPα Tyr789 impairs integrin-induced activation of Rac, Cdc42 and PAK. Wild-type (+/+) and PTPα-/- (-/-) MEFs, or PTPα-/- MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα were cultured 24h, starved with 0.5% FBS- DMEM for 24h, placed in suspension for 1 h, and then placed on FN-coated dishes for 15 min. Cell lysates were used in GST-PBD pull-down assays. (A) The pull-downs were probed for Rac1 or Cdc42 to detect active GTP-bound proteins (indicated as Rac1-GTP and Cdc42-GTP). Lysates were also probed for Rac1, Cdc42, phosphoTyr789-PTPα (pY789) and PTPα. The above four groups of MEFs were grown as adherent (Ad) cultures on plastic dishes and then placed in suspension for 1h (0) before plating on FN-coated dishes for 15 min (15). (B) Lysates were probed for phosphoSer144-PAK (pS144) and PAK. (C) Results from 3 experiments were quantified by densitometry and the PAK Ser144 phosphorylation per unit PAK is shown in the graphs as mean ± S.D. Asterisks indicate significant differences (*p=4.1×10-3; **p=0.047). PTPα PTPα‐Y789 Cdc42‐GTP Cdc42 Rac1‐GTP Rac1 WT Y789F +/+PTPα: AdV: ‐/‐ ‐/‐ ‐/‐ C pS144 PAK Ad 150 Ad 150 +/+ ‐/‐ Ad 150 Ad 150 WT Y789F ‐/‐ ‐/‐ AdV: 100 80 60 40 20 WT Y789F ‐/‐ ‐/‐ AdV: 100 80 60 40 20 120 +/+ ‐/‐PTPα: 140 PA K p S1 44 /P AK * ** 120 A 78 CFigure 3.3 Integrin-induced Src activation is not dependent on PTPα Tyr789 phosphorylation. Wild-type (+/+) and PTPα-/- (-/-) MEFs, or PTPα-/- MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα were cultured 24h, starved with 0.5% FBS- DMEM for 24h, placed in suspension for 1 h, and then placed on FN-coated dishes for 15 min. Src immunoprecipitates were probed for (A) Src phosphoTyr416 and Src, or (B) Src phosphoTyr527 and Src. Results from 3 experiments were quantified by densitometry. (C) The Src Tyr416 phosphorylation per unit Src and (D) Src Tyr527 phosphorylation per unit Src are shown in the graphs as mean ± S.D. Asterisks indicate significant differences (*, p= 0.007; **, p= 6×10-4 ). A Src pY416 Src IP‐Src FN15 +/+ ‐/‐ Y789FWTAdV: PTPα: ‐/‐ ‐/‐ Src pY527 Src IP‐Src FN15 +/+ ‐/‐ Y789FWTAdV: PTPα: ‐/‐ ‐/‐ B D 20S rc pY 41 6 / Sr c (% ) WT Y789F ‐/‐ ‐/‐ AdV: +/+ ‐/‐PTPα: 60 100 140 180 * Sr c pY 52 7 / Sr c (% ) WT Y789F ‐/‐ ‐/‐ AdV: +/+ ‐/‐PTPα: 100 60 20 140 180 ** 79 Figure 3.4 Integrin-induced FAK activation is independent of PTPα Tyr789. Wild-type (+/+) and PTPα-/- (-/-) MEFs, or PTPα-/- MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα were cultured 24h, starved with 0.5% FBS- DMEM for 24h, placed in suspension for 1 h, and then placed on FN-coated dishes for 15 min. (A) Cell lysates were probed for FAK phosphoTyr397 and phosphoTyr576. Results from 3 experiments were quantified by densitometry, and (B) the FAK Tyr397 phosphorylation per unit FAK and (C) FAK Tyr576 phosphorylation per unit FAK are shown in the graphs as mean ± S.D. Asterisks indicate significant differences (*, p=2×10-5; **, p=4×10-5). A +/+ 0 15 ‐/‐ 0 15 0 15 0 15 FAK pY397 FAK FAK pY576 WT Y789FAdV: ‐/‐ ‐/‐PTPα: B C FA K p Y3 97 / F AK (% ) WT Y789F ‐/‐ ‐/‐ AdV: +/+ ‐/‐PTPα: 150 120 90 60 30 * 180 WT Y789F ‐/‐ ‐/‐ AdV: +/+ ‐/‐PTPα: 100 80 60 40 20FA K p Y5 76 / F AK (% ) **120 80 Figure 3.5 PTPα Tyr789 is required for integrin-induced Cas phosphorylation. Wild-type (+/+) and PTPα-/- (-/-) MEFs, or PTPα-/- MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα were cultured 24h, starved with 0.5% FBS- DMEM for 24h, placed in suspension for 1h (0), and then plated on FN-coated dishes for 15 min (15). (A) Cas immunoprecipitates were probed for phosphotyrosine (pTyr) and Cas. Lysates were probed for phosphoTyr789-PTPα and PTPα. (B) Results from 3 experiments were quantified by densitometry and the tyrosine phosphorylation per unit Cas is shown in the graphs as mean ± S.D. Asterisks indicate significant differences (*, p=5×10-3; **, p=1.1 ×10-5). Cas pTyr PTPα PTPα pY789 ‐/‐+/+ 0 15 ‐/‐ 0 15 WT 0 15 0 15 ‐/‐ AdV: Y789F * PTPα: ‐/‐ ‐/‐ ** 100 80 60 40 20pT yr /C as (% ) ‐/‐ WTAdV: Y789F +/+ 81 B A Figure 3.6 PTPα Tyr789 is required for integrin-induced Cas phosphorylation. Wild-type (+/+) and PTPα-/- (-/-) MEFs, or PTPα-/- MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα were cultured 24h, starved with 0.5% FBS- DMEM for 24h as adherent (Ad) cultures, placed in suspension for 1h (0), and then plated on FN-coated dishes for 15 min (15). Ad represents adherent cells grown on plastic dishes. (A) Lysates were probed for phosphoTyr410-Cas, Cas, and PTPα. (B) Results from 4 experiments were quantified by densitometry and the Tyr410 phosphorylation per unit Cas is shown in the graphs as mean ± S.D. Asterisks indicate significant differences (*, p=8×10-3 ; **, p=0.04). Cas +/+ Ad 0 15 ‐/‐ Cas pY410 ‐/‐ WTAdV: Y789F PTPα ‐/‐ Ad 0 15 Ad 0 15 Ad 0 15 * PTPα: ** 100 80 60 40 20 120 Ca sp Y4 10 /C as (% ) AdV: +/+ ‐/‐ ‐/‐ ‐/‐ WT Y789F 82 B A Figure 3.7 PTPα Tyr789 is required for integrin-induced Cas-Crk association. Wild-type (+/+) and PTPα-/- (-/-) MEFs, or PTPα-/- MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα were cultured 24h, starved with 0.5% FBS- DMEM for 24h, placed in suspension for 1h (0), and then plated on FN-coated dishes for 15 min (15). (A) Cas immunoprecipitates were probed for Crk and Cas. (B) Crk immunoprecipitates were probed for Cas and Crk. Results from 3 experiments were quantified by densitometry. (C) The Crk per unit Cas and (D) Cas per unit Crk are shown on the graphs as mean ± S.D. Asterisks indicate significant differences (*, p=7×10-6; **, p=4×10-4; ***, p=2×10-4; ****, p=4×10-4). * ** 1.0 0.8 0.6 0.4 0.2 Cr k/ Ca s PTPα: AdV: +/+ ‐/‐ ‐/‐ ‐/‐ WT Y789F *** **** 1.0 0.8 0.6 0.4 0.2 Ca s/ Cr k PTPα: AdV: +/+ ‐/‐ ‐/‐ ‐/‐ WT Y789F Cas Crk IP: Crk +/+ 0 15 ‐/‐ 0 15 0 15 0 15 ‐/‐ WT Y789FAdV: ‐/‐ Cas Crk IP: Cas +/+ 0 15 ‐/‐ 0 15 0 15 0 15 ‐/‐ WT Y789FAdV: ‐/‐ A B C D 83 IP: Cas Figure 3.8 Integrin-stimulated Cas-Src association requires PTPα-Tyr789. Wild-type (PTPα+/+ or +/+) and PTPα-/- (-/-) MEFs, or PTPα-/- MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα, were cultured 24h, starved with 0.5% FBS-DMEM for 24h, placed in suspension for 1h (0), and then plated on FN- coated dishes for 15 min (15). (A) Cas immunoprecipitates were probed for Src and Cas. (B) Results from 3 experiments were quantified by densitometry and the Src per unit Cas is shown in the graphs as mean ± S.D. Asterisks indicate significant differences (*, p=5×10-4; **, p=1.5×10-8). Cas Src PTPα: 0 15 +/+ 0 15 WT 0 15 0 15 AdV: Y789F ‐/‐ ‐/‐ ‐/‐ * ** 1.0 0.8 0.6 0.4 0.2 Sr c/ Ca s PTPα: AdV: +/+ ‐/‐ ‐/‐ ‐/‐ WT Y789F 84 B A Figure 3.9 PTPα Tyr789 directs PTPα to integrin-mediated focal adhesion- enriched fractions. Wild-type (+/+) and PTPα-/- (-/-) MEFs, or PTPα-/- MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα were cultured 24h, starved with 0.5% FBS- DMEM for 24h, placed in suspension for 1h (0), and then plated on FN-coated dishes for 15 min (15). Cell lysates were fractionated into soluble (S) and insoluble (I) focal adhesion-enriched fractions as described in Methods. (A) Cell fractions were probed for phosphoTyr789-PTPα (pY789) and PTPα. (B ) Results from 3 experiments were quantified by densitometry and the amount of total PTPα in the soluble and insoluble fractions was taken as 100%. Graphs show the percent of PTPα in insoluble fractions as mean ± S.D. Asterisks indicate significant differences (*, p=0.0001; **, p=0.0004; ***, p=0.0094) and n.s. indicates no significant difference (p=0.0934). % ins ol ub le PT Pα 20 40 60 80 100 0 15 0 15 PTPα: +/+ * 100 20 40 60 80 0 15 0 15 WT Y789F ** n.s. *** ‐/‐ PTPα pY789 PTPα: AdV: 0 15 0 15 +/+ S IS S ISI I WT Y789F 0 15 0 15 S IS I S IS I ‐/‐ ‐/‐ ‐/‐ 85 B A Figure 3.10 PTPα-Tyr789 directs Cas and Crk, but not Src, to integrin-mediated focal adhesion-enriched fractions. Wild-type (PTPα+/+, +/+) and PTPα-/- (-/-) MEFs, or PTPα-/-MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα were cultured 24h, starved with 0.5% FBS-DMEM for 24h, placed in suspension for 1h (0), and then plated on FN- coated dishes for 15 min (15). Cell lysates were fractionated into soluble (S) and insoluble (I) focal adhesion-enriched fractions as described in Methods. (A) Cell fractions were probed for Cas, Crk, and Src. Results from 3 experiments were quantified by densitometry and the amount of total Cas or Src in the soluble and insoluble fractions was taken as 100%. Graphs show the percent of (B) Cas and (C) Src in insoluble fractions of FN-plated cells as mean ± S.D. Asterisks indicate significant differences (*, p=0.0070; **, p=0.0397), and n.s. indicates no significant differences (left, p=0.8535; right, p=0.6001). 20 40 30 10 20 40 30 10 % ins ol ub le Ca s at FN 15 * ** PTPα: ‐/‐ WT Y789F+/+ 20 40 60 80 100 20 40 60 80 100 % ins ol ub le Sr c at FN 15 n.s. n.s. PTPα: ‐/‐ WT Y789F+/+ Cas Src Crk PTPα: AdV: 0 15 0 15 +/+ S IS S ISI I WT Y789F 0 15 0 15 S IS I S IS I ‐/‐ ‐/‐ ‐/‐ 86 A CB 87 Chapter 4: Cas Complexes with PTPα via its Binding Protein BCAR3 to Regulate Cell Migration 4.1 Rationale BCAR3 was first discovered together with Cas (also called BCAR1 in the study) by a screen for genes responsible for conferring anti-estrogen resistance in breast cancer cells (van Agthoven et al. 1998).. The BCAR3 C-terminal region constitutively associates with Cas and is required for downstream signaling upon integrin activation (Cai et al. 1999; Riggins et al. 2003). BCAR3-enhanced migration is abolished when the BCAR3 GEF domain responsible for Cas binding is deleted or mutated (Riggins et al. 2003; Vanden Borre et al. 2011). SH2 domains are well-known ~110 amino acid modules that can bind to phosphotyrosine to regulate various signaling pathways (Pawson et al. 2001). The deletion or the defective point mutant of the BCAR3 SH2 domain failed to induce Rac1 activation in breast cancer cells (Felekkis et al. 2005). The specificity of the BCAR3 SH2 domain is still unknown, and no intracellular ligand has been identified yet (Vanden Borre et al. 2011). Interestingly, BCAR3 depletion in NIH 3T3 cells phenocopies the defects of Y789F PTPα-expressing MEFs, such as resulting in reduced Cas SD tyrosine phosphorylation and decreased Rac1/Cdc42 and PAK activation (Cai et al. 2003). BCAR3 overexpression promotes breast cancer cell or fibroblast spreading on fibronectin, migration, and invasion; while depletion of BCAR3 inhibits these cell properties (Riggins et al. 2003; Schuh et al. 2010). Importantly, BCAR3 can regulate Cas FA localization in MEFs and breast cancer cells and both its SH2 and GEF domains are required for this function (Schrecengost et al. 2007; Vanden Borre et al. 2011). These results suggest that the above actions of BCAR3 may 88 be dependent on its GEF domain-mediated interaction with Cas and, since its SH2 ligand is unknown, an unidentified phosphotyrosine site. Since BCAR3 structurally has the potential to function as a linker between PTPα phosphoTyr789 and Cas, and functionally acts in integrin signaling as well as BCAR3 deficiency phenocopying mutant PTPα Y789F signaling defects, I investigated the role of BCAR3 in PTPα Tyr789-dependent Cas regulation in integrin signaling. 4.2 BCAR3 knockdown cells have a similar phenotype to mutant Y789F PTPα- expressing cells Firstly, I confirmed that several of the reported molecular actions of BCAR3 mimicked those of PTPα Tyr789, namely those in Src activation, Cas tyrosine phosphorylation, Rac/Cdc42, and PAK activation in MEFs. Wildtype MEFs were depleted of about 90% of BCAR3 using siRNA (Fig. 4.1A, top panels). Consistent with Schuh et al. (2010), the depletion of BCAR3 inhibits the integrin-induced phosphorylation of Tyr410 in the Cas SD and almost eliminates downstream signaling pathway activation as evidenced by reduced PAK activation (Fig. 4.1A, middle and bottom panels). Src activation results from the phosphorylation of Tyr416 in its activation loop and the dephosphorylation of its inhibitory Tyr527 site which exposes the kinase domain by a conformational change (Smart et al. 1981). My results showed that Src Tyr416 phosphorylation is indeed abrogated in BCAR3-deficient MEFs (43±14% of that in control cells, p=0.001), while the phosphorylation of Tyr527 is similar between control siRNA and BCAR3 siRNA treated wildtype MEFs (92±11% of that in control cells, p=0.14) (Fig. 4.1B). This indicated that integrin-induced Src activation is indeed reduced upon the depletion of 89 BCAR3 (Sheetz et al. 2006; Schuh et al. 2010). As Src activation is necessary for the phosphorylation of PTPα Tyr789, I also probed the cell lysates for phospho-PTPα and PTPα. Notably, integrin-induced phosphorylation of PTPα Tyr789 was also reduced to 47±7% (p=7×10 -5 ) in BCAR3 knockdown cells (Fig. 4.1C). Thus, BCAR3 regulates not only Src activation through the phosphorylation of Tyr416 of Src, but also the phosphorylation of PTPα Tyr789 through Src activation. I then examined if integrin-induced Cas re-localization to focal adhesions (FAs) is regulated by BCAR3, using the FA enrichment assay described in Chapter 3 (Section 3.5). Cas localization in FA-enriched fractions was significantly reduced by about one-third upon BCAR3 depletion (from 17±2.5% in control cells to 12±1.5% in the BCAR3-deficient cells, p=0.04) (Fig. 4.2A). Src re-localization was unaffected (78±7.8% in control cells compared to 73±8.5% in BCAR3-depleted cells, p=0.49) (Fig. 4.2B), suggesting that BCAR3 is not required for Src FA translocation. PTPα re-localization to the FA-enriched fractions was also impaired by BCAR3 siRNA treatment (40±3% in BCAR3-deficient cells compared to 66±1% in control cells, p=2×10 -4 ) (Fig. 4.2C). This is likely due to the reduced phosphorylation of PTPα Tyr789, which in turn may underlie the reduced Cas re-localization in the BCAR3-depleted cells. The above findings confirmed the remarkably similar phenotype between BCAR3- deficient and Y789F PTPα-expressing MEFs, indeed suggesting that BCAR3 and phosphoTyr789-PTPα may act within the same signaling pathway to regulate cell migration. 90 4.3 Integrin-induced FA localization of the Cas binding protein BCAR3 requires PTPα Tyr789 The data presented in Chapter 3 demonstrated that the phosphorylation of PTPα at Tyr789 not only directs PTPα to forming FAs, but also plays an important role in bringing Cas to focal adhesions (Fig. 3.6). If BCAR3 and phosphoTyr789-PTPα act within the same signaling pathway, the structural features and binding properties of BCAR3 raise the possibility that it might be an intermediate in the PTPα-Tyr789-dependent recruitment of Cas to FAs. To test this hypothesis, I investigated if BCAR3 indeed underwent integrin- stimulated translocation to FA-enriched fractions and if this required an intact Tyr789 site within PTPα. As shown in Fig. 4.3, a portion (20.5%) of BCAR3 moved into FA-enriched fractions upon integrin engagement of wild-type MEFs, but much less BCAR3, only 2.7%, relocated to these fractions in cells lacking PTPα. Moreover, expression of WT PTPα in PTPα-/- cells effectively promoted BCAR3 movement into the FA-enriched fractions (29±3.5%), whereas expression of Y789F PTPα induced two-fold less BCAR3 re- localization (15±9.5%). These results showed that PTPα-phosphoTyr789 is important for the recruitment of not only Cas but also BCAR3 to focal adhesions. In summary, the above results reveal that BCAR3-mediated activation of Src, possibly acting in concert with PTPα-mediated activation of Src (Su et al., 1999; Zeng et al., 2003), promotes PTPα-Tyr789 phosphorylation to induce the re-localization of PTPα to focal adhesions. Thus not only phospho-Tyr789 of PTPα is required for BCAR3 and Cas FA localization, but BCAR3 also regulates PTPα and Cas localization to focal adhesions by stimulating Src activation and Src-dependent PTPα-Tyr789 phosphorylation. This suggests there are two different roles of BCAR3 in this signaling pathway: 1) an upstream function to 91 regulate Src activation and PTPα-Tyr789 phosphorylation; 2) a downstream function to promote integrin-induced Cas FA localization through phosphoTyr789-PTPα. Since the second role of BCAR3 is also dependent on PTPα-Tyr789 phosphorylation, I hypothesized that in this context, BCAR3 functions as an intermediate in PTPα-Tyr789-dependent recruitment of Cas to focal adhesions. 4.4 PhosphoTyr789 of PTPα directly interacts with the SH2 domain of BCAR3 The phosphorylation of PTPα Tyr789 and BCAR3 expression are necessary for Cas FA localization, and the fractionation studies revealed that phosphoTyr789-PTPα, BCAR3, and Cas translocate to FAs upon integrin activation. This raised the question of whether PTPα physically interacted with BCAR3, and via BCAR3, with Cas. 4.4.1 PTPα associates with Cas and Src in an integrin- and BCAR3- dependent manner To investigate if BCAR3 was required for the interaction between PTPα and Cas, VSVG- tagged WT PTPα was reconstituted in PTPα-/- MEFs that were treated with control non- targeting or BCAR3 siRNA. Lysates of adherent parental (untransfected PTPα-/- MEFs), control, and BCAR3 knockdown cells were probed with BCAR3 and actin antibodies to confirm the depletion of BCAR3 and equivalent sample loading, respectively (Fig. 4.4A, bottom panels). Probing VSVG-PTPα immunoprecipitates prepared from these lysates for VSVG (PTPα), BCAR3, Cas, and Src showed that BCAR3 indeed associated with PTPα as well as Cas and Src, since they were all present in the immunoprecipitates from the control cells that expressed BCAR3 (Fig. 4.4A, top panels). However, BCAR3 depletion virtually 92 abolished the association of Cas and Src with PTPα (Fig. 4.4A, top panels). This indicated that BCAR3 interacted with PTPα and was essential for the interaction of PTPα and Cas. However, these results could not differentiate between the possible roles of BCAR3 in mediating the PTPα-Cas interaction physically (i.e. as a molecular bridge) or by promoting the phosphorylation of PTPα Tyr789 (as shown in Fig. 4.1C) to induce an association between PTPα and Cas that was physically independent of BCAR3. However, the second possibility seemed less viable, since the ~50% reduction in PTPα Tyr789 phosphorylation in BCAR3 depleted cells (Fig. 4.1D and E) was unlikely sufficient to virtually abolish the interaction of PTPα with Cas. My results also established that PTPα is not required for the interaction of BCAR3 and Cas, since Cas immunoprecipitates from wild-type and PTPα-deficient MEFs contained equivalent amounts of associated BCAR3 (Fig. 4.4B). BCAR3-depleted MEFs (Fig. 4.4B, right lane) were used as a negative control in this experiment to confirm that the signal detected by anti-BCAR3-antibody on the blot (~95kD) was indeed endogenous BCAR3. To confirm that the PTPα interaction with BCAR3, Cas, and Src is an integrin-regulated event that occurs in concert with PTPα Tyr789 phosphorylation, the PTPα-/- MEFs expressing VSVG-tagged WT PTPα were maintained in suspension for one hour, and then placed on FN coated plates for 15 minutes to engage integrins. Cell lysates were used to prepare VSVG-PTPα immunoprecipitates that were probed for PTPα phosphoTyr789, VSVG (PTPα), Cas, BCAR3, and Src. In suspended cells where integrin signaling is not operational PTPα is not associated with BCAR3, Cas, or Src, and PTPα is not phosphorylated. Stimulating integrin signaling induced both the phosphorylation of PTPα-Tyr789 and the association of BCAR3, Cas, and Src with PTPα (Fig. 4.5). 93 These results showed that PTPα, BCAR3, Cas, and Src can form a complex in MEFs, and both integrin stimulation (probably through the integrin–induced phosphorylation of PTPα Tyr789) and BCAR3 are required for this complex formation. This indicates that phospho- Tyr789 PTPα may function through BCAR3 to form a complex with Cas and Src to promote integrin-induced cell migration. 4.4.2 PTPα Tyr789 phosphorylation is required for PTPα-BCAR3 interaction I next investigated whether Tyr789 of PTPα is required to mediate the association of PTPα with BCAR3, Cas, and Src. Lysates were prepared from VSVG-tagged WT and Y789F PTPα expressing cells (Fig. 4.6A, bottom panels), and VSVG immunoprecipitates were isolated and probed for VSVG-PTPα, BCAR3, Cas, and Src (Fig. 4.6A, top panels). BCAR3 associated with phosphoTyr789-containing WT PTPα but was not detected in association with unphosphorylatable Y789F PTPα. Cas and Src were also found in association with WT PTPα, but no Cas and a greatly reduced amount of Src interacted with Y789F PTPα. Together, these results indicated that an intact Tyr789 residue of PTPα is necessary for its interaction with BCAR3, Cas and Src. To investigate the role of PTPα Tyr789 phosphorylation in the PTPα-BCAR3 interaction, I further examined whether the PTPα phosphoTyr789-specific antibody could compete with BCAR3 for PTPα binding. Lysates from cells expressing VSVG-tagged WT PTPα were incubated with or without the anti-phosphoTyr789 antibody and PTPα was immunoprecipitated using bead-conjugated anti-VSVG antibody. Probing the immunoprecipitates for VSVG (PTPα), BCAR3, Cas, and Src showed that the interaction of BCAR3, as well as that of Cas and Src, with PTPα was virtually eliminated in the presence of 94 the phosphoTyr789 antibody (Fig. 4.6B), strongly suggesting that phosphoTyr789 was the site for BCAR3 binding. Such an interaction is most likely mediated through the SH2 domain of BCAR3, so I next investigated if the SH2 domain of BCAR3 is essential for this interaction. 4.4.3 BCAR3-SH2 domain is required for the PTPα-phosphoTyr789-BCAR3 interaction The role of the BCAR3 SH2 domain in mediating binding to phosphoTyr789 of PTPα was investigated using GST-pulldown assays where recombinant GST fusion protein containing the SH2 domain of BCAR3 (GST-SH2) was mixed with lysates prepared from wild-type and PTPα-/- MEFs, or from PTPα-/- MEFs expressing VSVG-WT PTPα or VSVG-Y789F PTPα. Glutathione-bead mediated pulldowns of the GST-SH2 domain from the latter two cell types contained associated WT PTPα but not Y789F PTPα (Fig. 4.7, lane 3 and 4). A lower amount of endogenous PTPα was also detected in pull-downs from wild-type MEFs (Fig. 4.7, lane 1). These results indicate that the BCAR3-SH2 domain can interact with phosphoTyr789 PTPα. To confirm that BCAR3 SH2 domain directly bound to phosphoTyr789-PTPα, I conducted a ‘Far Western’, where anti-VSVG immunoprecipitates from lysates of VSVG- WT or VSVG-Y789F PTPα-expressing cells were resolved by SDS-PAGE and transferred to a PVDF membrane, and then incubated with GST-SH2 (BCAR3) fusion protein. If the BCAR3 SH2 domain associates with the VSVG-PTPα, this interaction is detected by probing with an HRP-conjugated anti-GST antibody on the membrane with the same molecular weight as VSVG-PTPα. Subsequent probing with anti-GST antibody detected GST-SH2 95 (BCAR3) in the WT PTPα sample at a position that coincided with that of WT PTPα, whereas virtually no binding of BCAR3 SH2 domain to Y789F PTPα was observed (Fig. 4.8). These results demonstrated that the BCAR3 SH2 domain is sufficient to directly interact with PTPα Tyr789. 4.5 The binding of BCAR3 and Grb2 to PTPα phosphoTyr789 is mutually exclusive Since the adaptor protein Grb2 also binds through its SH2 domain to PTPα phosphoTyr789 (den Hertog et al. 1994; Su et al. 1994), it would be predicted that BCAR3 and Grb2 binding to phosphoTyr789 would be mutually exclusive. I therefore tested whether complexes of PTPα and Grb2 or BCAR3-SH2 could be distinguished. The GST-SH2 BCAR3 protein was added to lysates of cells expressing VSVG-WT PTPα, and portions of the mixture were immunoprecipitated with anti-Grb2 antibody or used for GST pulldown. The Grb2 immunoprecipitates contained PTPα and Grb2 but not GST-SH2 (BCAR3), whereas the GST pulldowns contained PTPα and GST-SH2 (BCAR3) but not Grb2 (Fig. 4.9), providing further evidence that the BCAR3 SH2 domain recognizes and binds to PTPα phosphoTyr789. 4.6 The SH2 and GEF domains of BCAR3 are respectively required for the physical linkage of PTPα with Cas/Src My results demonstrated that the BCAR3 SH2 domain directly interacts with PTPα Tyr789, so I further investigated whether two mutant forms of BCAR3 with a point mutation in either the SH2 domain (R177K) or the GEF domain (R748A) could bind to phosphoTyr789-PTPα and support the interaction of PTPα with Cas and Src. The SH2 96 domain mutation at R177 is a position in human BCAR3 analogous to a conserved critical Arg residue in SH2 domains that is required for phosphotyrosine binding (Felekkis et al. 2005), and the mutation in the GEF domain at R748 is a position in human BCAR3 that mediates its association with Cas (Vanden Borre et al. 2011). PTPα-/- cells were co- transfected with VSVG-WT PTPα and FLAG-tagged wild-type (WT) or mutant (SH2, R177K; or GEF, R748A) BCAR3 while the parental untransfected PTPα-/- cells served as a negative control. Anti-VSVG and anti-FLAG immunoprecipitates were analyzed for associated proteins. Cell lysates were probed for PTPα phosphoTyr789, PTPα, BCAR3, Cas, and Src as shown in Fig. 4.10C. The phosphorylation of PTPα Tyr789 and the expression of PTPα are similar in all three VSVG-WT PTPα over-expressing cells, which suggests that the over-expression of the BCAR3 mutants does not have a dominant-negative effect on PTPα Tyr789 phosphorylation even though BCAR3 is required for this event. The BCAR3 expression level is also comparable in the three FLAG-BCAR3 over-expressing cells, eliminating the possibility of any effects due to differential expression of BCAR3. The amounts of Cas, Src, and actin are similar to untransfected parental PTPα-/- MEFs and are not affected by the double trans-fection of PTPα- and BCAR3-plasmids (Fig. 4.10C). Both anti-VSVG (Fig. 4.10A) and anti-FLAG (Fig. 4.10B) immunoprecipitates from cells co-expressing WT forms of VSVG-PTPα and FLAG-BCAR3 contained the four members of the protein complex that I had previously detected: PTPα, BCAR3, Cas, and Src. The anti- VSVG immunoprecipitates from cells co-expressing tagged WT PTPα and the SH2 defective mutant of BCAR3 contained almost no BCAR3, Cas, or Src, as would be expected if BCAR3-SH2 mediated the association of PTPα with Cas and Src. In accord with this, virtually no PTPα was detectable in the anti-FLAG immunoprecipitates from these cells (Fig. 97 4.10B) despite the presence of phosphoTyr789-PTPα in the lysates (Fig. 4.10C). However Cas was present, consistent with its interaction with BCAR3 being independent of the BCAR3 SH2 domain, as was a reduced amount of Src. Reciprocal anti-VSVG or anti-FLAG immunoprecipitates from cells co-expressing tagged WT PTPα and the GEF mutant of BCAR3 respectively contained associated BCAR3 or PTPα, but the association appeared reduced compared to that detected in anti-VSVG immunoprecipitates from cells expressing WT BCAR3. Virtually no Cas or Src was co-immunoprecipitated (Fig. 4.10A and B). The GEF mutation in BCAR3 interferes with BCAR3-Cas association (Vanden Borre et al. 2011), suggesting that Cas binding to BCAR3 may modulate BCAR3 localization or conformation for optimal BCAR3-SH2-mediated interaction with PTPα. Together, the above results demonstrated that an intact SH2 domain of BCAR3 is required for binding to phosphoTyr789-PTPα, confirmed that an intact GEF domain within BCAR3 is required for its binding to Cas, and showed that these interactions mediate formation of the PTPα-BCAR3-Cas/Src complex. 4.7 Discussion Since Cas does not possess SH2 or PTB domains that could directly interact with phosphoTyr789, I postulated the involvement of an intermediary protein -BCAR3, which contains not only a GEF domain that constitutively binds to Cas but also an SH2 domain that has the potential to bind to PTPα phosphoTyr789. I found that in support of BCAR3 mediating the interaction of PTPα and Cas, a complex of PTPα-BCAR3-Cas-Src is detectable and the presence of Cas and Src in this complex is dependent on BCAR3, since Cas and Src showed almost no association with PTPα in BCAR3-depleted MEFs. As PTPα 98 Tyr789 phosphorylation is defective due to impaired Src activation in the BCAR3-depleted cells, this impaired interaction could be due to the absence of the phosphoTyr789 binding site on PTPα. However, the interaction of PTPα with BCAR3, Cas and Src is also abolished in cells over-expressing BCAR3 with a mutant SH2 domain, even though Tyr789 of PTPα is phosphorylated. In addition, the interaction of BCAR3 with Cas/Src is eliminated by a mutation in the BCAR3 GEF domain, while this mutant BCAR3 can still interact with phosphoTyr789-PTPα. Together, this evidence reveals BCAR3 as a molecular linker of PTPα and Cas. The ability of the BCAR3 SH2 domain to bind to PTPα phosphoTyr789 was unexpected, as this SH2 domain is not closely related to either of two other SH2 domains, those of Grb2 and Src (Sastry et al. 2006), that can bind phosphoTyr789. This study identifies phosphoTyr789 PTPα as the first cellular ligand for the SH2 domain of BCAR3. Not only does formation of the complex require PTPα Tyr789 and an intact SH2 domain of BCAR3, but recombinant BCAR3 SH2 domain interacts directly with WT PTPα and not with Y789F PTPα. Both the SH3 domain and the CCH domain (Cas family C-terminal homology domain, ~140 amino acid residues at Cas C-terminus) of Cas have been shown to mediate its FA localization, and both domains are required for optimal focal adhesion recruitment and tyrosine phosphorylation of Cas (Nakamoto et al. 1997; Harte et al. 2000; Donato et al. 2010). The N-terminal SH3 domain of Cas interacts with the FAK proline-rich region (residues 711-717) upon integrin stimulation (Polte and Hanks 1995), and as a prominent FA localized kinase FAK has been proposed to mediate the recruitment of Cas to FAs (Nakamoto et al. 1997). The deletion of the Cas SH3 domain reduces the Cas FA localization to only 1/6 of that observed with wildtype Cas, and re-expression of FAK in FAK -/- cells 99 increases Cas FA localization 3-fold, confirming that the association of the Cas SH3 domain with FAK is important for Cas FA localization (Donato et al. 2010). The CCH domain of Cas is also essential for its FA localization but the molecular basis for this is unclear (Harte et al. 2000; Donato et al. 2010). A series of recent studies suggested a BCAR3-mediated mechanism for the recruitment of Cas to FAs, since BCAR3 constitutively associates with Cas (Cai et al. 1999). Upon integrin stimulation BCAR3 and Cas co-localize at the cell periphery and in migration-associated structures such as ruffles and lamellipodia (Riggins et al. 2003), while depletion of BCAR3 eliminates the membrane ruffle localization of Cas (Schrecengost et al. 2007). This indicates that the interaction between the C-terminal GEF- like domain of BCAR3 and the CCH domain of Cas is required for optimal Cas FA localization and integrin-stimulated migration (Gotoh et al. 2000; Donato et al. 2010). The BCAR3 SH2 domain is required for BCAR3 FA localization, while the GEF domain is necessary for mediating Src activation and promoting cell migration (Riggins et al. 2003). The combined results described in Chapter 3 and 4 support a model where integrin activation induces Src-mediated phosphorylation of PTPα at Tyr789, resulting in formation and localization of a PTPα-BCAR3-Cas complex at nascent adhesions, which may underlie the CCH domain dependent FA localization of Cas. Another function of BCAR3 is to mediate Src activation (Schuh et al. 2010). In the present study I found that BCAR3 is required for Src Tyr416 phosphorylation and PTPα Tyr789 phosphorylation that promotes BCAR3-mediated Cas localization to FAs, suggesting that BCAR3 plays two roles of in integrin signaling. The first role of BCAR3 is to activate Src kinase, which is upstream of PTPα. Interestingly, PTPα activates Src by dephosphorylation of Src Tyr527 (inhibitory site) but BCAR3 promotes Src kinase activity 100 by phosphorylation of Src Tyr416 (kinase activation loop). They clearly target different regulatory tyrosine residues in Src and the absence of either protein leads to defective Src activation, suggesting that the roles and interactions of PTPα and BCAR3 in Src activation need further investigation. The second role is to bridge phosphoTyr789 PTPα and Cas association, which is downstream of PTPα and facilitates Cas FA localization to promote cell migration. In summary, this study reveals a novel molecular complex of PTPα-BCAR3-Cas-Src. This complex forms in response to PTPα Tyr789 phosphorylation and mediates Cas localization to focal adhesions and Cas downstream signaling to promote cell migration. As a critical regulator of cell motility, Cas overexpression in tumor tissue has been associated with increased proliferation, metastasis, acquired Tamoxifen-resistance, and poor prognosis (van der Flier et al. 2000; van der Flier et al. 2001; Dorssers et al. 2004; Tornillo et al. 2011). In cultured cell systems, both Cas overexpression and hyper-phosphorylation are correlated with an up-regulation of Src activity to induce cell transformation and increase the matrix metalloproteinases (MMPs) expression to promote invasion (Matsuda et al. 1990; Brabek et al. 2004). Overexpressing Cas and ERBB2 in mouse mammary gland induces enhanced tumor formation (Cabodi et al. 2006), while depletion of Cas in ERBB2 transformed cells inhibits tumor growth and Src, FAK, and Akt activation (Cabodi et al. 2010). Based on my results, disruption of the PTPα-BCAR3-Cas module may inhibit the movement and invasive ability of cancer cells. In the next chapter, I describe the investigation of whether this signaling complex has a role in regulating these properties of cancer cells. BCAR3 actin pS144 PAK pY410 Cas BCAR3siRNA: Con 20 60 100 pY 78 9 / PT Pα ** PTPα pY789 BCAR3siRNA: Con Con BCAR3siRNA: A Figure 4.1 BCAR3 promotes Src activation, PTPα Tyr789 phosphorylation, and downstream signaling. Wild-type MEFs were transfected with 100nM non-targeting (control, Con) or BCAR3-targeted siRNA. After 48 hours, the cells were suspended for 1h and then plated on FN for 15min. Cell lysates were prepared and were probed for (A) BCAR3, actin, phosphoTyr410-Cas, Cas, phosphoSer144-PAK, and PAK; and (B) for phospho- Tyr416- and phospho-Tyr527-Src and Src. The results from 3 experiments were quantified by densitometry, and the Src Tyr416 phosphorylation and Src Tyr527 phosphorylation per unit Src are shown in the graphs as mean ± S.D. The asterisk indicates a significant difference with the control (*, p=0.001). (C) Lysates from the cells were also probed for phosphoTyr789-PTPα and PTPα. The results from 3 experiments were quantified by densitometry, and the phospho-Tyr789 per unit PTPα is shown in the graphs as mean ± S.D. Asterisks indicate a significant difference with the control (**, p=7.3x10-5). B Src Con BCAR3 Ph os ph o‐S rc siRNA: pY416 pY527 pY527 pY416 BCAR3siRNA: Con 100 60 20 * C 101 PTPαCas Src siRNA: S IS I Con BCAR3 BA Figure 4.2 BCAR3 promotes PTPα and Cas FA localization. Wild-type MEFs were transfected with 100nM control non-targeting (Con) or BCAR3-targeted siRNA. After 48 hours, the cells were suspended for 1h and then plated on FN for 15min. (A) Soluble (S) and insoluble (I) focal adhesion-enriched fractions were prepared from the cells and probed for Cas. The results from 3 experiments were quantified by densitometry and the amount of total Cas in the soluble and insoluble fractions was taken as 100%. The graph shows the percent of Cas in the insoluble cell fractions as mean ± S.D. The asterisk denotes a significant difference (*, p=0.042). (B) Soluble (S) and insoluble (I) focal adhesion-enriched fractions were prepared from the cells and probed for Src. The results from 3 experiments were quantified by densitometry and the amount of total Src in the soluble and insoluble fractions was taken as 100%. The graph shows the percent of Src in the insoluble cell fractions as mean ± S.D. (p=0.49). (C) Soluble (S) and insoluble (I) focal adhesion-enriched fractions were prepared from the cells and probed for PTPα. The results from 3 experiments were quantified by densitometry and the amount of total PTPα in the soluble and insoluble fractions was taken as 100%. The graph shows the percent of PTPα in the insoluble cell fractions as mean ± S.D. The asterisks denote a significant difference (**, p=2×10-4). 5 10 15 20 * % ins ol ub le Ca s Con BCAR3siRNA: 20 40 60 80 ** % ins ol ub le PT Pα Con BCAR3 S IS I Con BCAR3 S IS I Con BCAR3C 20 40 60 80 % ins ol ub le Sr c Con BCAR3 100 102 Figure 4.3 PTPα Tyr789 regulates integrin-induced BCAR3 localization to focal adhesion-enriched fractions. Wild-type (+/+) and PTPα-/- (-/-) MEFs, or PTPα-/- MEFs infected with adenovirus (AdV) expressing WT or Y789F PTPα, were placed in suspension for 1 h (0) and then plated on FN-coated dishes for 15 min (15). Cell lysates were fractionated into soluble (S) and insoluble (I) focal adhesion-enriched fractions. (A) Cell fractions were probed for BCAR3 and PTPα. (B)The results from +/+ and -/- cells are from one experiment, and the results from WT and Y789F cells are representative of 3 experiments. The signals were quantified by densitometry and the amount of total BCAR3 in the soluble and insoluble fractions was taken as 100%. Graphs show the percent of BCAR3 in insoluble fractions as mean ± S.D. The asterisk indicates a significant differences (*, p=0.04). BCAR3 PTPα PTPα: AdV: 0 15 0 15 +/+ S IS S ISI I WT Y789F 0 15 0 15 S IS I S IS I ‐/‐ ‐/‐ ‐/‐ 10 20 15 5% ins ol ub le BC AR 3 at FN 15 * PTPα: ‐/‐ WT Y789F+/+ 25 30 35 103 Figure 4.4 BCAR3 is required for the formation of a PTPα-BCAR3-Cas-Src complex and BCAR3-Cas association is independent of PTPα. PTPα-/- MEFs were transfected with control non-targeting (Con) or BCAR3-targeted siRNA and with or without VSVG-tagged WT PTPα plasmid. (A )Anti-VSVG immuno-precipitates from lysates of adherent cells were probed for BCAR3, Cas, Src, and VSVG (VSVG-PTPα). Lysates were probed for BCAR3 and actin. (B) Cas immuno-precipitates were prepared from wild-type (+/+) or PTPα-/- (-/-) MEFS, and as a control, wild-type MEFs from which BCAR3 was depleted using siRNA. The immunoprecipitates were probed for BCAR3 and Cas. IP‐VSVG VSVG (PTPα) Src Cas BCAR3 BCAR3 actin lysate VSVG‐PTPα: WT WT Con BCAR3siRNA: ‐/‐ ‐ ‐ BCAR3 Cas IP‐Cas PTPα: +/+ BCAR3siRNA: ‐/‐ +/+ Con Con A B 104 Figure 4.5 The formation of the PTPα-BCAR3-Cas-Src complex is regulated by integrin. PTPα-/- MEFs were left untransfected or transfected with VSVG-WT PTPα plasmid. The transfected cells from cultures in uncoated dishes (Ad) were placed in suspension for 1h (0) and then plated on FN-coated dishes for 15 min (15). VSVG immuno- precipitates from cell lysates were probed for Cas, BCAR3, phosphoTyr789 PTPα, and VSVG (VSVG-PTPα). Src BCAR3 VSVG (PTPα) PTPα‐pY789 Cas IP‐VSVG Ad 0 15 ‐/‐ WT WTVSVG‐PTPα: ‐ 105 IP‐VSVG actin PTPα pY789 VSVG (PTPα) BCAR3 Cas Src PTPα: +/+ WT Y789FVSVG‐PTPα: ‐/‐ ‐/‐ ‐/‐ lysate Figure 4.6 PTPα Tyr789 is required for the formation of a PTPα-BCAR3-Cas- Src complex. (A) Lysates from adherent cultures of wild-type (+/+) and PTPα-/- MEFs (-/-), and from PTPα-/- MEFs transfected with VSVG-tagged WT PTPα or Y789F PTPα plasmids, were used to prepare VSVG immunoprecipitates. These were probed for BCAR3, Cas, Src, and VSVG (VSVG-PTPα). Lysates were probed for phosphoTyr789-PTPα, PTPα, and actin. (B) PTPα-/- MEFs were untransfected or transfected with VSVG-tagged WT PTPα plasmid. Lysates were mixed with anti-phosphoTyr789 PTPα antibody as indicated, and anti-VSVG immunoprecipitates were prepared and probed for BCAR3, Cas, Src, and VSVG (VSVG-PTPα). VSVG (PTPα) BCAR3 Cas Src IP‐VSVG WTVSVG‐PTPα: WT PTPα‐pY789 Ab: ‐ ‐ + PTPα‐/‐ ‐ A B ‐ ‐ 106 Figure 4.7 BCAR3-SH2 domain is required for BCAR3 to associate with phosphoTyr789 of PTPα. Lysates from adherent cultures of wild-type (+/+) and PTPα-/- MEFs (-/-), and from PTPα-/- MEFs transfected with VSVG-tagged WT PTPα or Y789F PTPα plasmids were used to prepare GST-BCAR3-SH2 pull-downs. Pull-downs were probed for VSVG (PTPα), PTPα, and GST (BCAR3-SH2). GST‐ pulldown VSVG GST‐SH2 PTPα PTPα: +/+ WT Y789FVSVG‐PTPα: ‐/‐ ‐/‐ ‐/‐ ‐‐ 107 IP‐VSVG GST‐SH2 VSVG (PTPα) ‐/‐ WT Y789FVSVG‐PTPα: Figure 4.8 PTPα phosphoTyr789 directly interacts with the BCAR3-SH2 domain. VSVG immunoprecipitates from adherent PTPα-/- MEFs (-/-) that were untransfected or transfected with VSVG-tagged WT and Y789F PTPα plasmids were probed with GST- BCAR3-SH2 fusion protein. Bound GST-SH2 domain was detected using anti-GST antibody. ‐ ‐ 108 VSVG GST Grb2 IP‐Grb2 GST‐ pulldown WT + + PTPα‐/‐ VSVG‐PTPα: GST‐SH2: WT lysate WT + PTPα Figure 4.9 BCAR3 and Grb2 binding to phosphoTyr789 PTPα are mutually exclusive. PTPα-/- MEFs were untransfected or transfected with VSVG-tagged WT PTPα plasmids. Lysates of VSVG-WT PTPα-expressing cells were mixed with recombinant GST-SH2 (BCAR3) fusion protein as indicated, and then these and the untransfected cell lysates were used to prepare anti-Grb2 immunoprecipitates or GST-SH2 pull- downs. Samples were probed for VSVG (PTPα), GST (BCAR3-SH2), and Grb2. ‐ ‐ ‐ ‐ 109 Figure 4.10 BCAR3-SH2 and -GEF domains are required for BCAR3 to associate with and link phosphoTyr789 of PTPα and Cas. PTPα-/- MEFs (-/-) were co-transfected with VSVG-WT PTPα and FLAG-tagged WT BCAR3 or mutant forms of BCAR3 (SH2, R177K mutation in the SH2 domain; GEF, R748A mutation in the GEF domain) plasmids. Anti-VSVG (A) and anti-FLAG (B) immuno-precipitates and cell lysates (C) were probed for the indicated proteins or epitope tags. BCAR3 Cas Src VSVG (PTPα) IP‐FLAG IP‐VSVG Cas VSVG (PTPα) Src FLAG (BCAR3) ‐/‐ VSVG‐PTPα: WT GEFFLAG‐BCAR3: WT WT WT‐ SH2‐ A B actin Lysate PTPα BCAR3 Src Cas FLAG VSVG pY789 C 110 111 Chapter 5: Disruption of PTPα-BCAR3-Cas Complex Formation Inhibits the Migration and Invasion of RD Rhabdomyosarcoma Cells 5.1 Rationale PTPα has been implicated in promoting carcinogenesis in several types of tissues. PTPα is overexpressed in late stage colon carcinomas, and it was proposed that PTPα can induce the transformation of colon epithelial cells (Tabiti et al. 1995). The expression of PTPα is also up-regulated in serum-stimulated androgen-sensitive prostate cancer cell lines, PC-3 and DU145 (Zelivianski et al. 2000). Another study reported that 15 of 51 breast cancer samples had increased expression of PTPα, which correlated with low-grade breast cancer (Ardini et al. 2000). In the same study, these authors demonstrated that expressing PTPα in invasive breast cancer cell lines inhibits growth and metastasis (Ardini et al. 2000). PTPα is also involved in the progression of gastric carcinoma by functioning as a receptor for VacA of Helicobacter pylori, a bacterium that can induce the apoptosis of gastric epithelial cells (Yahiro et al. 2003). However, a function of PTPα Tyr789 phosphorylation in cancer progression and metastasis has not been reported. BCAR3 expression and BCAR3-Cas association is required for anti-estrogen resistance in breast cancer cells (Cai et al. 2003; Near et al. 2007; Garron et al. 2009). BCAR3 overexpression has also been shown to promote cell cycle progression and DNA synthesis in the mammary epithelial cell line MCF-12A (Oh et al. 2008). Based on data collected from biopsies, the overexpression of Cas correlates with breast cancer and leukemia (Salgia et al. 1996; van der Flier et al. 2000; Dorssers et al. 2004; Cabodi et al. 2006). 112 The above studies indicate that PTPα, BCAR3, and Cas are required for or involved in promoting tumorigenesis, although the detailed molecular mechanisms of their actions are still not known. In my previous study in MEFs, I showed that integrin-induced PTPα- BCAR3-Cas-Src complex formation is dependent on PTPα Tyr789 phosphorylation, and this mediates Cas FA localization to promote cell spreading and migration. Since cell spreading and migration are two essential processes for cancer cell invasion and metastasis, I hypothesized that the PTPα-BCAR3-Cas complex plays a role in promoting tumor cell invasion. I investigated if 1) cancer cells use the same complex to regulate migration and invasion; and 2) disruption of this complex will inhibit these cancer cell properties. 5.2 Expression of PTPα, BCAR3, and Cas in cancer cell lines BCAR3 expression level is undetectable in less aggressive epithelial breast cancer cell lines, such as MCF7 and ZR-75, while it is highly expressed in aggressive breast cancer cell lines, such as BT-549 and MDA-321 (van Agthoven et al. 1998; Near et al. 2007; Makkinje et al. 2009). Depletion of BCAR3 from BT-549 cells inhibits migration and invasion by 50% or more (Schrecengost et al. 2007), while overexpression of BCAR3 in MCF-7 cells induces anti-estrogen resistance, cell migration and invasion (Cai et al. 2003; Near et al. 2007; Vanden Borre et al. 2011). Constitutive association of BCAR3 with Cas has been shown in both BT-549 and MCF-7 cells, and BCAR3 depletion in BT-549 cells results in the absence of Cas at the cell membrane, indicating that the BCAR3-Cas interaction has a role in promoting BCAR3-mediated cell migration and invasion (Schrecengost et al. 2007; Vanden Borre et al. 2011). However, it is not clear if this BCAR3-Cas complex plays a similar role in other tumor types. 113 My initial investigation of Cas and BCAR3 expression levels in some non-breast cancer cell lines revealed two extremes of expression: levels comparable to the high expression reported in aggressive BT-549 breast cancer cells, exemplified by the rhabdomyosarcoma RD cell line; and virtually undetectable expression as found in the Ewing’s sarcoma TC-32 cell line (Fig. 5.1A). All the cells were found to express PTPα that was phosphorylated at Tyr789. The presence of equivalent amounts of lysate protein from the different cell lines was confirmed by probing for actin. Rhabdomyosarcoma (RMS) is the most common soft tissue sarcoma in children (Kramer et al. 1983). About 20-25% of RMS patients have distant metastases, and the lung, bone, bone marrow, and lymph nodes are common metastatic sites (Koscielniak et al. 1992). Two subtypes of this malignancy are embryonal RMS (ERMS) and alveolar RMS (ARMS). ERMS features small spindle-shaped cells and genetic instability with increased expression of IGF-II (Zhan et al. 1994). ARMS exhibits a small and rounded cell shape reminiscent of pulmonary alveoli (Newton 1995). The majority of ARMS tumors have chromosome translocations, t(2;13)(q35;q14) or t(1;13)(p36;q14), which result in PAX3-FKHR or PAX7- FKHR fusion proteins. These proteins fuse the DNA binding domain of PAX3/PAX7 and the transactivation domain of the forkhead family of transcription factors (FKHR) to induce inappropriate gene transcription (Galili et al. 1993; Fredericks et al. 1995). ARMS tumors tend to grow faster than ERMS tumors and usually require more intensive treatment (Kelly et al. 1997). The RD cell line (ATCC ® Catalog No. CCL-136 TM ) is derived from the rhabdomyosarcoma (RMS) of a seven-year old Caucasian girl. RMS is the most common soft tissue sarcoma of childhood, and a majority of RMS patients are under 18 years of age. Two 114 histologic subtypes of this tumor are embryonal RMS (60% frequency) and alveolar RMS (20% frequency) (Hynes 2002). To determine if high BCAR3 and Cas expression was common in other RMS cell lines and shared by both RMS subtypes, I further examined a panel of RMS cell lines representing alveolar RMS (CW9019, Rh18, and RH30) and embryonal RMS (RD and A204) for the expression of PTPα, BCAR3, and Cas. MEF cell lysate was used as a reference sample and equivalent loading amounts of lysates from different cell lines was confirmed by probing for actin (Fig. 5.1B). The cell lines express varying amounts of PTPα that were not consistently associated with tumor subtype, and the slight size shifts of the PTPα band that were observed in some cell lines may be due to different levels of glycosylation in the extracellular domain of PTPα (Tonks 2006) (Fig. 5.1B). The expression of Cas in all the cell lines is similar, whereas the expression of BCAR3 is very different. The CW9019 and RD cells exhibited highest BCAR3 expression, the Rh18 cells have intermediate BCAR3 expression, and the Rh30 cells display low BCAR3 expression that is comparable to MEFs. Little to no BCAR3 was detected in A204 cells (Fig. 5.1B). As with PTPα, BCAR3 and Cas expression levels did not vary in association with alveolar and embryonal subtypes. Since RD cells have a relatively high level expression of BCAR3 and Cas, together with intermediate level PTPα expression, I decided to use the RD cell line as a model system for further investigation. 5.3 PTPα, BCAR3, and Cas form a complex in RD cells To determine if PTPα, BCAR3, and Cas form a complex in the RD tumor cells as in MEFs, Cas was immunoprecipitated from RD cell lysates and the presence of associated PTPα and BCAR3, as well as Src, was determined by immunoblotting. Preliminary 115 experiments (not shown) revealed that these proteins were all present, indicating their association with Cas. In subsequent experiments, these findings were confirmed and extended by examining Cas immunoprecipitates prepared from RD cells depleted of PTPα or BCAR3 using siRNA-mediated silencing (Fig. 5.2A). A high knockdown efficiency of PTPα and BCAR3 was achieved (>95%) and probing the cell lysates confirmed that Src, Cas, and actin expression was equivalent in all three siRNA-treated groups of RD cells (Fig. 5.2B). Cas immunoprecipitates prepared from the cells were probed for PTPα, BCAR3, and Src. Cas was associated with PTPα, BCAR3, and Src in the control siRNA treated cells, confirming that the PTPα-BCAR3-Cas-Src module indeed exists in the RD cells (Fig. 5.2A, lane 1). Cas was associated with BCAR3 and Src in the PTPα knockdown cells, confirming that BCAR3-Cas-Src association is independent of PTPα (Fig. 5.2A, lane 2). BCAR3 was not immunoprecipitated with Cas in the BCAR3 knockdown cells. Importantly, no PTPα was present in Cas immunoprecipitates from BCAR3 depleted cells (Fig. 5.2A, lane 3), confirming that BCAR3 is a requisite intermediary protein for PTPα and Cas association. The above results established the PTPα-BCAR3-Cas-Src complex to be a molecular module that forms in RD cells as well as MEFs. I next investigated if this module regulates RD cell migration and invasion. 5.4 RD cell migration and invasion are dependent on PTPα and PTPα Tyr789 PhosphoTyr789 of PTPα mediates its interaction with BCAR3-Cas. I therefore investigated whether PTPα, and most importantly, PTPα Tyr789, are required for mediating RD cell migration and invasion. To accomplish this I used siRNA-mediated silencing 116 combined with an RNAi-resistant PTPα expression system developed in our lab. The RD cells were transfected with PTPα targeted siRNA to deplete endogenous PTPα. Some cells were co-transfected with siRNA resistant wildtype (WT) PTPα and mutant Y789F PTPα plasmids. Resistance to the PTPα targeted siRNA (two siRNA oligos) is conferred by mutating 2-3 base pairs in each of the regions of PTPα mRNA that is the targeted by the siRNAs. As shown in Fig. 5.3A (lanes 1 and 2) the siRNA treatment ablated >95% of PTPα in RD cells, and re-expression of RNAi-resistant WT or Y789F PTPα at physiological levels was confirmed by immunoblotting for PTPα and PTPα phosphoTyr789 (lanes 3 and 4). Migration was assessed by microscopic imaging (OlympusIX81 system) of cell movement into a created “wound” or empty space. The BioFlux200 system was used because of its ability to generate uniform 200μm standard wounds for different cell groups by controlled flow trypsinization-mediated cell removal. Cells depleted of PTPα showed a significantly reduced migration ability (Fig. 5.3B), indicating that PTPα is required for RD cell migration. The re-expression of WT PTPα in PTPα depleted cells rescued the defective cell migration caused by PTPα knockdown (Fig. 5.3B). However, the expression of Y789F PTPα failed to rescue the defective cell migration in PTPα depleted cells (Fig. 5.3B). The same experimental approach was used to investigate the roles of PTPα and PTPα Tyr789 in RD cell invasion. Invasion was determined using Boyden chambers coated with a layer of Matrigel (BioCoatTM BD MatrigelTM Invasion Chamber), since the ability of malignant cells to move through Matrigel represents their invasiveness. PTPα depletion inhibited the ability of RD cells to invade by nearly 90% (12±2.3% invasion index of PTPα depleted cells compared to non-targeted control siRNA treated RD cells) (Fig. 5.4). Re- 117 constitution of WT PTPα expression rescued this invasion defect (99±12% invasion index of PTPα depleted cells expressing WT PTPα compared to non-targeted control siRNA treated RD cells), but RD cells expressing Y789F PTPα still showed significantly reduced invasion (18±2.5% invasion index of PTPα depleted cells expressing Y789F PTPα compared to non- targeted control siRNA treated RD cells) (Fig. 5.4). The above results demonstrated that PTPα and Tyr789 are required for RD cell migration and invasion. Given the essential role of PTPα Tyr789 in formation of the PTPα-BCAR3-Cas complex, this suggests that this intact complex may be a positive regulator of RD cancer cell migration and invasion. 5.5 BCAR3 is required for RD cell migration and invasion The siRNA-mediated depletion of BCAR3 in BT-549 cells inhibits migration and invasion by 50% or more (Schrecengost et al. 2007). Whether or not BCAR3-mediated migration and invasion are dependent upon the ability of BCAR3 to associate with Cas and/or PTPα is not known. In support of the BCAR3-Cas interaction being involved in BCAR3-mediated migration and invasion, the depletion of BCAR3 from BT-549 cells results in the absence of Cas at the cell membrane (Schrecengost et al. 2007). In the work described in Chapter 4, I demonstrated that in MEFs, mutant BCAR3 with a GEF domain point mutation (R748A) cannot interact with Cas, and mutant BCAR3 with an SH2 point mutation (R177K) cannot interact with phospho-Tyr789-PTPα. Here I investigated if BCAR3 was required for RD cell invasion, and if the PTPα-interacting SH2 domain of BCAR3 and the Cas-interacting GEF domain of BCAR3 played a role in supporting this ability. 118 5.5.1 BCAR3 silencing reduces RD cell invasion To investigate whether RD cell invasion requires BCAR3, RD cells were depleted of BCAR3 using siRNA-mediated silencing, or treated with non-targeting siRNA as a control. More than 95% of BCAR3 was depleted from RD cells by this treatment (Fig. 5.5A). Invasion assays showed that BCAR3 depletion resulted in significantly reduced RD cell invasion (17±1.6% invasion index of BCAR3 depleted cells compared to non-targeted control siRNA treated RD cells) (Fig. 5.5B), confirming that BCAR3 is required for this property of RD cells. 5.5.2 BCAR3 SH2 mutant and GEF mutant inhibit RD cell invasion To investigate if the role of BCAR3 as PTPα-Cas molecular linker is important for RD cell invasion, I reasoned that the expression of mutant BCAR3 lacking the ability to bind to either PTPα or Cas might compete with endogenous BCAR3 to disrupt the endogenous PTPα-BCAR3-Cas complex. To this end, FLAG-tagged WT BCAR3 or mutant BCAR3 with inactivating point mutations in the SH2 domain (R177K) or the GEF domain (R748A) were overexpressed in RD cells (Fig. 5.6A). The expression of WT BCAR3 had little effect on RD cell invasive ability (90±5.1% invasion index of WT BCAR3-overexpressing RD cells compared to parental non-transfected RD cells) (Fig. 5.6B), probably due to the high levels of endogenous BCAR3 and Cas in these cells that support maximal invasion. However the invasive properties of the cells were significantly inhibited by the overexpression of SH2 and GEF mutants of BCAR3 (53±5.2% invasion index of by SH2 mutant BCAR3-overexpressing 119 RD cells and 52±2.1% invasion index of GEF mutant BCAR3-overexpressing RD cells compared to parental non-transfected RD cells) (Fig. 5.6B). These results suggested that the mutant non-interactive forms of BCAR3 indeed exert dominant-negative effects and that intact BCAR3 is required for RD cell invasion. 5.6 Discussion My findings described in previous chapters showed that the PTPα-BCAR3-Cas complex mediates Cas localization to FAs and Cas downstream signaling. The integrin-stimulated phosphorylation of PTPα Tyr789, intact SH2 domain and GEF domain of BCAR3 are all required for transducing the signal via Cas to promote migration. Disruption of BCAR3-Cas interaction blocks the Cas-mediated signaling cascade that is essential for cell migration (Bouton et al. 2001). If this complex is a common module that also functions as a positive regulator of cell motility in cancer cells, it represents a potential target for cancer therapy. Since BCAR3 is highly expressed in aggressive ER- (estrogen receptor negative) breast cancer cell lines (van Agthoven et al. 1998; Cai et al. 2003), the role of BCAR3 has for most part been studied in breast cancer context. To expand the knowledge of BCAR3 functions in other types of cells, I initially examined BCAR3 expression in two pediatric cancer cell lines. My results showed that RD cells (RMS) have high expression of BCAR3, coupled with high expression of Cas and readily detectable expression of PTPα. High BCAR3 expression was detected in one other type (CW9019) of the 5 RMS cells tested. Since RD cells have the highest combined expression of these three proteins, I decided to use RD cells as a model system to investigate the formation and role of the PTPα, BCAR3, and Cas complex. 120 Consistent with the results from MEF cell studies described in Chapter 4, PTPα, BCAR3, and Src form a complex with Cas in RD cells, and depletion of BCAR3 disrupts this complex, confirming the existence of this molecular module in RD cancer cells and that PTPα association with Cas is indeed mediated by BCAR3. PTPα silencing does not affect the interaction between Cas and BCAR3, though the Cas-associated Src is reduced in the absence of PTPα. These findings suggest that the PTPα-BCAR3-Cas-Src signaling complex may play a role similar to that in MEFs to regulate RD cell migration. Since these are cancer cells, it is also important to examine if this module can regulate some of the motility-related cancer properties, such as invasive ability. Previous studies indicate that PTPα or BCAR3 functions as positive regulator of cell migration and invasion in certain cancer cell lines. RNAi-mediated PTPα knockdown reduces Src activity by 2 to 4-fold in breast, colon and other human cancer cell lines (Zheng et al. 2008), and Src activity is correlated with cancer cell invasiveness in various tumors (Guarino 2010). Prostate cancer cells depleted of Src display reduced cell migration and treatment of prostate cancer or sarcoma cell lines with the small molecule SFK kinase inhibitor dasatinib in also results in decreased cellular proliferation, migration, and invasion (Shor et al. 2007; Park et al. 2008). BCAR3 depletion in BT-549 breast cancer cells inhibits cell migration and invasion while overexpression of BCAR3 in MCF7 breast cancer cells increases cell migration and invasion (Near et al. 2007; Schrecengost et al. 2007). In RD cells, I found that PTPα knockdown severely inhibits cell migration and this migration defect can be rescued by the expression of WT PTPα, confirming that this effect on cell motility is caused by the depletion of PTPα. The role of PTPα Tyr789 in cell migration has been shown in MEFs, where the expression of Y789F mutant PTPα is unable to restore the defective cell 121 migration of PTPα-/- MEFs (Chen et al. 2006). My studies establish that expression of siRNA-resistant Y789F PTPα in PTPα-depleted RD cells likewise fails to rescue impaired migration, implicating Tyr789 of PTPα as essential for the motility of RD cells. Acquisition of invasiveness and metastatic ability is a hallmark of cancer cells. The gain of these properties includes two key events: the activation of extracellular proteases (metalloproteases, serine proteases, and cysteine proteases) and the altered binding specificities of adhesion molecules (cadherins, CAMs, and integrins) (Hanahan and Weinberg 2000; Cudic and Fields 2009). The regulatory molecular mechanisms that are involved in this are still unclear. In this chapter, I presented the first evidence for a role of the PTPα-BCAR3-Cas-Src signaling complex in regulating invasion of RD cells. Similar to migration, PTPα and its Tyr789 residue are required for RD cell invasion, indicating the importance of PTPα Tyr789 in regulating RD cancer cell invasion. BCAR3 silencing in RD cells also significantly impaired the invasive ability of these cells, expanding a role for BCAR3 in non-breast cancer cell invasion. To determine whether the defective invasion is caused by disruption of the PTPα-BCAR3-Cas complex, I further examined RD cells overexpressing SH2 mutant BCAR3 that cannot interact with Cas or GEF mutant BCAR3 that cannot interact with PTPα phosphoTyr789. Interestingly, cells expressing either mutant showed reduced invasive ability, suggesting that mutant BCAR3 can compete with endogenous BCAR3 and disrupt the PTPα-BCAR3-Cas complex to act in dominant-negative manners. These results indicate that the PTPα-BCAR3-Cas-Src signaling complex is a positive regulator of RD cell invasion, which likely requires events additional to those involved in cell migration to remodel the ECM. It will be very interesting to identify the downstream effectors and pathways involved in this process. 122 In summary, I found that BCAR3 and Cas are highly expressed in RD cells, a cell line derived from an embryonal rhabdomyosarcoma (eRMS). I demonstrated that PTPα, PTPα Tyr789, and intact wildtype BCAR3 are all required for RD cell migration and invasion, indicating a new role for the PTPα-BCAR3-Cas complex in regulating cancer cell invasion. This information is relevant to personalized screening of tumors for PTPα, BCAR3, and Cas expression to determine if the tumor is likely to metastasize, and may warrant the development of novel therapeutics that disrupt the complex to inhibit cancer metastasis. PTPα +/+ ‐/‐ MEFs Alveolar RMS Embryonal RMS BCAR3 Cas actin PTPα pY789 PTPα Cas BCAR3 actin A B Figure 5.1 Expression of PTPα, BCAR3, and Cas in cancer cell lines Lysates from wild type MEFs and different types of cancer cell lines were prepared for immunoblotting. (A) The indicated cell lysates were probed for PTPα phospho- Tyr789, PTPα, Cas, BCAR3 and actin. BT-549, breast cancer cells; RD, rhabdo- myosarcoma (RMS) cells; TC-32, Ewing’s sarcoma cells. (B) Lysates from wild type (+/+) and PTPα-/- (-/-) MEFs, and different types of RMS cell lines representing alveolar and embryonal subtypes were probed for PTPα, Cas, BCAR3 and actin. 123 actin PTPα BCAR3 Src Cas Lysate + IP Cas Control: PTPα: BCAR3: + + PTPα BCAR3 Src Cas siRNA Figure 5.2 PTPα, BCAR3, and Cas form a complex in RD cells that is BCAR3-dependent. RD cells were transfected with control non-targeting (50nM), PTPα-targeted (20nM) or BCAR3-targeted (50nM) siRNA. After 48 hours, lysates were prepared and used for Cas immuno-precipitation. (A) Immunoprecipitates were probed for PTPα, BCAR3, Src, and Cas. (B) Cell lysates were probed for the indicated proteins. 124 1 2 3 A B Figure 5.3 RD cell migration is dependent on PTPα and PTPα Tyr789. RD cells were transfected with 20nM control non-targeting (Ctrl) or PTPα-targeted siRNA (PTPα KD), or co-transfected with PTPα-targeted siRNA and RNAi-resistant WT PTPα (WTα) or Y789F PTPα (Y789Fα) as indicated. (A) The cells were lysed 48 hours later and 30 μg whole cell lysate was immunoblotted for the indicated proteins. (B) After 24 hours, the cells were trypsinized and replated into FN-coated channels of BioFlux 24-well plate. The plate was incubated at 37ᵒC for 24 hours to allow the cells to attach the bottom of each channel. The cells on the right half of each channel were then removed by trypsinization using the BioFlux200 system (top panels, 0h: cells immediately after trypsinization). Other cells were incubated at 37ᵒC for 24 hours following trypsinization to allow the cells migrate into the cleared right hand side of the channel (bottom panels, 24h). At 0h and 24h, cells were stained with Calcein AM and imaged using the Olympus IX81system equipped with a CoolSnap HQ2 CCD camera. A B Ctrl PTPα KD 0h 24h WTα Y789Fα 100μm PTPα siRNA: Ctrl siRNA: PTPα actin PTPα pY789 PTPα: WT Y789F ++ + + 125 1 2 3 4 Ctrl PTPα KD WTα Y789Fα 500μm 1.0 0.8 0.6 0.4 0.2 Ctrl αKD WTα Y789Fα In va sio n In de x * **1.2 126 Figure 5.4 PTPα and PTPα Tyr789 are required for the invasion of RD cells. RD cells were transfected with 20nM control non-targeting (Ctrl) or PTPα-targeted siRNA (PTPα KD), or co-transfected with PTPα-targeted siRNA and RNAi resistant WT PTPα (WTα) or Y789F PTPα (Y789Fα) as indicated for 24 hours. The cells were trypsinized and seeded into invasion chambers (6×104 cells/chamber). After incubating at 37°C for 24 hours, the Matrigel and cells on the top of the membrane in each chamber were removed using cotton swabs. (A) The cells remaining on the bottom of the membrane were stained with Calcein AM , and scanned using an Olympus FV10i microscope. (B) The numbers of invaded cells present at the bottom of each chamber membrane were counted using ImageJ software and confirmed by manual counting. Data from 3 independent experiments are shown in the graph as mean±SD, and asterisks indicate significant differences (*, p=7×10-7; **, p=1×10-6). AB PTPα actin BCAR3 : + +Ctrl: BCAR3 Ctrl BCAR3 KD 500μm Figure 5.5 BCAR3 is required for RD cell invasion. RD cells were transfected with 50nM control non-targeting (Ctrl) or BCAR3- targeted siRNA (BCAR3 KD). (A) The cells were lysed 48 hours later and 30 μg whole cell lysate was immunoblotted for the indicated proteins. (B) After 24h of transfection, the cells were trypsinized and seeded into invasion chambers (6×104 cells/chamber) . After incubating at 37°C for 24 hours, the Matrigel and cells on the top of the membrane in each chamber were removed using cotton swabs. The cells remaining on the bottom of the membrane were stained with Calcein AM , and scanned using an Olympus FV10i microscope. The numbers of invaded cells present at the bottom of each chamber membrane were counted using ImageJ software and confirmed by manual counting. Data from 3 independent experiments are shown in the graph as mean ± SD, and the asterisk indicates a significant difference (*, p=1×10-7). 127 1.0 0.8 0.6 0.4 0.2 Ctrl BCAR3 KD In va sio n In de x * siRNA AB Figure 5.6 An intact BCAR3 SH2 domain or GEF domain is required for RD cell invasion. RD cells were transfected with FLAG-tagged wildtype BCAR3 (WT) or mutant forms of BCAR3 (SH2, R177K mutation in SH2 domain; GEF, R748A mutation in GEF domain). (A) The cells and control non-transfected parental cells (Ctrl) were lysed 48 hours later and 30 μg whole cell lysate was immunoblotted for the indicated proteins. (B) After 24h of transfection, the cells were trypsinized and seeded into invasion chambers (6×104 cells/chamber). After incubating at 37°C for 24 hours, the Matrigel and cells on the top of the membrane in each chamber were removed using cotton swabs. The cells remaining on the bottom of the membrane were stained with Calcein AM , and scanned using an Olympus FV10i microscope. The numbers of invaded cells present at the bottom of each chamber membrane were counted using ImageJ software and confirmed by manual counting. Data from 3 independent experiments are shown in the graph as mean±SD, and the asterisks indicate significant differences (*, p=1×10-6; **, p=3×10-4). 128 * ** 1.0 0.8 0.6 0.4 0.2In va sio n In de x WT SH2 GEFCtrl WT SH2 GEF 500μm Ctrl PTPα WT SH2 GEF FLAG BCAR3 actin Ctrl 129 Chapter 6: Summary and Future Directions 6.1 The novel PTPα-BCAR3-Cas-Src signaling module in integrin signaling My thesis describes a novel non-catalytic mechanism of action of PTPα in integrin- regulated cell migration through PTPα-BCAR3-Cas-Src signaling complex, which forms in response to integrin-stimulated phosphorylation of PTPα Tyr789. This complex mediates Cas translocation to FAs, where it undergoes tyrosine phosphorylation to activate downstream integrin signaling events that promotes cell migration. Furthermore, I extended this study into a cancer model by showing that disruption of this signaling complex inhibits the invasive motility of RD rhabdomyosarcoma cells. PTPα was cloned by several groups in 1990 (Kaplan et al. 1990; Krueger et al. 1990; Matthews et al. 1990; Sap et al. 1990). Shortly after that, it was identified as a positive regulator of Src, dephosphorylating Src at the inhibitory Tyr527 residue to activate it (Zheng et al. 1992). In 1994, den Hertog and his colleagues identified a constitutively phosphorylated tyrosine at the C-terminus of PTPα and mapped it to Tyr789 (den Hertog et al. 1994). Previous work in our lab demonstrated two roles of PTPα in integrin signaling: 1) PTPα activates SFKs (and FAK) in a manner dependent on its catalytic activity but independent of PTPα Tyr789 phosphorylation and 2) promotes downstream cytoskeleton- associated events such as cell spreading and migration in a manner that requires PTPα phosphorylation at Tyr789 (Chen et al. 2006) (Fig. 6.1). My study focuses on this second role of PTPα, and my findings in Chapter 3 clearly showed that Cas tyrosine phosphorylation is defective and the Cas/Crk-Rac/Cdc42-PAK signaling cascade is severely impaired in the absence of this single Tyr789 residue in PTPα. The key role played by tyrosine 130 phosphorylated Cas in promoting cell migration strongly indicates that these molecular defects account for the reduced cell spreading and migration previously linked to mutated Tyr789 in PTPα (Chen et al. 2006). I further demonstrated in Chapter 4 that the phosphoTyr789 site functions as a docking site for Cas recruitment to newly formed FA complexes through a molecular linker known as BCAR3. The discovery of this PTPα- phosphoTyr789-BCAR3-Cas signaling module further revealed the specific mechanism of action underlying the second role of PTPα (Fig. 6.1), which is essential for integrin- stimulated cell spreading and migration. Cell migration is a process that occurs in response to extracellular chemical signals and mechanical forces through the temporal and spatial organization of signaling complexes and the cytoskeleton (Lauffenburger and Horwitz 1996; Hood et al. 2003; Gardel et al. 2010). Integrin ligation initiates a hierarchy of recruitment of signaling proteins and cytoskeletal molecules to FAs, which includes FAK, Src, PTPα, BCAR3, Cas, actin and many other molecules. (Larsen et al. 2003; Parson et al. 2010; Nagano et al. 2012). Interestingly, Cas was proposed to act as a mechanical force sensor in the cell since mechanical extension of the Cas SD has been shown to promote its tyrosine phosphorylation and force-dependent activation of the small GTPase Rap1 (Sawada et al. 2006). Based on this extension model, theoretically Cas needs at least two anchor points for the applied mechanic force to stretch its SD and increase the accessibility of the region to phosphorylation. The N-terminal SH3 domain of Cas associates with FAK, a protein that contains multiple cytoskeletal-associated domains, such as an actin-binding FERM domain (Chen et al. 1995; Lee et al. 2004) and a paxillin-binding FAT domain (Tachibana et al. 1995). Thus FAK can recruit Cas to FAs as well as mediate the N-terminal anchorage of Cas (Burnham et al. 1996; Sawada et al. 2006). 131 Although the C-terminal CCH (Cas family C-terminal homology domain) region of Cas is required for optimal FA localization and Cas phosphorylation (Donato et al. 2010), so far no other anchor mechanism for this region of Cas has been identified. My results support a model (Fig. 6.2) that may provide a mechanism for the C-terminal anchorage of Cas. Integrin activation induces Src-mediated phosphorylation of PTPα at Tyr789, resulting in the formation and localization of the PTPα-BCAR3-Cas complex at nascent adhesion sites. BCAR3 connects PTPα and Cas through the BCAR3 N-terminal SH2 domain association with phosphoTyr789 of PTPα and its C-terminal GEF domain interaction with the CCH region of Cas (Gotoh et al. 2000; Donato et al. 2010). PTPα is also required for force- dependent FA formation by associating with αvβ3 integrins during the initial phase of cell- ECM adhesion (von Wichert et al. 2003). Since integrin-stimulated assembly of the PTPα phosphoTyr789-BCAR3-Cas-Src complex recruits Cas to FAs and places Cas in proximity to Src, I propose that this mechanism provides a C-terminal anchor point for Cas at FAs. In my model (Fig. 6.2), the PTPα-BCAR3 C-terminal Cas anchor cooperates with FAK-mediated N-terminal anchoring of Cas to provide opposing traction that allows the mechanical force- dependent conformational extension of Cas to expose its SD for phosphorylation by Src (Sawada et al. 2006). My findings also demonstrate a new level of functional interdependency between PTPα and Src in integrin signaling as not only does PTPα dephosphorylate and activate Src (Zeng et al. 2003; Chen et al. 2006), but Src-mediated phosphorylation of PTPα also promotes the accessibility of Cas as a substrate for Src. Integrin engagement induces the auto- phosphorylation of FAK at Tyr397 site, which creates a binding site for the SH2 domain of Src (Parsons 2003). The interaction of the Src SH2 domain with FAK phosphoTyr397 and 132 the association of the Src SH3 domain with the integrin β subunit disrupt the inhibitory conformation of Src and increase the accessibility of the Src C-terminal phosphoTyr527. Integrin-associated PTPα dephosphorylates this phosphotyrosine and facilitates the optimal activation of Src (the early upstream role of PTPα, role 1). The active FAK-associated Src phosphorylates FAK at Tyr576/Tyr577 to activate FAK kinase activity and promote further FAK Tyr397 auto-phosphorylation, resulting in the full activation of the Src-FAK complex (Zeng et al. 2003). The active Src-FAK complexes in turn phosphorylate PTPα Tyr789 and downstream scaffolding proteins such as Cas and paxillin (Chen et al. 2006). The phosphoTyr789 residue of PTPα cooperatively transduces a signal downstream via the PTPα- BCAR3-Cas-Src signaling module to promote cytoskeletal rearrangement, FA formation, and cell migration. Besides the role downstream of PTPα phosphoTyr789, BCAR3 has also been shown to mediate Src activation. It was proposed that BCAR3-mediated Src-Cas association at FAs may promote full activation of Src (Schuh et al. 2010), and I confirmed that BCAR3 depleted cells indeed display reduced Src activation. In the present study I demonstrated that BCAR3 is required for Src Tyr416 and PTPα Tyr789 phosphorylations to promote BCAR3- mediated Cas localization to FAs and Cas tyrosine phosphorylation, raising the possibility that BCAR3 may operate upstream as well as downstream of PTPα. Interestingly PTPα and BCAR3 clearly target different regulatory tyrosine residues in Src and the absence of either of these proteins leads to defective Src activation, suggesting that the roles and interactions of PTPα and BCAR3 in Src activation merit further investigation. My experimental evidences of integrin-stimulated PTPα-BCAR3 association result from biochemical experiments in vitro, demonstrating that the SH2 domain of BCAR3 is able to interact with PTPα phosphoTyr789. My colleague, Suzanne Cheng, further confirmed that 133 PTPα Tyr789 is required for FA localization of Cas and BCAR3 using total internal reflection fluorescence (TIRF) microscopy (Sun et al. 2012). It would be ideal to visualize the complex formation of PTPα, BCAR3 and Cas at FAs in vivo using fluorescent tagged fusion proteins. Specifically, GFP-tagged PTPα (WT and Y789F) and mCherry-tagged BCAR3 could be used to examine the integrin-stimulated co-localization of PTPα with BCAR3 and whether Tyr789 of PTPα is required for their FA translocation by TIRF microscopy. YFP-tagged PTPα (WT and Y789F) and CFP-tagged BCAR3 may also be used to examine the interaction of PTPα and BCAR3 in live cells using fluorescence resonance energy transfer (FRET). Furthermore, we could use YFP-tagged PTPα, mCherry-tagged BCAR3, and GFP-tagged Cas to reveal their real-time translocation in the cell by fluorescence recovery after photobleaching (FRAP). The above experiments would provide more detailed information about the temporal and spatial regulation of PTPα-BCAR3-Cas complex formation in vivo, allowing us a better understanding of how these molecules cooperate to promote cell migration. However, the major current limitation putting this experimental approach into practice is the difficulty in appropriately placing the florescent tag on PTPα. An N-terminal tag cannot be used because N-terminus of PTPα is the signal peptide, and placing the tag at C-terminus prevents the tyrosine phosphorylation of PTPα (Suzanne Cheng unpublished result). Additionally we could try to visualize endogenous PTPα, but we do not have antibody with sufficient quality for immunofluorescent detection. 6.2 PTPα-BCAR3-Cas-Src signaling in promoting cancer cell invasive motility As a ubiquitously expressed scaffolding protein, Cas integrates signals from growth factor receptors and integrins to control cell migration and survival (Stupack et al. 2000; Defilippi 134 et al. 2006). Overexpression and hyper-phosphorylation of Cas are involved in driving cancer cell migration and invasion as well as promoting cell proliferation and drug resistance (Brinkman et al. 2000; van der Flier et al. 2000; Ta et al. 2008). Cas and BCAR3 were initially identified as genes responsible for conferring anti-estrogen (tamoxifen) resistance to breast cancer cells (Dorssers et al. 1993; van Agthoven et al. 1998), and expression of exogenous Cas in colon cancer cells confers resistance to COX-2 inhibitor celecoxib-induced cell death (Casanova et al. 2006). Ionizing radiation strongly induces Cas expression and tyrosine phosphorylation, which are essential for post-radiation cell survival in lung carcinoma cells and fibroblasts (Beinke et al. 2003; Seidler et al. 2005). Cas overexpression can abolish the PTEN-induced inhibition of glioblastoma cell spreading and migration (Tamura et al. 1999). Cas is often overexpressed in association with enhanced metastasis and poor prognosis in breast cancer (Cabodi et al. 2010; Tikhmyanova et al. 2010). Furthermore the high expression of Cas and BCAR3 correlates with more aggressive and invasive breast cancer cell lines (Near et al. 2007; Schrecengost et al. 2007; Makkinje et al. 2009). The results described in Chapter 5 showed that the PTPα-BCAR3-Cas signaling module promotes the invasion of RD rhabdomyosarcoma cells in a PTPα Tyr789-, BCAR3 SH2/GEF domain- dependent manner, indicating their involvement in regulating cancer cell invasion. Furthermore, the RD cells and PTPα depleted RD cells, and PTPα depleted RD cells re- expressing siRNA resistant wildtype (WT) PTPα and mutant Y789F PTPα could be injected into immunodeficient mice to examine their ability for xenograft tumor formation Experimental evidence from these in vivo xenograft assays in animal model will be necessary to further examine the role of the PTPα-BCAR3-Cas complex in promoting invasive cancer properties, such as metastasis. 135 Degradation of the ECM is one of the key events that occurs during cancer invasion and metastasis (van Zijl et al. 2011). As an extraordinary complicated process, cancer metastasis encompasses 1) local invasion of primary tumor cells through basement membrane into adjacent tissues, 2) intravasation of tumor cells into blood or lymphatic vessels and survival in the circulatory system, 3) tumor cells arrest and extravasation, and 4) colonization and outgrowth of tumor cells at the distal secondary sites (Hunter et al. 2008; Bozzuto et al. 2010). ECM degradation mediated by specialized organelles termed invadopodia and is involved in all the steps above (Linder 2007; Buccione et al. 2009; Caldieri and Buccione 2010). Based on the similarities between invadopodia and FAs in composition and architecture, it is proposed that invadopodia may develop from FAs (Linder and Aepfelbacher 2003; Buccione et al. 2004). Interestingly, a novel activity of FAs to degrade the ECM and thus initiate cancer cell invasion has been recently described (Wang and McNiven 2012). This process is mediated by the regulated interaction of the matrix metalloproteinase MT1-MMP with Cas at FAs. MT-MMPs are synthesized as inactive zymogens (pro-MT-MMP), which are activated in the Golgi by convertase-mediated proteolytic removal of the pro-peptide (Holmbeck et al. 2004). MT1-MMP has been reported to promote cancer invasion either directly by degrading the ECM components or indirectly by activating other MMPs (Seiki 1999). Disruption of the Cas-MT1-MMP interaction impairs tumor cell invasion (Wang and McNiven 2012). The MT1-MMP-Cas interaction requires phosphorylation of MT1-MMP and the Cas SD by Src. Here I postulate that since PTPα phosphoTyr789 recruits Cas into proximity with Src at FAs (via the PTPα-BCAR3-Cas association), this not only enables Src to phosphorylate Cas and MT1-MMP, but also induces the recruitment of the active MT1-MMP to FAs. This would facilitate MT1-MMP-mediated 136 ECM degradation at FAs to promote tumor cell invasion. It will be important to investigate the role of PTPα Tyr789 and the PTPα-BCAR3-Cas complex in regulating MT1-MMP and its FA-localized matrix degradation activity. 6.3 The PTPα phosphoTyr789 binding proteins When Tony Hunter and his colleagues identified the phosphorylation of Tyr789 in PTPα, they proposed that phosphoTyr789 functions as a docking site for SH2 domain-containing proteins and identified its first binding partner -the well-known adapter protein Grb2 (den Hertog et al. 1994). Besides the tight association of PTPα phosphoTyr789 with the Grb2 SH2 domain, the same group later showed that the C-terminal SH3 domain of Grb2 also interacts with the D1 domain of PTPα and blocks its catalytic cleft, inhibiting the phosphatase activity of PTPα (den Hertog et al. 1996). Src was also identified as an associated protein of PTPα phosphoTyr789 (Zheng et al. 2000). My investigation identifies phosphoTyr789 PTPα as the first cellular ligand for the SH2 domain of BCAR3 by demonstrating that not only does the formation of this signaling complex require PTPα Tyr789 and an intact SH2 domain of BCAR3, but also that recombinant BCAR3 SH2 domain interacts directly with WT PTPα and not with Y789F PTPα. The ability of the BCAR3 SH2 domain to bind to PTPα phosphoTyr789 was unexpected as this SH2 domain is not closely related to either of the SH2 domain in Grb2 and Src (Liu et al. 2006) that also can bind to phosphoTyr789. However, virtually nothing is known of BCAR3-SH2 specificity and besides PTPα, no other target phosphotyrosyl proteins have been found. BCAR3 (also known as NSP2, Shep2, or AND-34) (van Agthoven et al. 1998; Cai et al. 1999) is a member of an SH2 domain- containing family of Cas-binding proteins that includes NSP1 (Lu et al. 1999) and Shep1 137 (also known as NSP3 or Chat) (Dodelet et al. 1999; Sakakibara and Hattori 2000). Nsp1 has been shown to interact with activated EGF receptor through an unidentified mechanism (Lu et al. 1999), and the SH2 domain of Shep1 can bind to tyrosine phosphorylated EphB2 and EphA4 receptors (Dodelet et al. 1999). These binding partners are all receptor tyrosine kinases, and my findings demonstrate that BCAR3 can interact with a receptor protein tyrosine phosphatase, PTPα. This raises the possibility that the SH2 domains of NSP family members have distinct binding specificities. Functionally, BCAR3 has been shown to induce anti-estrogen resistance while other NSP homologs do not have this ability, suggesting that three members of NSP family may regulate different effectors though they all have similar structures (Near et al. 2009). To begin to investigate if these functional differences are regulated by the distinct binding specificity of their SH2 domain, it is important to identify SH2 domain binding partners. To do this, the GST-tagged SH2 domain of each NSP could be used as the bait for pull-down experiments, and tagged full length NSPs could be useful to identify other binding proteins, including direct downstream effectors. If suitable specific NSP1 and Shep1 antibodies are available, it will be interesting to examine if NSP1 and Shep1 can also interact with phosphoTyr789 of PTPα and/or regulate cell migration. The SH2 domain of Shep1 has been implicated to bind Tyr605 and Tyr611 in the juxtamembrane region of EphB2, and mutation of these tyrosines to Phe or Glu indicated that Tyr605 is the SH2 binding site (Dodelet et al. 1999; Zisch et al. 2000). Alignment of the PTPα tail sequence with this region of EphB2 reveals a similar positioning of two PTPα tyrosine residues, Tyr789 and Tyr782 (Fig. 6.3). Moreover, the sequences on the C-terminal sides of PTPα Tyr782 and EphB2 Tyr605 are highly homologous, suggesting that these represent respective recognition motifs for the related SH2 domains (Lu et al. 1999) of 138 BCAR3 and Shep1. The binding of Shep1-SH2 to phosphoTyr605-EphB2 at the start of this potential recognition motif raises the interesting question of whether Tyr782 of PTPα plays any role in binding to the SH2 domain of BCAR3, although this site in PTPα has never been shown to be phosphorylated. To investigate the requirement of Tyr782 for the interaction of PTPα and BCAR3, I constructed VSVG-tagged Y782F and Y782/789F double mutant PTPα plasmids. Lysates were prepared from PTPα+/+ and PTPα-/- MEFs, and PTPα-/- MEFs expressing VSVG-tagged Y782F, Y789F, Y789F/Y782F, and WT PTPα (Appendix Fig.1, bottom panels), and VSVG immunoprecipitates were isolated and probed for phospho- tyrosine, BCAR3, VSVG, and PTPα. Not surprisingly, VSVG immunoprecipitates from WT and Y782F PTPα expressing cells showed detectable tyrosine phosphorylation, confirming that Tyr789 is the major phosphorylation site in PTPα (den Hertog et al. 1994). Consistent with this, BCAR3 associated with WT and Y782F PTPα that have intact Tyr789 residues, but was not detected in association with Y789F and Y782/789F mutant PTPα that both lack of Tyr789 (Appendix Fig.1, top panels). This indicates that phosphoTyr789 is indeed the binding site for the SH2 domain of BCAR3 and Tyr782 is not required for the interaction between PTPα and BCAR3. Nevertheless, the possibility that phosphorylation of Tyr789 is prerequisite for phosphorylation of Tyr782, in this way enabling the formation of a BCAR3- SH2 binding site at phosphoTyr782, cannot at present be present rule out. Thus it will be important to examine if the Y789E mutant PTPα (with the glutamic acid substitution mimicking the negative charge of the phosphotyrosine) can promote PTPα Tyr782 phosphorylation and so induce the interaction between PTPα and BCAR3. This piece of data is essential to unequivocally identify the exact tyrosine binding site of BCAR3 on PTPα. 139 6.4 Conclusion In summary, I have described a novel molecular complex of PTPα-BCAR3-Cas-Src. This complex forms in response to PTPα Tyr789 phosphorylation and mediates Cas localization to FAs and Cas downstream signaling to promote cell migration. Since Cas is a critical regulator of cell movement in pathological contexts such as cancer metastasis (Cabodi et al. 2010), my findings that the PTPα-BCAR3-Cas-Src signaling complex regulates cell migration and RD cancer cell invasive motility, suggesting that this module may be the central to the action of Cas in cancer cell invasion and metastasis. This novel information may be useful for designing molecular targeting strategies to ablate cancer metastasis. Ligands Integrin FAK‐Src Figure 6.1 Schematic diagram illustrating the dual roles of PTPα in integrin- mediated cell migration Integrin engagement by ligands induces the activation of Src and FAK. Active Src and FAK forms a complex, and phosphorylates scaffolding proteins, such as Cas and paxillin, leading to the phosphotyrosyl-dependent assembly of multi-protein signaling complexes -termed focal adhesions (FAs) (denoted by shaded area)- and transduce the signals to downstream pathways. (1) PTPα-mediated activation of SFKs; (2) active Src-FAK complex phosphorylates PTPα at Tyr789; (3) PTPα phosphoTyr789-mediated PTPα-BCAR3-Cas complex formation and Cas recruitment to FAs; (4) Src phosphorylates Cas at FAs. BCAR3 PTPα PIX PKL Rac/Cdc42 Dock180/ELMO Crk Paxillin‐P Cas‐P PAK 1 2 PTPα‐P 140 Src 3 4              !"  # $%  &'()*+,-.-/012+345'16'4(*437852+3+1249'03:;<2'12=270=27>;*?@ABC>CD177*6'945+0E40=70'5'79'9(496=270=27*;F45'79'9&/0-GHIJKLMHJHKNKJOJHIMHPQRJSTUTVWXYSWXYLZ[NIMYHNIUZL\]̂_ZNRIM̀NIJPaLRMHIXJaLRbcdeRYOW[JfgTXYSWXYUZL\]̂bTUTVMS[YRN[MhJPIYiYLOMHKcdSj_[QJNLJNkglmdnoiYLOSNOY[JRQ[NL_LMPKJ_JIpJJHWXYSWXYbTUTVNHPmNSjpMIXIXJaqrPYONMHYilmdno_MHPMHKIYWXYSWXYUZL\]̂YHTUTVNHPIXJstcPYONMHYilmdnoNSSYRMNIMHKpMIXIXJmmqLJKMYHYimNSkuIXQSWYSMIMYHMHKmNSMHcdSgUXJTUTVblmdnobPJWJHPJHImbIJLOMHN[NHRXYLMHKYimNSWLYOYIJSmNSMHIJLNRIMYHpMIXaLRNIIXJaLRb_MHPMHKPYONMHjalvkNHPaLRbOJPMNIJPWXYSWXYLZ[NIMYHjLJPNLLYpSkNHPNRIM̀NIMYHYimNSgUXMSONZ_JiNRM[MINIJP_ZcdebOJPMNIJPNHRXYLMHKYiIXJwbIJLOMHN[YimNSMHiYRN[NPXJSMYHSSYIXNIPQN[YWWYSMHKmNSNHRXYLSJHN_[JSILJIRXMHKNHPJHXNHRMHKJfWYSQLJYiIXJSQ_SILNIJPYONMHjavkLJKMYHIYaLRiYLWXYSWXYLZ[NIMYHgmNSavIZLYSMHJWXYSWXYLZ[NIMYHJHN_[JSYWIMON[PYpHSILJNOSMKHN[MHKIYWLYOYIJRJ[OMKLNIMYHg xyx z   { $| }# ~  Figure 6.3 Alignment of the homologous region in PTPα and EphB2 Amino acid sequences in the tail region of PTPα (777-793) and the juxta-membrane region of EphB2 (600-615) are aligned. The dash in the EphB2 sequence represents an introduced gap. The region of high homology is shown in green. The red circles/P depict known tyrosine phosphorylation sites. K V V Q E Y I D A F S D Y A N F K KP MG I Y I D P F T Y E D P N‐EphB2 PTPα 789 611605 782 PP P 142 143 Bibliography Ahmad, K. F. and W. A. Lim (2010) The minimal autoinhibited unit of the guanine nucleotide exchange factor intersectin. PLoS One 5, e11291 DOI: 10.1371/journal.pone.0011291 Alberts, B., Alexander Johnson, Julian Lewis, Martin Raff, Keith Roberts and P. Walter (2007) Molecular Biology of the Cell, Fifth Edition. Alexandrova, A. Y., K. Arnold, S. Schaub, J. M. Vasiliev, J. J. Meister, A. D. Bershadsky and A. B. Verkhovsky (2008) Comparative dynamics of retrograde actin flow and focal adhesions: formation of nascent adhesions triggers transition from fast to slow flow. PLoS One 3, e3234 DOI: 10.1371/journal.pone.0003234 Alonso, A., J. Sasin, N. Bottini, I. Friedberg, A. Osterman, A. Godzik, T. Hunter, J. 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Immunoprecipitates prepared from adherent cultures of wild-type (+/+) and PTPα-/- MEFs (-/-), and from PTPα-/- MEFs transfected with VSVG-tagged VSVG-Y782F, Y789F, Y782/789F, or WT PTPα plasmids were probed for phosphotyrosine (PY20), BCAR3,VSVG (PTPα), and PTPα. Lysates from the above cells were probed for PTPα phosphoTyr789, VSVG (PTPα), BCAR3, and actin. pY789 BCAR3 VSVG actin PTPα VSVG pY20 +/+ ‐/‐ ‐/‐ ‐/‐PTPα: Y782F Y782/ 789F‐/‐ ‐/‐ Y789F WTVSVG‐PTPα: VSVG‐IP Lysate BCAR3 ‐ ‐ 179 Appendix
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The role of protein tyrosine phosphatase alpha tyrosine 789 phosphorylation in integrin signaling
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