Regulation of Syk activity in GPVI-mediated activation

A Dissertation

Submitted to

The Temple University Graduate Board

In Partial Fulfillment

Of the Requirements for the Degree

DOCTOR OF PHILOSOPHY

by

Dafydd Thomas

August 2010

Dissertation Examining Committee: James L. Daniel, Pharmacology Alexander Tsygankov, Microbiology & Immunology Satya Kunapuli, Physiology Barrie Ashby, Pharmacology Hong Wang, Pharmacology Wolfgang Bergmeier, External Reader, Thomas Jefferson University ABSTRACT

REGULATION OF SYK ACTIVITY IN GPVI-MEDIATED PLATELET

ACTIVATION

Dafydd Thomas

Doctor of Philosophy

Temple University School of Medicine, 2010

Doctoral Advisory Committee Chair: James L. Daniel, Ph.D.

Activation of is essential for hemostasis. Following damage to the vascular endothelium is exposed, to which platelets stably adhere. After adhesion on collagen, a signaling cascade is initiated, mediated by Glycoprotein VI (GPVI), which results in platelet activation. A major signaling protein in GPVI signaling is Spleen tyrosine kinase (Syk). It undergoes phosphorylation and activation following GPVI stimulation. Syk's central role in this physiological process suggests regulation of its activity is required to maintain the platelets response to collagen within physiological limits. The regulation of Syk activation is the focus of this work.

Previously published reports implicate the phosphatases SHP-1, SHP-2 and TULA-2 in the negative regulation of Syk. Therefore, we tested these phosphatases possible role in platelets. We show that SHP-1 can dephosphorylate Syk in vitro , but is unable to bind

Syk. Also, Syk is hypophosphorylated in GPVI-stimulated SHP-1 deficient platelets and platelet functional responses are minimally affected compared to wild-type platelets.

ii SHP-2 is unable to bind Syk and Syk is not a good substrate for SHP-2 in vitro . TULA-2 dephosphorylated Syk in vitro and associated with Syk in platelets. In TULA-2 deficient platelets, Syk and PLC γ2 were hyperphosphorylated compared to wild-type platelets.

Deletion of TULA-2 resulted in enhanced GPVI-dependent platelet functional responses and a prothrombotic phenotype.

c-Cbl has been shown to be a negative regulator of GPVI signaling, possibly by regulating Syk phosphorylation. Thus, SHP-1, SHP-2 and TULA-2’s role in c-Cbl regulation of GPVI was also investigated. We show that TULA-2 is able to bind c-Cbl in platelets. SHP-1 and SHP-2 do not. Furthermore, we show a striking similarity between the phenotype of TULA-2 and c-Cbl deficient platelets. However, in vitro binding studies show TULA-2 is able to bind Syk independently of c-Cbl. Thus, the exact role of c-Cbl in regulating Syk dephosphorylation is unclear.

In conclusion, we show SHP-1 and SHP-2 are probably not involved in the negative regulation of Syk. However, TULA-2 is the major phosphatase responsible for the negative regulation of Syk in GPVI signaling. This serves to negatively regulate GPVI- mediated platelet function and prevent uncontrolled platelet activation that could lead to thrombosis.

iii DEDICATION

This dissertation is dedicated to my wife, Meredith, and my daughter, Natalie.

iv ACKNOWLEDGEMENTS

I would first like to thank my mentor, Dr. Daniel, for opening his laboratory to me and for giving me the chance to learn about platelets during the last five years. I am truly grateful for his guidance in all aspects of my graduate studies and his willingness to always help me and discuss things whenever it was needed. His dedication has provided me a firm scientific foundation on which I can build a career.

I would like to thank my committee members: Dr. Alexander Tsygankov, Dr. Satya

Kunapuli, Dr. Barrie Ashby, Dr. Hong Wang and Dr. Nick Carpino. Their feedback during my studies and for their time in reviewing my progress and thesis is greatly appreciated. I would especially like to thank Dr. Kunapuli for his support during my time at Temple, Dr. Tsygankov for leading us to TULA-2 and Dr. Ashby for his help with all things non-science related.

In the laboratory I am particularly indebted to Carol Dangelmaier. I would like to thank her for her friendship during the last five years and for her invaluable training in platelets and laboratory skills. The technical lessons and skills she has taught me I will carry through the rest of my scientific career.

I would like to thank the faculty and staff in Department of Pharmacology for their warmth and friendship during my time and Temple, all the members of Dr. Kunapuli’s laboratory for their support and all the students past and present for their friendship. I

v would particularly like to thank Todd Getz and Neil Lamarre for providing some comic relief during the day and their friendship throughout my time at Temple.

Finally, I would like to thank my family. I would like to thank my Mum and Dad and the rest of the Thomas family for giving me the best chance possible to reach my goals throughout my childhood and into adulthood. I would like to thank the Trapp family and

Heinz family for their support during my time in USA and welcoming me into their family. Last, but by no means least, I would like to thank my wife and daughter. To

Meredith, for supporting me throughout this whole process and giving me the drive to keep going when things got tough. To Natalie, for being such a source of inspiration and love and such a bundle of joy. I love you guys.

vi TABLE OF CONTENTS Page

ABSTRACT ...... II

DEDICATION...... IV

ACKNOWLEDGEMENTS ...... V

LIST OF FIGURES ...... IX

LIST OF ABBREVIATIONS ...... XI

INTRODUCTION...... 1

Platelets and Hemostasis ...... 1

Platelet Receptors ...... 3

Spleen Tyrosine Kinase (Syk) ...... 12

Tyrosine Phosphatases ...... 14

Casitas B-lineage Lymphoma (Cbl) Family Proteins ...... 21

MATERIALS AND METHODS ...... 26

Materials...... 26

Synthesis and Purification of GST-TULA-2 and GST-TULA-2 H380A ...... 27

Sf9 Cells and Protein Production ...... 28

In vitro Phosphatase Assays ...... 31

In vitro Association Assays ...... 31

Human Platelet Isolation and Preparation ...... 32

Murine Platelet Isolation and Preparation ...... 32

Murine Platelet Preparation for Ca 2+ mobilization ...... 33

Platelet Activation ...... 34

Platelet Dense Granule Secretion Measurements ...... 34

Lysate Preparation ...... 34

vii GST-pulldown Assay ...... 35

Coimmunoprecipitation ...... 35

Immunoblotting ...... 36

In vitro Kinase Assay ...... 36

FeCl 3-induced in vivo Thrombosis Injury Model...... 37

Statistical Analysis ...... 38

RESULTS ...... 39

SH2-domain Containing Phosphatases ...... 39

Summary of SH2 Domain Containing Phosphatases ...... 54

TULA Family Proteins ...... 54

Phosphatases role in the c-Cbl-mediated negative regulation of Syk ...... 96

DISCUSSION ...... 105

Is Syk a Substrate For a Specific Phosphatase Expressed In Platelets? ...... 106

Do Syk And a Specific Phosphatase Associate In GPVI activated Platelets? ...... 109

Do Phosphatase-null Murine Platelets Display a Phenotype In Which Syk Dephosphorylation Is Impaired? ...... 113

Do Phosphatases Play a Role in the c-Cbl-mediated Negative Regulation of Syk? ...... 122

Summary and Conclusions ...... 127

Future Experiments ...... 128

REFERENCES ...... 131

viii LIST OF FIGURES

Figure 1. The GPVI signaling cascade ...... 6

Figure 2. SHP-1 can dephosphorylate Syk in vitro ...... 41

Figure 3. SHP-1 does not coimmunoprecipitate with Syk in platelets ...... 42

Figure 4. GPVI-mediated platelet aggregation is no different in WT and mev mice...... 45

Figure 5. GPVI-mediated dense granule secretion is slightly suppressed in mev mice...... 46

Figure 6. Syk is hypophosphorylated in GPVI-stimulated mev platelets...... 47

Figure 7. SHP-1 does not associate with c-Cbl in platelets...... 48

Figure 8. Syk is not a good substrate for SHP-2 in vitro ...... 50

Figure 9. Syk does not coimmunoprecipitate with SHP-2...... 51

Figure 10. SHP-2 does not coimmunoprecipitate with c-Cbl in platelets...... 53

Figure 11. TULA-2 but not TULA is expressed in human and murine platelets and deletion of TULA proteins does not affect protein expression...... 56

Figure 12. TULA-2 can be successfully expressed in Sf9 cells and purified to homogeneity...... 59

Figure 13. TULA-2 dephosphorylation of Syk...... 63

Figure 14. Syk and c-Cbl associate with GST-TULA-2 H380A and GST-TULA-2...... 64

Figure 15. Syk and c-Cbl can associate with TULA-2...... 65

Figure 16. Syk is hyperphosphorylated in TULA-dKO platelets in a time dependent manner...... 68

Figure 17. Syk is hyperphosphorylated in TULA-dKO platelets in a dose dependent manner...... 72

Figure 18. Syk is more active in TULA-dKO platelets than WT platelets...... 73

Figure 19. PLC γ2 is hyperphosphorylated in TULA-dKO platelets in a time dependent manner...... 75

ix Figure 20. PLC γ2 is hyperphosphorylated in TULA-dKO platelets in a dose dependent manner...... 77

Figure 21. Ca 2+ mobilization is potentiated in TULA-dKO platelets...... 79

Figure 22. GPVI-mediated platelet aggregation is potentiated in TULA-dKO platelets. 82

Figure 23. GPVI-mediated platelet dense granule secretion is enhanced in TULA-dKO platelets...... 83

Figure 24. PAR4-mediated platelet aggregation is not affected in TULA-dKO platelets. 85

Figure 25. PAR4-,mediated dense granule secretion is not affected in TULA-dKO platelets...... 86

Figure 26. TULA-dKO mice are prothrombotic...... 88

Figure 27. Syk is hyperphosphorylated in TULA-2 deficient platelets in a time dependent manner...... 90

Figure 28. Syk is hyperphosphorylated in TULA-2 deficient platelets in a dose dependent manner...... 92

Figure 29. GPVI-mediated platelet aggregation is enhanced in TULA-2 deficient platelets...... 94

Figure 30. GPVI-mediated dense granule secretion is enhanced in TULA-2 deficient platelets...... 95

Figure 31. Syk is hyperphosphorylated in c-Cbl deficient platelets...... 98

Figure 32. PLC γ2 is hyperphosphorylated in c-Cbl deficient platelets...... 99

Figure 33. Ca 2+ mobilization is not potentiated in c-Cbl deficient platelets...... 101

Figure 34. GPVI-mediated platelet aggregation is enhanced in c-Cbl deficient platelets...... 102

Figure 35. TULA-2 does not require c-Cbl to bind to Syk...... 104

x LIST OF ABBREVIATIONS

4H Four-helix bundle AA Arachidonic Acid ADP Adenosine diphosphate ADAP Adhesion and degranulation-promoting adapter protein ANOVA Analysis of Variance ATP Adenosine triphosphate Btk Brutons tyrosine kinase c-Cbl cellular-Casitas B-lineage CEACAM Carcinoembryonic antigen-related cell adhesion molecule CLEC-2 C-type lectin-like 2 COX Cyclooxygenase CRP Collagen-related peptide DAG Diacyl glycerol EGFR Epidermal growth factor receptor FcR γ Fragement crystallizable receptor gamma chain Gads Grb2-related adaptor down-stream of Shc GEF Guanine nucleotide exchange factor GPIa Glycoprotein Ia GPVI Glycoprotein VI GST Glutathione-S-transferase HRP Horseradish Peroxidase Ig Immunoglobulin IL-2 Interleukin-2 IP3 Inositol trisphosphate ITAM Immunoreceptor tyrosine activation motif ITIM Immunoreceptor tyrosine inhibition motif kDa kiloDalton LAT Linker-for-activation of T-cells mev Motheaten viable PAR Protease activated receptor PBMC Peripheral blood mononucleocytes PECAM Platelet endothelial cell adhesion molecule PGH2 Prostoglandin H2 PGM Phosphoglycerate mutase PI3K Phosphoinositide-3-kinase PIP2 Phosphatidylinositol 4,5-bisphosphate PIP3 Phosphatidylinositol 3,4,5-trisphosphate PLC γ2 Phospholipase C γ2 PRP Platelet rich plasma

xi PS Phosphatidylserine PTP Protein tyrosine phosphatase RING Really interesting new gene SH2 Src-homology domain 2 SH3 Src-homology domain 3 SHP SH2 domain containing phosphatase SLP76 SH2 domain containing leukocyte protein of 76kDa STS Supressor of T-cell signaling Syk Spleen tyrosine kinase TCR T-cell receptor TKB Tyrosine kinase binding domain TULA T-cell ubiquitin TULA-dKO T-cell ubiquitin ligand double deficient Tyk2 Tyrosine kinase 2 TxA2 UBA Ubiquitin-associated VASP Vasodilator-stimulated phosphoprotein WASP Wiskott-Aldrich syndrome protein WT Wild-type ZAP70 Zeta-chain-associated protein kinase 70

xii CHAPTER 1

INTRODUCTION

Platelets and Hemostasis

Hemostasis, the cessation of bleeding, is an important process that maintains the integrity of the circulatory system and prevents hemorrhage. It is a complex process that requires three main components, vascular spasm, blood and platelet activation. Vascular spasm serves to temporarily reduce blood flow to the vessels at the site of injury. Platelet activation occurs to allow the formation of a platelet plug at the site of injury to prevent blood loss from the injured vessel and blood coagulation occurs to strengthen the platelet plug.

Platelets critical role in hemostasis is highlighted in the disease states thrombocytopenia, where platelet count is low and Glanzmanns thrombasthenia, where platelets are dysfunctional. In thrombocytopenia, bleeding in the mouth occurs as well as bruising due to the lack of platelets available to arrest bleeding (Mullally

2006). Glanzmanns thrombasthenia is characterized by increased susceptibility to bruising, menorrhagia, and increased post-operative bleeding (Franchini, Favaloro, and Lippi 2010). Additionally platelets have a pathogenic role in myocardial infarction and stroke (Helft and Worthley 2001). Rupture of unstable atherosclerotic plaques causes the exposure of thrombogenic surfaces. Circulating platelets will be activated by the thrombogenic surface leading to the formation of a thrombus which,

1 if large enough, can fully occlude the artery and cause ischemic events (Helft and

Worthley 2001).

Platelets are generated in the bone marrow by megakaryocytes. Platelets bud off from the megakaryocyte and are released into the bloodstream (Blockmans,

Deckmyn, and Vermylen 1995; Patel, Hartwig, and Italiano 2005). Here they circulate for eight to twelve days until they are cleared by the liver and spleen. As a result of being budded from megakaryocytes platelets are anucleated and are smaller than other circulating cells with a diameter of 2 to 3 m.

While circulating, platelets exist in a quiescent discoid state and must undergo a series of complex processes to become activated in order to arrest bleeding as reviewed by Jurk and Kehrel (Jurk and Kehrel 2005). The process begins with the rolling of platelets along the newly exposed subendothelial collagen at the site of injury and eventual stable adherence to the exposed collagen. The collagen bound platelet then signals via a receptor for collagen found on the platelet surface. This causes the platelet to undergo a process known as shape-change, extending filopodia and then lamellipodia. In addition, the platelet then secretes prothrombotic autocoids from stored granules, generates the prothrombotic compound thromboxane A2

(TxA2) and activates the receptor for fibrinogen found on the platelet surface. The secretion of prothrombotic autocoids and generation of TxA2 serves to recruit more circulating platelets to the site of injury and also feedback on the collagen bound platelets to cause further secretion, TxA2 generation and fibrinogen receptor

2 activation (Holmsen 1989). The activation of the fibrinogen receptor allows

fibrinogen to bind to its receptor and form a bridge between adjacent platelets to form

a platelet plug to arrest bleeding.

Platelet Receptors

In order for the platelet to respond to prothrombotic agents it expresses a number of

receptors to transduce the signal from outside the cell and evoke activation of the

platelet. The major platelet agonist receptors are as follows:

Glycoprotein VI

It was first thought that the stable adhesion of platelets to collagen was mediated by

the α2β1 integrin found on the platelet surface based on observations by Nieuwenhuis

et al. and Kehrel et al. (Nieuwenhuis et al. 1985) (Kehrel et al. 1988). They identified

two patients who had prolonged bleeding times and whose platelets failed to respond

to collagen. They concluded this was due to defects in Glycoprotein Ia (GPIa)/ α2 integrin expression on the platelet surface. In 1989 Moroi et al. proposed that a 62 kiloDalton (kDa) protein identified as GPVI was also important in collagen-mediated platelet activation. This was based on observations of a patient who had a mild bleeding disorder. The investigators discovered that the patient’s platelets were unresponsive to collagen but not other platelet agonists and subsequently found the platelets lacked GPVI expression on their surface (Moroi et al. 1989). Later work by

3 Santoro et al. lead to a ‘two-site two-step’ model of collagen mediated platelet

activation being proposed (Santoro et al. 1991). The authors hypothesized that the

α2β1 integrin initially bound to collagen in a high affinity interaction that then allowed another collagen receptor on the platelet surface, now known to be GPVI, to bind to collagen and cause platelet activation. Discoveries made in the nineties led to a modification of the two-site two-step model to accommodate the finding that the

α2β1 integrin is required to be in a high affinity state before it can stably bind to collagen (Jung and Moroi 1998) (Polanowska-Grabowska, Simon, and Gear 1999).

Watson et al. proposed that initial binding of platelets to collagen is mediated by either GPVI or α2β1 and the subsequent binding of the complementary receptor serves to strengthen the platelet binding (Watson et al. 2000). Signals from GPVI then convert α2β1 from its low affinity state to a high affinity state, further stabilizing the platelet binding. GPVI continues to signal, inducing secretion of prothrombotic autocoids thus leading to formation of a stable platelet plug.

GPVI is thought to be the primary signaling receptor for collagen on the platelet surface (Nieswandt and Watson 2003). Following the exposure of subendothelial collagen it is thought that GPVI is the first collagen receptor to evoke a signal to activate platelets. It is a member of the Immunoglobulin (Ig) family of receptors that localizes to the plasma membrane of the platelet, traversing it once (Watson et al.

2005). It is made up of a number of domains, two Ig domains, a juxtamembrane domain and a proline-rich domain (Watson et al. 2005). A salt bridge and cytosolic domains link the receptor to an Fc receptor γ-chain homodimer (FcR γ) (Watson et al.

4 2005). This interaction is critical for GPVI signaling as the binding of collagen to the

GPVI receptor is transduced via a motif known as the immunoreceptor, tyrosine-

based activation motif (ITAM) in the FcR γ-chain. On binding of collagen to GPVI, it is thought that clustering of the receptor occurs and phosphorylation of the ITAM results (Gibbins et al. 1996; Poole et al. 1997). This is thought to be mediated by Src- family tyrosine kinases Fyn and Lyn which are constitutively associated with GPVI via their Src-homology-3 (SH3) domains interacting with the proline rich domain of

GPVI (Ezumi et al. 1998; Quek et al. 2000; Suzuki-Inoue et al. 2002). Following phosphorylation of the ITAM, the tyrosine kinase Syk is recruited to the phosphorylated ITAM and interacts via its dual Src-homology-2 (SH2) domains.

Once bound, Syk undergoes tyrosine phosphorylation at multiple sites including

Y525 and Y526 (Y519/20 in murine Syk) to increase its kinase activity (Furlong et al.

1997). The activation of Syk leads to the formation of a protein complex containing a number of adapter proteins, kinases and other enzymes (Gross, Melford, and Watson

1999; Pasquet, Gross et al. 1999). As can be seen in figure 1, the Syk mediated phosphorylation of the adapter proteins Linker-for-activation-of-T-cells (LAT) and

SH2-domain-containing-leukocyte-protein-of-76kD (SLP-76) and the recruitment of

GRB2-related adapter protein 2 (Gads) allows the colocalization of Brutons tyrosine kinase (Btk), Phosphatidyl-inositol-3-kinase (PI3K) and phospholipase C γ2 (PLC γ2).

PLC γ2 activation and recruitment to the complex is aided by Btk and PI3K's conversion of Phosphatidylinositol bisphosphate (PIP2) to Phosphatidylinositol trisphosphate (PIP3), providing a binding site for PLC γ2‘s PH domain (Humphries et al. 2004; Quek, Bolen, and Watson 1998) (Pasquet, Bobe et al. 1999). In addition, 5 PLC γ2's SH2 domain can also interact with LAT to further aid recruitment (Gross,

Melford, and Watson 1999). PLC γ2 is then able to catalyze the conversion of PIP2 to diacylglycerol (DAG) and Inositol Trisphosphate (IP3) causing the mobilization of

Ca 2+ from intracellular stores, critical for platelet activation (Daniel, Dangelmaier, and Smith 1994).

Figure 1. The GPVI signaling cascade

6 GPVI agonists

A number of agonists for GPVI are used experimentally to study GPVI signaling.

These are as follows:

Convulxin

Convulxin is a purified from Crotalus durissus terrificus . It is made up of α and β chains linked by disulfide bonds (Batuwangala et al. 2004). These multimerize to form a ring-like structure to allow it to cluster multiple GPVI receptors. It is considered a potent agonist due to its high affinity for GPVI and its ability to activate platelets without the need for feedback from secreted platelet agonists (Atkinson et al. 2001; Francischetti et al. 1997; Quinton et al. 2002).

Convulxin can also interact with GPIb α however (Kanaji et al. 2003). This is a caveat when using convulxin.

Collagen-related peptide

Collagen-related peptide (CRP) is a synthetically produced agonist. It is made up of a triple helical peptide that is cross-linked at both termini via cysteine residues. The primary structure consists 10 glycine-proline-hydroxyproline (GPO) repeats. Unlike convulxin, CRP requires feedback from secreted platelet agonists in order to cause full platelet activation (Jarvis, Best, and Watson 2004; Quinton et al. 2002).

Therefore, is more like the natural agonist, collagen, in terms of platelet activation and can be considered a more physiological agonist than convulxin. 7 Collagen

Collagen is the physiological agonist for GPVI. It can be classified into fibrous and non-fibrous and exists in up to 25 forms, with types I, III, IV, V and VI being most abundant in the vessel wall. Like CRP it is made up of a triple helical structure that is required for GPVI binding. The primary structure contains approximately 10% GPO repeats which mediate GPVI binding. Collagen also requires feedback from secreted platelet agonists to cause full platelet activation (Farndale et al. 2004). In these studies we will use Type I collagen from an equine source for platelet activation.

Convulxin is a powerful tool for studying GPVI-mediated platelet activation and complementing these studies with the CRP and collagen can give a more physiological aspect to GPVI-mediated platelet activation.

G-protein coupled receptors

ADP-receptors

One major prothrombotic autocoid secreted by the platelet is adenosine diphosphate

(ADP) (Kunapuli et al. 2003). Following secretion, ADP can bind to three receptors

expressed on the platelet surface, two of which are GPCR’s, namely P 2Y1 and P 2Y12

(Daniel et al. 1998). Each GPCR receptor couples to a different G-protein to evoke a

different intracellular response. P 2Y1 couples to Gq to mobilize stored calcium. P 2Y12 couples to Gi thus inhibiting adenylyl cyclase and lowering intracellular cyclic-AMP 8 (Dorsam, Tuluc, and Kunapuli 2004; Jin, Daniel, and Kunapuli 1998). The action of

ADP on these receptors following its release from intracellular stores serves to strengthen the interaction of collagen bound platelets as well as recruit circulating platelets to the site of injury (Dangelmaier et al. 2001; Quinton et al. 2002; Storey,

Newby, and Heptinstall 2001).

ThromboxaneA2 receptor

Thromboxane A2 is generated by a multistep enzymatic process that first involves the action of phospholipases on glycerophospholipids found in the inner leaflet of the platelet membrane that causes the release of arachidonic acid (AA) into the cytosol of the platelet. Once released, the AA is converted into prostoglandin H2 (PGH2) by cyclooxygenase-1 (COX-1) which is quickly isomerized into TxA2 by thromboxane synthase (Foegh 2001). The soluble TxA2 then diffuses from the platelets cytosol to act on circulating platelets and feedback on the platelet generating the TxA2. The

TxA2 achieves this by binding to the TP receptor. The TP receptor couples to both

Gq and G12/13 thus mobilizes calcium from intracellular stores via Gq and activates

RhoA via G12/13 which results in the phosphorylation of myosin light chain phosphatase and down regulation of its activity (Allan et al. 1996). This causes an increase in cytoskeletal rearrangements and platelet activation.

9 Thrombin receptors

In addition to sub-endothelial collagen being exposed at the site of vascular injury,

thrombin is also generated. Thrombin can activate platelets, causing platelet shape-

change, secretion, TxA2 generation, and αIIb β3 activation (Kahn et al. 1998). Like

ADP, thrombin acts on two receptors in human platelets, protease activated receptor-

1 (PAR-1) and PAR-4, however unlike ADP they couple through the same G-

proteins, Gq and G12/13. Thus mobilizing stored calcium and causing increased

myosin phosphorylation (Kahn et al. 1998). Members of the PAR family of receptors

have a unique mechanism of activation. Their N-terminus undergoes enzymatic

cleavage by proteases such as thrombin and the resulting shortened N-terminus acts

as a tethered ligand and binds to the exposed extracellular loops of the receptor to

cause receptor activation (Coughlin 1999). An interesting difference between murine

and human platelets is that while human platelets express PAR-1 and PAR-4, murine

platelets express PAR-3 and PAR-4 and PAR-3 only acts as a coreceptor for PAR-4

and cannot signal individually (Nakanishi-Matsui et al. 2000).

αIIb β3 integrin

The αIIb β3 integrin, or glycoprotein IIb/IIIa is the receptor for fibrinogen. It is made

up of two proteins, αIIb integrin and β3 integrin which exist on the platelet surface in a low affinity state (Shattil, Kashiwagi, and Pampori 1998). Binding of the platelet to collagen and feedback from ADP and TxA2 receptor signaling, as well as platelet activation by thrombin, causes the transition of the integrin to its high affinity state

10 (activation), termed inside-out signaling (Shattil and Newman 2004). This allows fibrinogen to bind and bridge between adjacent platelets that have been recruited to the site of injury. Thus allowing the formation of a platelet aggregate crosslinked by fibrin at the site of injury. Additionally, the integrin forms multi-protein complexes via the cytoplasmic tail of the β3 chain. Briefly, upon binding of fibrinogen to the integrin, c-Src, which is constitutively associated with the β3 cytoplasmic tail, undergoes deinhibition by dephosphorylation (Arias-Salgado et al. 2003; Obergfell et al. 2002). It is then able to phosphorylate recruited Syk by virtue of its tyrosine kinase activity. The two tyrosine kinases then phosphorylate a number of downstream substrates such as the guanine nucleotide exchange factor (GEF) Vav.

This causes the activation of the small G-protein Rac which leads to cytoskeletal rearrangements and platelets spreading. The phosphorylation of the adapter proteins

SLP-76, adhesion and degranulation-promoting adapter protein (ADAP), and cellular- casistas B-lineage lymphoma protein (c-Cbl) also occurs which provide a platform for the recruitment of PI3K, the cytoskeletal modulators Vasodilator-stimulated

Phosphoprotein (VASP) and Wiskott-Aldrich Syndrome Protein (WASP) thus aiding cytoskeletal rearrangements (Bearer, Prakash, and Li 2002; Judd et al. 2000; Miranti et al. 1998; Obergfell et al. 2001). Additionally, the phosphorylation of SLP-76 allows for the recruitment and activation of PLC γ2 that leads to calcium mobilization and PKC activation, which is required for full platelet activation (Wonerow et al.

2003). This is termed outside-in signaling.

11 Working in concert, these platelet agonists serve to cause complete platelet activation at the site of injury to cause the cessation of bleeding.

Spleen Tyrosine Kinase (Syk)

The tyrosine kinase Syk was first described by Zioncheck et al. in 1986 (Zioncheck,

Harrison, and Geahlen 1986). Subsequent work lead to Syk’s isolation from the porcine spleen (Kobayashi et al. 1990; Sakai et al. 1988), hence its name.

Syk structure and phosphorylation

Cloning of Syk by Taniguchi et al. showed that Syk contains two SH2 domains and a tyrosine kinase domain (Taniguchi et al. 1991). The tandem SH2 domains allow Syk to interact with tyrosine phosphorylated ITAMs to localize Syk to specific areas within the cell for . C-terminal to both SH2 domains are interdomains A and B. Interdomain A can associate with other proteins by virtue of its coiled coil domain and Interdomain B contains five tyrosine residues that can undergo phosphorylation allowing Syk to bind with SH2 domain containing proteins

(Sada et al. 2001). Thus far three of these five phosphorylation sites binding partners have been identified. Deckert et al. have documented that phosphorylation at tyrosine

317 in murine Syk (Y323 in human Syk) allows Syk to bind to c-Cbl (Deckert et al.

1998). This has been reported to be important for Syk negative regulation (Lupher et al. 1998; Ota et al. 2000; Yankee et al. 1999). Tyrosine 342 (Y348 in human Syk) and 346 (Y352 in human Syk) phosphorylation have been reported to form a docking

12 site for proteins that interact with PLC γ1 (Finco et al. 1998; Kurosaki et al. 1995;

Pivniouk et al. 1999; Saitoh et al. 2000). Within the activation loop of the kinase domain tyrosines 519 and 520 (525 and 526 in human Syk) undergo phosphorylation and are critical phosphorylation events for the transduction of cellular signaling events and can be used as a marker of Syk activation (Kurosaki and Tsukada 2000;

Zhang, Kimura, and Siraganian 1998). The most C-terminal phosphorylations of Syk are at tyrosines 624 and 625 and have been reported to negatively regulate Syk function downstream of the T-cell receptor (TCR) (Zeitlmann et al. 1998). Due to

Syk’s tandem SH2 domains and multiple phosphorylation sites it can be regulated by an array of mechanisms.

Syk in platelets

Syk was first implicated in platelet activation in 1992 by Ohta et al. who showed that

Syk undergoes autophosphorylation in response to wheat-germ agglutinin (Ohta et al.

1992). Later work by Fujii et al. showed that collagen could induce tyrosine phosphorylation of Syk and increase its kinase activity and was a result of direct collagen-induced signaling and not a function of feedback (Fujii et al. 1994). Later work by Poole et al. demonstrated that platelets deficient in Syk have many deficits in

GPVI mediated platelet activation. It was shown that platelets devoid of Syk fail to aggregate, secrete stored granules, form arachidonic acid and exhibit no PLC γ2 phosphorylation in response to GPVI agonists (Poole et al. 1997). Syk’s role in GPVI signaling is therefore undisputed and tight regulation of its activation and deactivation

13 are of critical importance to ensure necessary platelet activation to prevent bleeding while preventing uncontrolled platelet activation that may lead to thrombosis.

Tyrosine Phosphatases

Phosphorylation is the addition of a phosphate group to a compound. In the biological setting this is often the addition of phosphate to a protein and is catalyzed by an enzyme class known as kinases. This process requires adenosine triphosphate

(ATP) and is used as molecular switch to change the activity state and/or localization of the target protein. Classically the phosphorylation of substrates such as kinases leads to an increase in their kinase activity and the propagation of signaling events.

Conversely, dephosphorylation is the removal of a phosphate group and in the biological setting this is catalyzed by a class of proteins known as phosphatases.

These classically serve to deactivate kinases and terminate signaling events.

Given the involvement of unregulated platelet activation in thromboembolic events and their potential for serious morbidity and mortality, the tight regulation of platelet activation is of critical importance. As described above, tyrosine phosphorylation events are paramount to the activation of the platelet by collagen. Therefore, tyrosine phosphatases are likely to have an important role in regulating collagen mediated platelet activation and more specifically the dephosphorylation of the critically important kinase Syk.

14 SH2-domain containing phosphatases

To date 107 human cysteinyl protein-tyrosine-phosphatases (PTPs) have been

identified, 81 of which specifically dephosphorylate protein substrates. The

superfamily catalytic domain is comprised of a conserved Cysteine (C) X 5 Arginine

(R) motif with the C residue acting as a nucleophile to attack the phosphate attached

to the substrate and the R residue binds the phosphate group (Kim and Denu 2003).

The superfamily is subdivided into four families based on sequence homology within

the catalytic domain. The SH2-domain containing phosphatases belong to a

subdivision of the class I PTPs which exclusively dephosphorylate phosphorylated

tyrosines, the other subdivision being dual specificity phosphatases,

dephosphorylating tyrosine and threonine residues. Of the 17 reported cytosolic, non-

receptor PTPs only two contain dual SH2 domains, namely Src homology domain-

containing PTP-1 (SHP-1) and SHP-2 (Alonso et al. 2004; Poole and Jones 2005).

SHP-1

Four isoforms of SHP-1 exist, encoded by one gene. Three variants differ due to the site of transcription initiation with one of these transcripts being exclusively expressed in cells of hematopoietic origin (Banville, Stocco, and Shen 1995). The other isoform is a splice variant alternatively spliced at the C-terminus leading to the elongation of SHP-1 by 66 amino acids and is termed SHP-1L (Jin, Yu, and Burakoff

1999). Extensive studies of SHP-1 have been undertaken, not least with respect to

TCR signaling. With regard to TCR signaling, SHP-1 appears to be a negative regulator with T-cells isolated from the spontaneously occurring SHP-1 inactivating 15 mutant, motheaten (me/me) mouse displaying a hyper-responsive phenotype in terms of protein phosphorylation and functional responses (Carter, Neel, and Lorenz 1999;

Lorenz et al. 1996). These data were confirmed in T-cell cell lines where SHP-1 was overexpressed. A suppression of T-cell receptor signaling protein phosphorylation was observed as well as Interleukin-2 (IL-2) production (Fawcett and Lorenz 2005;

Sankarshanan et al. 2007). To date however the exact substrates for SHP-1 have not been identified but may include Zeta-chain-associated protein kinase 70 (Zap70), Vav and Lck (Lorenz 2009). Given the similarities between TCR signaling and platelet

GPVI signaling SHP-1 could have an important role in collagen signaling in platelets.

In 2000, Pasquet et al. presented evidence for SHP-1’s role in GPVI signaling. They showed that SHP-1 can coimmunoprecipitate with Syk following GPVI stimulation and showed a robust pulldown of Syk in a GST-pulldown assay using the tandem

SH2 domain of SHP-1 (Pasquet et al. 2000). However, when investigating SHP-1’s function using SHP-1 deficient platelets, they show the unexpected result of tyrosine hypophosphorylation of Syk following GPVI stimulation. This would suggest that

SHP-1 could potentiate signaling in platelets, contrary to previously published data from immune cells. An important observation made by Pasquet and coworkers was that SHP-1’s role in the regulation of GPVI signaling occurred after the mobilization of Ca 2+ . Therefore a possible role of SHP-1 in negative regulation of Syk and GPVI signaling prior to Ca 2+ mobilization remains to be investigated and could correlate with the regulatory mechanisms seen in T-cells.

16 SHP-1 has also been shown to associate indirectly with the adapter protein and E3

ligase c-Cbl (Uddin et al. 1996). Earlier work showed that SHP-1 could associate

with the kinase Interferon-dependent Tyrosine Kinase (Tyk2) in cell lines (Yetter et

al. 1995). A follow-up paper then showed that the Tyk2 could associate with c-Cbl in

various hematopoietic cells lines (Uddin et al. 1996). The implications of this

reported c-Cbl binding is discussed further below.

SHP-2

The role of SHP-2 in TCR signaling remains equivocal. A positive and negative regulatory role has been proposed depending on the experimental methodologies utilized. Kwon et al. have reported that overexpression of a phosphatase inactive mutant of SHP-2 leads to a potentiation of the phosphorylation of the signaling molecules SLP-76, Vav1 and ADAP and an increased adhesion of T-cells to fibronectin (Kwon et al. 2005). Additionally, Salmond et al. have shown that SHP-2 suppresses the differentiation of T-cells but appeared to be independent of the TCR

(Salmond et al. 2005). Conversely, Nguyen et al. have shown that conditional deletion of SHP-2 causes a suppression of TCR signaling as measured by a suppression of ERK phosphorylation as well as a reduction in IL-2 secretion (Nguyen et al. 2006). These data suggest that SHP-2 is a positive regulator of TCR signaling.

In addition, Xu and Pecht have shown SHP-2 can associate with Syk in RBL cells.

In platelets SHP-2 has been reported to be a negative regulator of GPVI signaling by virtue of its association with the inhibitory receptor platelet endothelial cell adhesion molecule-1 (PECAM-1). Utilizing PECAM-1 deficient platelets Patil et al. have

17 shown that a lack of SHP-2 activation when PECAM-1 is not present leads to a

potentiation of GPVI-mediated signaling (Patil, Newman, and Newman 2001). The

role of SHP-2 was inferred from previously published work by the same group that

SHP-2 binds to PECAM-1 (Jackson et al. 1997).

SHP-2 has also been shown to associate with c-Cbl. In 2001 Chernock et al. showed

that SHP-2 was present in c-Cbl immunoprecipitates and c-Cbl was present in SHP-2

immunoprecipitates following stimulation of Jurkat T-cells with stromal derived

factor-1α ( SDF −1α) (Chernock, Cherla, and Ganju 2001). Additionally, GST- pulldowns using the SH-2 domain of SHP-2 were able to pull down c-Cbl. The implications of this association with c-Cbl are discussed further below.

Given the possible negative regulatory role of SHP-2 in TCR signaling, the reported association of SHP-2 with Syk in RBL cells and the inferred negative regulatory role of SHP-2 in platelets, consideration of SHP-2 as a regulator of Syk phosphorylation will be made in this study.

Histidine phosphatases

The histidine phosphatase superfamily encompasses a wide array of proteins originally classified as mutases, which transfer phosphate from one residue on a protein to another. Further work demonstrated that a large number of proteins within the superfamily were actually phosphatases (Rigden 2008). As their name suggests,

18 histidine is an important residue utilized by the superfamily for catalysis. A conserved histidine found in the catalytic pocket undergoes phosphorylation and dephosphorylation during the dephosphorylation of substrates. In addition to the conserved histidine, the superfamily also conserves an Arginine (R) and Glycine (G) residue within the catalytic pocket which makeup the RHG motif essential for catalysis. Additionally, another H and R are conserved within the catalytic pocket which hydrogen bond with the phosphate group to aid catalysis.

T-cell Ubiquitin Ligand (TULA) Proteins

The TULA family of proteins is comprised of two proteins: TULA, also known as either suppressor of T-cell receptor signaling (STS-1) or ubiquitin binding domain

(UBA), and SH3 domain-containing protein A (UBASH3A) and TULA-2 (STS-1,

UBASH3B) (Tsygankov 2008). TULA is primarily expressed in lymphoid cells whereas TULA-2 is ubiquitously expressed. TULA was first cloned by Wattenhofer et al. who were looking for candidate genes for autosomal recessive nonsyndromic deafness (Wattenhofer et al. 2001). While searching for c-Cbl binding proteins

Feshchenko et al. discovered that TULA could associate with c-Cbl in T-cells

(Feshchenko et al. 2004). Work performed by Carpino et al. led to the cloning of a protein with similar homology to TULA and later work resulted in the cloning of both

TULA family members (Carpino et al. 2002; Carpino et al. 2004). Kowanetz et al. further confirmed the identity of the TULA family proteins and the ability to bind c-

Cbl (Kowanetz et al. 2004).

19 In the same report in which Carpino et al. cloned both TULA family members they also generated mice deficient in both family members. Characterization of these mice revealed that the TULA family proteins had a suppressive role in TCR signaling observed as a hyperphosphorylation of Zap70, a protein that shares 50% structural identity with Syk, as well as a hyperproliferative T-cell response but only when both family members were deleted. The mechanism of this TULA family protein negative regulation of TCR signaling however remained unclear.

During these initial studies, it was remarked that proteins belonging to the phosphoglyceromutase (PGM) family can also act as phosphatases not just phosphoglyceromutases. Additionally, the TULA family proteins PGM domain also bore structural resemblance to the acid phosphatase family (Jedrzejas 2000).

Therefore TULA family proteins were investigated as possible phosphatases.

Mikhailik et al. confirmed that TULA family proteins were indeed phosphatases

(Mikhailik et al. 2007). The group showed that TULA proteins could liberate phosphate from the prototypic phosphotyrosine substrate para-Nitrophenol phosphate, a tyrosine phosphorylated peptide as well as dephosphorylate phosphorylated Zap70 downstream of the TCR although TULA-2 had approximately 2000-fold greater phosphatase activity than TULA. In this study, it was also shown that retroviral reconstitution of TULA-2 into primary T-cells deficient in both TULA members reversed the hyperproliferative phenotype. More recently, Agrawal et al. reported that the TULA-2 phosphatase domain could dephosphorylate Syk and this was a preferential dephosphorylation over Src-family kinases (Agrawal, Carpino, and

20 Tsygankov 2008). Additionally, the same group showed that TULA-2 could coimmunoprecipitate with Syk when both were overexpressed in HEK293T cells.

With regard to TULA’s phosphatase activity, the original description of TULA proteins phosphatase activity showed that TULA-2 has 2000 fold greater phosphatase activity than TULA at physiological pH. However, a follow up study by the same group now reports that optimal phosphatase activity for TULA may occur at acidic pH, therefore TULA may be an acid-dependent phosphatase but still remains 200 to

500-fold less active that TULA-2 at acid pH (Chen et al. 2009).

TULA proteins are also able to bind the adapter and E3 ligase c-Cbl. As discussed above, Feschenko et al. described TULA as a c-Cbl binding protein. This was elucidated using affinity chromatography using c-Cbl as a bait protein and subsequent mass spectroscopy of associating protein bands. This association was found to be

SH3 and UBA domain dependent. TULA-2 binding to c-Cbl was reported by

Kowanetz et al. who showed by GST-pulldown that the SH3 domain of TULA-2 could interact with c-Cbl (Kowanetz et al. 2004). The implications of TULA family protein interaction with c-Cbl are discussed below.

Casitas B-lineage Lymphoma (Cbl) Family Proteins

The Cbl family of proteins have been recently been implicated in the negative regulation of GPVI signaling. Three mammalian forms of this family of proteins have been discovered to date, namely c-Cbl, Cbl-b and Cbl-3, as well as other non-

21 mammalian orthologs. c-Cbl and Cbl-b are ubiquitously expressed with notable expression of Cbl-b in the spleen and c-Cbl in hematopoietic cells (Tsygankov et al.

2001). Cbl-3 expression, however, is mostly confined to the gastrointestinal tract

(Tsygankov et al. 2001). The Cbl family of proteins have a large variety of domains, a number of which are shared across the three mammalian homologues (Thien and

Langdon 2001). At the N-terminus is the tyrosine kinase binding domain (TKB) which is made up of a four helix bundle (4H), a calcium binding EF hand motif and an SH2 domain (Meng et al. 1999). C-terminal to this is the really interesting new gene (RING) finger domain, which facilitates the E3 ligase function of c-Cbl

(Joazeiro et al. 1999; Levkowitz et al. 1999). C-terminal to the RING domain is a proline-rich motif (Kowanetz et al. 2003) followed by a UBA (Bertolaet et al. 2001;

Chen et al. 2001; Wilkinson et al. 2001). The multiple domains found in the Cbl family proteins would suggest that they could interact with a number of proteins, which is indeed the case. Notably, proteins known to be involved in GPVI signaling such as Syk, Lyn, PLC γ, albeit isoform 1, and PI3K can associate with c-Cbl via the various domains found in c-Cbl (Donovan et al. 1994; Fukazawa et al. 1995; Graham et al. 1998; Marcilla et al. 1995). c-Cbl has also been shown to interact with a whole host of other proteins depending on the cell line, as reviewed by Tsygankov et al.

(2001), suggesting c-Cbl can act as a multivalent adapter protein to colocalize various proteins. It has also been reported that c-Cbl undergoes tyrosine phosphorylation in

T-cell antigen receptor stimulated Jurkat cells suggesting c-Cbl has a role in signal transduction (Donovan et al. 1994) and furthers c-Cbl adapter function as SH2 domain containing proteins can bind to the phosphorylated tyrosines.

22

Not only can c-Cbl act as a scaffolding protein, it can also act as a ubiquitin ligase via its RING domain as mentioned above. Ubiquitination is the enzymatic addition of the

8.5kDa protein, ubiquitin, to a lysine residue on target proteins. This can be mono-, di- or tri- ubiquitination sequentially attached to the preceding ubiquitin or the target protein at multiple sites. Classically, ubiquitination is a signal for degradation of a protein and a number of groups have shown ubiquitination of proteins by c-Cbl results in reduced protein expression, such as the Epidermal Growth Factor Receptor

(EGFR) (Levkowitz et al. 1998), Fyn (Andoniou et al. 2000) and Syk (Lupher et al.

1998), in various cell lines. However, mounting evidence also implicates ubiquitination as a signal other than for protein degradation (Schnell and Hicke

2003). Despite the mechanism of c-Cbl regulation of signaling not yet being fully understood it clearly has an important role to play.

c-Cbl's importance in platelet signaling was first reported by Oda et al. (Oda et al.

1996). The authors report that c-Cbl is expressed in platelets and undergoes tyrosine phosphorylation on stimulation of the platelets with Thrombopoeitin. They also found that c-Cbl was constitutively associated with the adapter protein Grb2. Further work by Auger et al. suggested that c-Cbl was involved in GPVI signaling and acted as a negative regulator of such signaling (Auger et al. 2003). Utilizing mice heterozygous for active c-Cbl they show that CRP treated c-Cbl mutant platelets exhibit hyperphosphorylation of a number of tyrosine phosphorylated proteins. They identify these proteins as FcR γ, LAT, Syk and PLC γ2. This hyperphosphorylation

23 manifests itself functionally as potentiation of platelet aggregation in the c-Cbl deficient platelets. These data suggest c-Cbl has an important negative regulatory role in GPVI signaling. In agreement with Auger et al., Dangelmaier et al. also show that c-Cbl deficient murine platelets exhibit hyperphosphorylated Syk when activated via GPVI which supports the notion that c-Cbl is a negative regulator of GPVI signaling (Dangelmaier et al. 2005).

Given the reported findings that c-Cbl null mice platelets exhibit a potentiation of Syk phosphorylation following activation of GPVI and c-Cbl exhibits no putative phosphatase domain, a phosphatase may be involved in the negative regulatory function of c-Cbl. Therefore, since SHP-1, SHP-2 and TULA-2 can associate with c-

Cbl, it is conceivable that these phosphatases could associate with c-Cbl in order for c-Cbl to negatively regulate GPVI signaling.

As mentioned above, c-Cbl also has E3 ligase activity and data from this laboratory suggests that Syk is rapidly ubiquitinated in GPVI activated platelets but this does not act as a signal for degradation of Syk (Dangelmaier et al. 2005). It was observed that c-Cbl was the E3 ligase responsible for Syk ubiquitination as c-Cbl null murine platelets did not ubiquitinate Syk. Therefore while c-Cbl’s possible interaction with a phosphatase may be the mechanism by which c-Cbl negatively regulates platelets ubiquitination of Syk may also be important.

24 Hypothesis

Given the implication of the phosphatases SHP-1, SHP-2 and TULA-2 in the negative regulation of signaling and more specifically TCR signaling, we hypothesize that these phosphatases may have a role in the negative regulation of the TCR-like signaling of GPVI and more specifically the negative regulation of Syk. This hypothesis will be broken down into four parts and addressed as follows: Is Syk a substrate for a specific phosphatase expressed in platelets? Do Syk and a specific phosphatase associate in GPVI activated platelets? Do phosphatase-null murine platelets display a phenotype in which Syk dephosphorylation is impaired? Do phosphatases play a role in the c-Cbl-mediated negative regulation of Syk?

25 CHAPTER 2

MATERIALS AND METHODS

Materials

All reagents were from Sigma (St Louis, MO) unless stated. Anti-SHP-1 (C-19), anti-SHP-1 (D-1), anti-SHP-2 (B-1), anti-c-Cbl (C-15), anti-Syk (4D10), anti-Syk

(01), anti-Syk (N-19), anti-PLC γ2 (B10), anti-α Tubulin (B7), anti-β Tubulin (D10),

anti-Horseradish peroxidase (HRP)-conjugated goat anti-mouse, anti-rat or anti-rabbit

immunoglobulin G (IgG) antibodies and protein A/G PLUS agarose were purchased

from Santa Cruz Biotechnology (Santa Cruz, CA). Antiphosphospecific-Syk

(Tyr525/6), PLC γ2 (Tyr759) and anti-Erk 1/2 (3A7) antibodies were purchased from

Cell Signaling Technology (Beverly, MA). Anti-TULA-2 antibody was purchased

from Rockland Immunochemicals (Gilbertsville, PA). Anti-phosphotyrosine (4G10)

and His-Syk was purchased from Millipore (Billerica, MA). Anti-GPVI (JAQ1) was

purchased from Emfret (Eibelstadt, Germany). Infrared dye-labelled goat anti-mouse

and anti-rabbit immunoglobulin G (IgG) antibodies were purchased from Licor

(Lincoln, NE). Type I collagen was purchased from Chronolog (Havertown, PA).

Convulxin was purified as described by Polgar et al. (Polgar et al. 1997). Collagen-

related peptide was purchased from Dr. Richard Farndale (Cambridge, UK).

AYPGKF was purchased from Genscript (Piscataway, NJ). Anti-TULA antibody

was produced as previously described (Feshchenko et al. 2004). GST-Syk and

FURA2-AM were purchased from Invitrogen (Carlsbad, California). Gelcode Blue 26 was purchased from Pierce (Rockford, IL). Recombinant SHP-1 and SHP-2 protein

were purchased from Enzo Life Sciences (Plymouth Meeting, PA). P 32 ATP was

purchased from Perkin Elmer (Waltham, MA). TULA family deficient mice were a

gift from Dr. Nick Carpino (Stoney Brook University, Stoney Brook, NY) via Dr.

Alexander Tsygankov (Temple University, Philadelphia, PA). mev mice were

purchased from Jackson Laboratories (Bar Harbor, ME). c-Cbl (-/-) mice were

acquired as previously described (Dangelmaier et al. 2005).

Synthesis and Purification of GST-TULA-2 and GST-TULA-2 H380A

TULA-2 and TULA-2 H380A DNAs were amplified by polymerase chain reaction

(PCR) from a pAlterMax plasmid containing the TULA-2 or TULA-2 H380A gene

by 30 cycles of a 1 minute 95 oC melting step, a 1 minute 64 oC annealing step and a

72 oC elongation step using Pfu Ultra II (Agilent Technologies, Santa Clara, CA). The

PCR products were then purified by separation on a 1% agarose gel run at 100V and

the appropriate band excised and isolated into 50ul dH 2O using the Qiagen Qiaquick kit (Catalogue # 28704). The resulting DNA was then quantitated using optical density and 1 g of TULA-2 DNA and the baculovirus vector pAcGHLT-B were cut with KpnI and XhoI. The pAcGHLT-B was also incubated with alkaline phosphates to remove phosphates for improved ligation (Catalogue # M1821, Agilent,

Technologies, Santa Clara, CA). A 1:1 ratio of vector and PCR product was then ligated for 5 minutes at room temperature using T4 DNA ligase from the Roche ligation kit (Catalogue # 16353791, Roche, Nutley, NJ). Following successful

27 ligation 100 l XL-10 Gold Ultracompetent cells were transformed, plated and grown overnight at 37 oC. Colonies were then isolated and grown overnight in LB broth to replicate the DNA. DNA was then isolated from the bacterial culture using the

Wizard Plus Minipreps DNA Purification System (Catalogue # TB117, Promega,

Madison, WI). Successful ligation of the TULA-2 gene was confirmed using a restriction digestion with XhoI and KpnI. Vectors with successful TULA-2 inserts were then sequenced by IDT Technologies (Coralville, IA) to confirm no mutations in the TULA-2 gene had been introduced.

Sf9 Cells and Protein Production

Transfection, amplification, expression and purification of proteins from Sf9 cells was performed using the Baculogold Transfection kit (BD Biosciences, San Jose,

CA) and was as follows:

Sf9 cell transfection

1x10 6 cells/well were plated on a six well plate at 70% confluency and allowed to attach for 15 minutes. The media was then aspirated and 0.5ml of Transfection Buffer

A (Grace’s medium, 10% FBS, pH 6.0-6.2) added to the cells. 1 g of purified

TULA-2 or TULA-2 H380A containing pAcGHLT-B was then mixed with 0.25g of

Baculogold Baculovirus DNA for 5 minutes. 0.5ml of Transfection buffer B (25mM

HEPES pH 7.1, 125mM CaCl 2, 140mM NaCl) was then added to the DNA mixture.

28 The Transfection buffer B solution was then added dropwise to the cells and the DNA

was transfected based on the calcium phosphate principle of cell transfection. The

cells were then incubated for 4 hours at 27 oC following which the transfection solution was removed and TNM-FH media added and the cells incubated for 5 days at

27 oC. After 5 days the media was aspirated and centrifuged at 2500g for 5 minutes at

4oC and the resulting supernatant containing the virus collected and stored at 4 oC for

amplification.

Baculovirus amplification

Following transfection, a sequential amplification of baculovirus was performed. For

the first amplification step 2.2x10 6 cells were plated on 60mm dish with 3ml of media. To this, 300ul (1:10 dilution) of media containing the baculorvirus from the transfection step was added and incubated for 3 days at 27 oC. The baculovirus was then collected by centrifugation of the supernatant at 2500g for 5 minutes at 4 oC. The amplification process was then repeated 3 more times each time increasing the volume and cell number the virus was amplified in at a 1:10 dilution. These volumes and cell number are as follows: 3.3x10 6 cells in 10ml of media, 1x10 7 cells in 20ml of media and 2x10 7 cells in 40ml of media.

Expression and purification of proteins

Following baculovirus amplification 2x10 7 cells were plated in 27ml of media. To this 3ml of baculovirus solution from the amplification step was added and incubated

29 for 3 days at 27 oC. Three days post infection the cells were harvested by scraping and centrifuged at 2500g for 5 minutes at 4 oC followed by resuspension in 30ml lysis

buffer (10mM Tris pH 7.5, 130mM NaCl, 1% Triton-X-100). The lysate was then

centrifuged at 40,000g for 45 minutes at 4 oC and then incubated with glutathione

agarose beads for one hour at 4 oC. The beads were then washed 3 times in 10

volumes of PBS wash buffer (10mM NaCl, 2,7mM KCl, 10mM Na 2HPO 4, 1.8mM

o KH 2PO 4, pH 7.4) and either aliquoted and stored at -80 C on PBS wash buffer supplemented with 10% glycerol or eluted 3 times for 2 minutes using GST elution buffer (50mM Tris-HCl pH 8.0, 5mM Glutathione, 10% glycerol) and stored at -

80 oC.

Protein concentration determination

In order to determine the concentration of the synthesized protein from the baculorvirus expression vector system a SDS-polyacrylamide gel was run as described in Immunoblotting below with a standard curve of BSA run against the eluted or non-eluted protein samples. The gel was stained using Gelcode blue for 1 hour and then destained using dH 2O and the resulting bands visualized using the

Fujifilm LAS-1000 (Fujifilm, Valhalla, NY) and the bands quantitated using Image

Gauge software (Fujifilm, Valhalla, NY).

Recombinant c-Cbl was also synthesized in the same manner by Carol Dangelmaier

(Temple University, Department of Physiology).

30 In vitro Phosphatase Assays

Phosphorylated GST-Syk was diluted in phosphatase assay buffer (25mM Tris pH

7.0, 50mM NaCl, 2mM EDTA, 5mM DTT, 0.01% NP-40, 1mg/ml BSA) to a final

concentration of 50nM. Prior to the addition of a phosphatase an aliquot was taken

for the 0 time-point. Then, at room temperature, either 50nM SHP-1, SHP-2 or

TULA-2 was added and aliquots taken at specific time points. Each aliquot was

stopped using 2x sample buffer (125 mM Tris pH 6.8, 4% SDS, 20% glycerol, 200

mM DTT, 0.02% bromphenol blue) and the sample boiled and immunoblotted as

described in Immunoblotting.

In vitro Association Assays

50nM agarose-coupled GST, GST-TULA-2 or TULA-2-H380A was incubated with

either 50nM His-Syk, 50nM c-Cbl or both together in Association Assay buffer (1x

NP-40 supplemented with 0.1% SDS). The mixtures were shaken for 15 minutes at

200rpm at room temperature with a stir bar to keep the agarose in suspension. After

fifteen minutes the agarose was pelleted by centrifugation at 2500g for 30 seconds at

4oC. The agarose beads were then washed 5 times for 5 minutes with 4 ml

Association Assay buffer. The agarose beads were then resuspended in 2x sample buffer and immunoblotted.

31 Human Platelet Isolation and Preparation

Blood was drawn from informed healthy volunteers according to a protocol approved

by the Institutional Review Board of Temple University into one-sixth volume ACD

(85mM sodium citrate, 111mM glucose, 71.4mM citric acid). Platelet-rich plasma

(PRP) was isolated by centrifugation at 200g for 15 minutes and incubated with 1mM

aspirin for 30 minutes at 37 oC. Platelets were obtained by centrifugation for 10 minutes at 800 g and resuspended in HEPES buffered Tyrodes solution (10mM

HEPES, pH7.4, 137mM NaCl, 2.7mM KCl, 2mM MgCl 2, 0.42mM NaH 2PO 4, 5mM

glucose, 0.2% bovine serum albumin [BSA]) and 0.2U/ml apyrase. Platelets counts

were adjusted using a Z1 Coulter Particle Counter (Beckman Coulter, Brea, CA) or

Hemavet 950 FS (Drew Scientific, Dallas, TX) and adjusted to 2x10 8 cells/ml unless

otherwise stated.

For isolating washed platelets, following isolation of PRP platelets were

resusupended in PIPES buffered Tyrodes solution (25mM PIPES pH 6.8, 137mM

NaCl, 2.7mM KCl, 2mM MgCl 2, 0.42mM NaH 2PO 4, 5mM glucose, 0.2%BAS),

0.2U/ml apyrase and 100nM carbacyclin.

Murine Platelet Isolation and Preparation

All mice were maintained and housed in a specific pathogen-free facility, and animal procedures were carried out in accordance with the institutional guidelines after the

Temple University Animal Care and Use Committee approved the study protocol.

32 Blood was drawn via cardiac puncture into one-tenth volume of 3.8% sodium citrate.

Blood was then spun at 100 g for 10 minutes and the PRP removed. Red blood cells

were mixed with 400ul 3.8% sodium citrate and spun for a further 10 minutes at

100g. Resulting PRPs were combined, 100nM carbacyclin added and centrifuged for

10 minutes at 400 g. Platelet-poor plasma was removed and the platelet pellet

resuspended in HEPES buffered Tyrodes solution (pH 7.4) containing 0.2U/mL

apyrase. Platelets counts were adjusted using Hemavet 950 FS (Drew Scientific,

Dallas, TX) and adjusted to 2x10 8 cells/ml except for single TULA family deficient

murine platelets which were adjusted to 1x10 8 cells/ml.

Murine Platelet Preparation for Ca 2+ mobilization

Murine blood was drawn as described above and mixed with 4 volumes of PIPES- buffered Tyrode’s solution (pH 6.8) containing 500 M EGTA, 10 M indomethacin and 100nM carbacyclin. The mixture was centrifuged at 100g for 15 minutes at room temperature. The PRP was removed, diluted to 10 ml with the above buffer and centrifuged at 800g for 15 minutes at room temperature. The platelet pellet was resuspended in the above buffer (1ml per mouse) and incubated with 5 M FURA-2

AM for 45 minutes at RT. The platelets were centrifuged at 800g for 10 minutes at

room temperature and finally resuspended in HEPES-buffered Tyrode’s solution. The

platelets were allowed to recover for 15 minutes at room temperature prior to starting

the experiment. A Mark 1 spectrofluorometer (Farrand Optical Company, Valhalla,

NY) with an excitation wavelength of 340 nm and an emission wavelength of 510nm

33 was used to measure fluorescence. The details of Ca 2+ measurement have been previously documented (Quinter et al. 2007). The data in the Results section represents the maximal Ca 2+ concentration minus basal Ca 2+ concentration.

Platelet Activation

300 l or 500 l samples of washed platelets were placed into a lumi-dual aggregometer (Chronolog, Havertown, PA) set at a temperature of 37 oC and stirring rate of 1200rpm. The samples were then treated as described in the results section for each experiment and the change in light transmission recorded using the chart recorder BD41 (Kipp and Zonen, Bohemia, NY) set at 0.2mm/s.

Platelet Dense Granule Secretion Measurements

ATP secretion was monitored using a luciferin-luciferase reagent (DuPont,

Wilmington, DE). Prior to activation of the sample, 1.6mg/ml, final concentration, of the reagent was added.

Lysate Preparation

Following activation, platelets were either treated with one tenth volume 6.6N HClO 4 to precipitate the protein, centrifuged for 10 minutes at 12,000 g at 4 oC, washed with deionized H 20 and resuspended in 1x sample buffer (62.5 mM Tris pH 6.8, 2% SDS,

10% glycerol, 100 mM DTT, 0.01% bromphenol blue) or lysed with 300/500 l 2x

34 RIPA (100mM Tris, pH 7.4, 300mM NaCl, 2% Nonidet P-40, 0.2% SDS, 1%

deoxycholate, 4 mM EDTA [ethylenediaminetetraacetic acid], 2 mM EGTA

[ethylene glycol tetraacetic acid], 2 mM sodium orthovanadate, 20 mM sodium

fluoride, 200 µM phenylmethylsulfonyl fluoride [PMSF] and 20 µg/ml leupeptin) or

2x NP-40 (as 2x RIPA except no SDS or deoxycholate added) lysis buffer, or 2x NP-

40 supplemented with 0.2% SDS, incubated on ice for 10 minutes and centrifuged for

10 minutes at 12,000 g at 4 oC or centrifuged at 100,000g for 30 mins @ 4 oC.

GST-pulldown Assay

Cleared lysates were incubated with 2 g of agarose-coupled GST-TULA-2 H380A

for 1 hour at 4 oC while rocking, or equimolar agarose-coupled GST as a negative

control. After 1 hour, the beads were pelleted and washed three times with 1x NP-40

lysis buffer. 2x sample buffer was then added to the samples, boiled for 5 minutes

and the samples subjected to SDS-PAGE and immunoblotting.

Coimmunoprecipitation

Following lysis with 2x NP-40 supplemented with 0.2% SDS and ultracentrifugation,

platelet lysates were incubated with 2 g/ml of appropriate antibody at 4 oC while

rocking. After 1 hour, 20 l/ml Protein A/G agarose was added to the lysates and incubated overnight at 4 oC with rocking. The next day, agarose beads were washed 5 times for 5 minutes with 4 ml of 1x NP-40 lysis buffer with 0.1% SDS. After washing the beads were resuspended in 2x sample buffer and subject to immunoblotting. 35 Immunoblotting

Prepared proteins were resolved on SDS/polyacrylamide gels and transferred to either

Immobilon-P (Millipore, Billerica, MA) or Nitrocellulose membranes (Whatman,

Piscataway, NJ). Immobilon-P membranes were blocked with 5% Non-fat dry milk

in Tris buffered saline with 0.05 % Tween 20 (TBS-T) and nitrocellulose membranes

were blocked with Odyssey blocking buffer (Licor, Lincoln, NE). Membranes were

probed overnight at 4 oC with the desired primary antibody and then washed four times with TBS-T. Immobilon-P membranes were incubated with HRP-conjugated secondary antibody and nitrocellulose membranes incubated with Infrared dye labelled secondary antibodies for 60 minutes at room temperature and washed four times with TBS-T. Immobilon-P membranes were developed using chemiluminescent substrate (Millipore, Billerica, MA) and visualized using the

Fujifilm LAS-1000 (Fujifilm, Valhalla, NY). Nitrocellulose membranes were developed using the Odyssey imaging system (Licor, Lincoln, NE)

In vitro Kinase Assay

Murine platelets were stimulated with convulxin for 4 minutes and lysed with 2x NP-

40 lysis buffer. Following centrifugation at 12,000g, Syk was immunoprecipitated using anti-Syk clone N-19 polyclonal antibody. After 1 hour, 20 l of protein A/G agarose was added. After another hour, the beads were washed 3 times with 1xNP-40 lysis buffer and once with kinase assay buffer (50mM MOPS pH 7.5, 150mM NaCl,

5mM MgCl 2, 5mM MnCl 2 and 1mM DTT). Immunoprecipitates were then incubated

36 with kinase assay buffer containing 5 g of Tubulin and the reactions started by the addition of 5 l of 25 M ATP containing 5 Ci [ γ-32 P] ATP. Reactions were then

stopped by the addition of 15 l of sample buffer. The samples were boiled and

immunoblotted and Tubulin phosphorylation measured using a Phosphor screen and a

Packard Cyclone Storage Phosphor Screen (Packard Instruments, Meridian, CT).

Tubulin was used as a loading control.

FeCl 3-induced in vivo Thrombosis Injury Model

The collagen-mediated FeCl 3 thrombosis injury model was performed similarly to

Wang et al. (Wang and Xu 2005). Briefly, the left common carotid artery from mice

was exposed and a miniature Doppler flow probe placed around the artery. To induce

injury, the arterial blood flow was stopped and a 1 x 2mm 7.5% FeCl 3-soaked piece

of filter paper was placed on the artery for 2 minutes followed by rinsing with saline.

Blood flow was then resumed and monitored for 30 minutes and the time to occlusion

and thrombus stability noted. A stable thrombus was defined as one that fully

occluded the vessel for at least five minutes.

37 Statistical Analysis

All statistics were calculated using Graphpad Prism . Time courses were analyzed

by Two-way analysis of variance (ANOVA). Dose response curves were fit to a

simple hyperbolic equation and differences between the fits of wild-type and mutant

cells determined using comparisons built into Graphpad Prism . All other data was analyzed using either a Student’s T-test or a One-way ANOVA with Tukey post-hoc test.

38 CHAPTER 3

RESULTS

Syk undergoes substantial tyrosine phosphorylation following stimulation of GPVI to increase its kinase activity and elicit platelet activation. It also undergoes dephosphorylation at later time-points following GPVI stimulation. This dephosphorylation implicates a phosphatase in regulating Syk kinase activity. SHP-

1’s ability to associate with Syk, SHP-2’s reported negative regulatory role in platelets and TULA-2’s reported negative regulatory role in TCR signaling warrants study of these phosphatases as possible negative regulators of Syk in platelet GPVI signaling. In this work we present evidence that rules out involvement of SHP-1 and

SHP-2 and points to TULA-2 as the negative regulator of Syk in platelet GPVI signaling.

SH2-domain Containing Phosphatases

SHP-1 can dephosphorylate Syk in vitro

Pasquet et al. previously showed that, following GPVI stimulation, Syk can associate with SHP-1. Therefore Syk may be a SHP-1 substrate. Additionally, Plas et al. documented that Zap70 could be dephosphorylated by SHP-1 in a T-cell cell-line

(Plas et al. 1996). We therefore investigated the ability of SHP-1 to dephosphorylate

39 Syk. To achieve this, recombinant SHP-1 was incubated with recombinant

phosphorylated Syk for a 1 hour time-course and aliquots taken at various time-

points. Figure 2 shows a Western blot of the samples taken from the time-course and

shows that SHP-1 can cause a reduction in total tyrosine phosphorylation of Syk as

well as dephosphorylate Syk at Y323 and Y525/6 whereas control samples without

SHP-1 show no dephosphorylation of Syk. This suggests that Syk is a substrate for

SHP-1, at least in vitro .

SHP-1 does not coimmunoprecipitate with Syk in platelets

As additional evidence that Syk is a substrate for SHP-1 in platelets the well- established technique of coimmunoprecipitation was employed. Pasquet et al. previously reported that SHP-1 could coimmunoprecipitate with Syk following GPVI stimulation. To confirm this and further evaluate whether SHP-1 is involved in Syk dephosphorylation we performed an immunoprecipitation of Syk from unstimulated and convulxin stimulated platelets and looked for association of SHP-1. Contrary to the Pasquet et al. report, SHP-1 was not present in Syk immunoprecipitates from unstimulated or convulxin stimulated platelets. This result suggests that SHP-1 may not negatively regulate Syk during GPVI signaling.

40

Figure 2. SHP-1 can dephosphorylate Syk in vitro . Samples taken from a phosphatase assay containing SHP-1 and Syk or Syk alone were subjected to SDS-PAGE, blotted and probed for total tyrosine phosphorylation (pY), pY323, pY525/6 and total Syk. Ratios are equal to band intensity for the phosphorylated Syk blots divided by band intensity for the total Syk blot and normalized to the 0 minute timepoint.

41

Figure 3. SHP-1 does not coimmunoprecipitate with Syk in platelets For each sample 1ml of 4x10 8cells/ml were lysed with NP-40, precleared and incubated with 2ug of anti-Syk (4D10) antibody. Immunoprecipitates were Western blotted and probed for Syk and SHP-1.

42 SHP-1 deficient platelet aggregation is similar to wild-type (WT)

A spontaneous mutation in C57BL/J6 mice causes the synthesis of SHP-1 that lacks phosphatase activity. The strain is termed motheaten due to the loss of patches of the animals fur. Life expectancy is approximately 3 weeks. A second spontaneous mutation in the motheaten locus extends life expectancy to approximately 9 weeks and maintains the loss-of-function mutation in SHP-1. These mice are termed motheaten viable (mev) and can be used as a tool for studying SHP-1 deficiency

(Shultz et al. 1984). We isolated platelets from WT and mev mice and monitored platelet aggregation following GPVI stimulation. As shown in figure 4, no difference in platelet aggregation was seen between WT and mev mice suggesting that SHP-1 may not regulate GPVI-mediated platelet functional responses.

GPVI-mediated dense granule secretion is suppressed in SHP-1 deficient platelets

In addition to performing platelet aggregation studies in mev mice we also performed dense granule secretion experiments. Pasquet et al. have shown that α-granule secretion is suppressed in mev mice following GPVI stimulation suggesting that

SHP-1 may be a positive regulator of GPVI signaling. We show a small suppression of dense granule secretion in mev mice compared to wild type which suggests that

SHP-1 acts as a positive regulator of GPVI-mediated dense granule secretion and is in agreement with secretion data published by Pasquet et al (Figure 5).

43 Syk is hypophosphorylated in SHP-1 deficient platelets

Work by Pasquet et al. showed that Syk is hypophosphorylated in mev platelets compared to WT thus suggesting that SHP-1 is a positive regulator of GPVI signaling. We sought to confirm this result by performing similar Syk phosphorylation studies. In the same SHP-1 deficient platelets that the functional studies were performed Syk phosphorylation was also investigated. Figure 6 shows a

Western blot of Syk phosphorylation following GPVI stimulation in WT and mev platelets and shows a hypophosphorylation of Syk at Y519/20 in mev platelets compared to WT. These data therefore confirm the initial observation by Pasquet et al. that SHP-1 is a positive regulator of GPVI signaling.

SHP-1 does not coimmunoprecipitate with c-Cbl in platelets

The reported hyperphosphorylation of Syk observed in c-Cbl deficient platelets suggests a phosphatase may be binding to this adapter protein in order for it to regulate Syk phosphorylation, as c-Cbl lacks a putative phosphatase domain.

Furthermore, Uddin et al. have reported that SHP-1 is able to bind to c-Cbl in other cell systems. To look for a potential interaction between c-Cbl and SHP-1 in platelets we performed a coimmunoprecipitation experiment looking for a complex between c-

Cbl and SHP-1. We observed no association between SHP-1 and c-Cbl in unstimulated or GPVI stimulated platelets suggesting that SHP-1 does not play a role in c-Cbl’s negative regulation of GPVI signaling (Figure 7). This lack of interaction is contrary to previously reported studies in other cell systems however these studies

44 only reported an indirect association between SHP-1 and c-Cbl and if the necessary binding partner is not present in platelets then this may explain a lack of interaction between c-Cbl and SHP-1 (Uddin et al. 1996).

Figure 4. GPVI-mediated platelet aggregation is no different in WT and mev mice. Isolated WT and mev platelets were stimulated with 100ng/ml convulxin and the change in light transmittance measured.

45

Figure 5. GPVI-mediated dense granule secretion is slightly suppressed in mev mice . Isolated WT and mev platelets were stimulated with 100ng/ml convulxin and the dense granule secretion was measured by monitoring ATP release using a luciferin- luciferase reagent.

46

Figure 6. Syk is hypophosphorylated in GPVI-stimulated mev platelets . Isolated platelets from WT or mev mice were stimulated with convulxin for the 30 seconds, the protein precipitated and probed for pY519/20 Syk and total Syk.

47

Figure 7. SHP-1 does not associate with c-Cbl in platelets. For each sample, 1ml of 4x10 8cells/ml were lysed with NP-40, precleared and incubated with 2 g of anti-c-Cbl (C-15). Immunoprecipitates were Western blotted and probed for SHP-1 and c-Cbl.

48 Syk is not a good substrate for SHP-2 in vitro

Given the possible role of SHP-2 in the GPVI-like signaling of the TCR, the reported

association between SHP-2 and Syk in RBL cells, as well as the reported negative

regulatory function of SHP-2 in platelet GPVI signaling we investigated the ability of

SHP-2 to dephosphorylate Syk (Kwon et al. 2005; Patil, Newman, and Newman

2001; Xu and Pecht 2001). Similarly to SHP-1, an in vitro phosphatase approach was first used using recombinant SHP-2 in place of SHP-1. Figure 8 shows a Western blot of samples taken from the in vitro phosphatase assay and shows that SHP-2 does not dephosphorylate Syk as rapidly as the positive control SHP-1. This datum suggests that SHP-2 may not be the phosphatase that negatively regulates Syk in platelets.

Syk does not coimmunoprecipitate with SHP-2

While a less rapid dephosphorylation of Syk in vitro suggests that Syk may not be a substrate for SHP-2, in the intact cell other cofactors may facilitate Syk dephosphorylation by SHP-2. Alternatively, concentration of the two proteins to a specific compartment within the cell may occur for more efficient dephosphorylation of Syk by SHP-2. Therefore we performed an immunoprecipitation of SHP-2 and looked for an association with Syk. Figure 9 shows that Syk does not associate with

SHP-2 in unstimulated or convulxin stimulated platelets. Therefore it is unlikely that

SHP-2 is the phosphatase responsible for the negative regulation of Syk downstream of the GPVI receptor.

49

Figure 8. Syk is not a good substrate for SHP-2 in vitro . Samples taken from a phosphatase assay containing SHP-2 and Syk, Syk alone or SHP-1 and Syk alone were subjected to SDS-PAGE, blotted and probed for total tyrosine phosphorylation (pY), pY525/6 and total Syk. Ratios are equal to band intensity for the phosphorylated Syk blots divided by band intensity for the total Syk blot and normalized to the 0 minute timepoint.

50

Figure 9. Syk does not coimmunoprecipitate with SHP-2. For each sample, 1ml of 4x10 8cells/ml were lysed with NP-40, precleared and incubated with 2 g of anti-SHP-2 (B-1) antibody. Immunoprecipitates were Western blotted and probed for SHP-2 and Syk.

51 SHP-2 does not coimmunoprecipitate with c-Cbl

As discussed above, deletion of c-Cbl causes a hyperphosphorylation of Syk in

GPVI-stimulated platelets compared to WT (Auger et al. 2003; Dangelmaier et al.

2005). The reason for the observed hyperphosphorylation remains unknown but it is reasonable to assume that a phosphatase may be involved. To evaluate SHP-2 as the phosphatase in mediating c-Cbl-dependent negative regulation of Syk phosphorylation an immunoprecipitation of c-Cbl was performed and an association of SHP-2 was probed for. Figure 10 shows the Western blot of the immunoprecipitation and indicates that SHP-2 does not specifically interact with c-

Cbl in platelets. This datum suggests that SHP-2 does not mediate c-Cbl-mediated negative regulation of GPVI signaling. Additionally, it is in disagreement with

Chernock et al. who have shown an association in Jurkat T-cells so this interaction may be a T-cell specific phenomenon (Chernock, Cherla, and Ganju 2001).

These data do not provide strong evidence for SHP-2 being the phosphatase that negatively regulates Syk in GPVI signaling cascade. Since SHP-2 deficiency is embryonic lethal and a transgenic mouse model of SHP-2 deficiency/inactivation is not readily available, SHP-2 was not further pursued as a regulator of Syk in GPVI signaling.

52

Figure 10. SHP-2 does not coimmunoprecipitate with c-Cbl in platelets. For each sample, 1ml of 4x10 8cells/ml were lysed with NP-40, precleared and incubated with 2 g of anti-c-Cbl (C-15) antibody. Immunoprecipitates were Western blotted and probed for SHP-2 and c-Cbl.

53 Summary of SH2 Domain Containing Phosphatases

Previous reports have provided evidence that, in T-cells, SHP-1 is a negative regulator of signaling. However, work in platelets suggests that SHP-1 is a positive regulator. Our studies confirm the studies performed in platelets, therefore suggesting that SHP-1 is not the phosphatase responsible for the negative regulation of Syk in GPVI signaling. Additionally, some reports have suggested that SHP-2 is a negative regulator of TCR signaling and a negative regulator of GPVI-mediated platelet activation. However, our data suggests that SHP-2 is not the phosphatase responsible for negatively regulating Syk in GPVI signaling. Furthermore, our data suggests that SHP-1 nor SHP-2 are responsible for the hyperphosphorylation of Syk seen c-Cbl deficient platelets. Therefore other phosphatases were evaluated.

TULA Family Proteins

In search of an alternative, Dr. Alexander Tsygankov (Department of Microbiology and Immunology, Temple University School of Medicine) suggested that we investigate the role of TULA family proteins in regulation of Syk activation. TULA family proteins have been shown to be negative regulators of signal transduction and

TULA-2 has been shown to have preferential phosphatase activity towards Syk.

They have also been shown to associate with c-Cbl. Therefore TULA proteins may play a critical role in the negative regulation of Syk in GPVI signaling.

54 TULA-2 but not TULA is expressed in human and murine platelets

In order to evaluate the role of the TULA family proteins in platelet GPVI signaling, we investigated whether these proteins are expressed in human and murine platelets using Western blot analysis. Previous reports indicate that TULA-2 is ubiquitously expressed however TULA expression is confined to lymphoid cells (Carpino et al.

2002; Carpino et al. 2004; Feshchenko et al. 2004; Tsygankov 2009; Wattenhofer et al. 2001). Figure 11A shows a Western blot of increasing amounts of human platelet protein and peripheral blood mononucleocytes (PBMCs) probed with antibodies against TULA and TULA-2. The results indicate a notably high expression of

TULA-2 in human platelets compared to PMBCs and no detectable expression of

TULA in human platelets. The doublet for TULA seen in the PBMCs represents both the long and short form splice variants of TULA, which are both found in PBMCs

(Tsygankov 2009). We further analyzed platelet lysates from WT mice and mice deficient in both TULA family members (TULA-dKO) as well as HEK293T cells overexpressing each family member. As shown in Figure 11B, TULA-2 is clearly present in the WT platelets and not in TULA-dKO platelets, whereas no TULA is detectable in either set of murine platelets. These data suggest that TULA-2 is the only TULA family member expressed in human and murine platelets.

55 To ensure that deletion of TULA family members had no effect on relative protein

expression of major GPVI signaling molecules a Western blot was performed of

GPVI, Syk and PLC γ2 and shows no difference in the protein expression (figure

11C). Therefore, any enhanced signaling or platelet functional responses documented below for the TULA-dKO platelets were not due increased expression of GPVI, Syk or PLC γ2.

Figure 11. TULA-2 but not TULA is expressed in human and murine platelets and deletion of TULA proteins does not affect protein expression. (A) Increasing amounts of human platelet and peripheral blood mononucleocyte protein were blotted and probed for TULA family members or the loading control Erk. (B) TULA family protein expression is compared in WT and TULA-dKO murine platelets with HEK293T cells overexpressing each member as a positive control. (C) Relative expression of GPVI, Syk and PLC γ2 are compared between WT and TULA-dKO murine platelets.

56 Synthesis of GST-TULA-2 and GST-TULA H380A

It has previously been reported that the catalytic domain of TULA-2 can dephosphorylate Syk but no report has been made regarding the full-length protein

(Agrawal, Carpino, and Tsygankov 2008). Therefore, we utilized the baculovirus expression vector system to synthesize full-length GST-TULA-2 to evaluate its function. Additionally, we synthesized full-length phosphatase inactive GST-TULA-

2 (TULA-2 H380A) for use in GST-pulldown assays as phosphatase inactive mutants can be used a substrate traps to identify substrates (Blanchetot et al. 2005). To achieve this, pAcGHLT-B vector containing either the TULA-2 gene or TULA-2

H380A gene were cotransfected into Sf9 cells with linearized baculovirus DNA containing a GFP tag. Following successful transfection, the baculovirus was amplified three times by sequentially infecting more cells to increase the viral titer.

After amplification, the Sf9 cells were then infected with a high concentration of baculovirus to induce larger scale protein synthesis for protein purification. TULA-2 and TULA-2 H380A were then purified from lysed Sf9 cells using glutathione- agarose. The proteins were then left conjugated to agarose or eluted using glutathione. Figure 12A shows a fluorescence microscopic image of untransfected cells to indicate the background level of fluorescence. Figure 12B shows an overlay of the visible frame (to indicate the presence of the Sf9 cells) with the fluorescence image. Figure 12C shows a fluorescence image of Sf9 cells transfected with 1 g of pAcGHLT-B containing TULA-2. Figure 12D shows the overlay with the visible frame showing successful cotransfection and synthesis of TULA-2. Figure 12E shows a Gelcode blue stained gel of purified TULA-2 and a BSA standard curve to 57 show successful synthesis and purification of TULA-2 and figure 12F shows a

Western blot probing for TULA-2 to show successful synthesis of TULA-2 protein.

Thus, we were able to use agarose-conjugated or eluted GST-TULA-2 and GST-

TULA-2 H380A as tools to evaluate TULA-2’s possible role in platelets. TULA was not synthesized due to its apparent absence of expression in platelets (Figure 11).

58

Figure 12. TULA-2 can be successfully expressed in Sf9 cells and purified to homogeneity. Sf9 cells were cotransfected with pAcGHLT-B-TULA-2 and linearized baculovirus DNA or left untreated. (A) Fluorescence image of untransfected cells. (B) Overlay of fluorescence image with visible frame of untransfected cells. (C) Fluorescence image of Sf9 cells cotransfected with 1 g of pAcGHLT-B-TULA-2. (D) Overlay of fluorescence image with visible frame of pAcGHLT-B-TULA-2 transfected cells. (E) Gelcode blue treated SDS-PAGE gel of purified GST-TULA-2 and BSA standards, E denotes elution fractions. (F) Western blot of purified TULA-2. E denotes eluted fractions, SB denotes 2X sample buffer treated glutathione agarose beads, AC denotes glutathione agarose-conjugated fraction.

59 TULA-2 can dephosphorylate Syk

The ability of the catalytic domain of TULA-2 to dephosphorylate Syk in vitro as

well as Zap70 in vitro has previously been reported (Agrawal, Carpino, and

Tsygankov 2008; Mikhailik et al. 2007). However, no reports exist using the full- length protein. We therefore evaluated the ability of full-length TULA-2, synthesized in Sf9 cells, to dephosphorylate Syk. As for the SHP-1 and SHP-2 phosphatase assays we incubated full-length TULA-2 with phosphorylated Syk, sampling at specific time-points and Western blotting to probe for phosphorylated Syk. As shown in Figure 13, full-length TULA-2 caused a reduction in the total tyrosine phosphorylation of Syk as well as dephosphorylation at Y525/6 in the activation loop of Syk. These data confirm the results of Agrawal et al. and suggest that Syk is a substrate for TULA-2.

TULA-2 can associate with Syk and c-Cbl in platelets (GST-pulldowns)

Previous work has shown that TULA-2 can coimmunoprecipitate with Syk in

HEK293T cells overexpressing Syk and TULA-2 (Agrawal, Carpino, and Tsygankov

2008). It has also been reported that TULA family proteins can bind to c-Cbl

(Kowanetz et al. 2004). Therefore, to provide further evidence that Syk could be a substrate for TULA-2 and that TULA-2 could be mediating the Syk hyperphosphorylation phenotype in c-Cbl deficient platelets we performed GST- pulldowns looking for an association between Syk, c-Cbl and TULA-2 in platelets.

Unstimulated and convulxin-stimulated precleared platelet lysates were incubated

60 with agarose-coupled full-length GST-TULA-2 H380A or GST-TULA-2. The resulting pulldowns were then probed for Syk and c-Cbl. Figure 14A shows the

Western blots from the GST-pulldown using TULA-2 H380A and shows constitutive association between Syk and TULA-2 and between TULA-2 and c-Cbl, that increases following stimulation of GPVI, suggesting that a complex may form between TULA-

2, Syk and c-Cbl in platelets. Theses data are only qualitative however, so to quantitate these data, mean band intensities from 3 experiments for Syk and c-Cbl were calculated and plotted as a histogram. As shown in figure 14B, there is a statistically significant association between Syk and TULA-2 H380A in both unstimulated and convulxin stimulated platelets. This effect is mirrored in c-Cbl association with TULA-2 H380A (figure 14C). Performing the same studies using

WT TULA-2 the same pattern of association was seen (figure 14D), demonstrating that the association is not non-specific association only observed when using the phosphatase inactive TULA-2.

TULA-2 can associate with Syk and c-Cbl in platelets (Coimmunoprecipitations)

While GST-pulldown studies may be indicative of protein-protein interactions a major caveat is that it involves the addition of an exogenous protein that may not reflect true protein-protein interactions inside the cell. Thus, coimmunoprecipitation studies were also performed to ascertain whether the protein complexes seen in the

GST-pulldowns could occur with endogenously expressed proteins. Precleared lysates from unstimulated and convulxin stimulated proteins were incubated with an

61 anti-TULA-2 antibody and the associated proteins probed for on a Western blot.

Figure 15A shows these Western blots and shows that Syk and c-Cbl can

coimmunoprecipitate with TULA-2 in unstimulated and convulxin-stimulated platelets, confirming the observations made in the GST-pulldown assay. When these data were quantitated, similarly to the pulldown experiments, a statistically significant association of c-Cbl and TULA-2 was observed, however the data for Syk are not statistically significant which could be attributable to the high level of non-specific binding of Syk (Figures 15B and C). In addition, in the unstimulated and convulxin- stimulated groups Syk was phosphorylated as indicated by probing the Syk blot for pY525/6 Syk.

The GST-pulldown and coimmunoprecipitation experiments suggest that a complex exists between TULA-2, Syk and c-Cbl, which may allow TULA-2 to negatively regulate Syk phosphorylation downstream of GPVI signaling. The presence of c-Cbl in this complex may also explain the Syk hyperphosphorylation seen in c-Cbl deficient platelets. This possibility is considered further in the discussion section.

62

Figure 13. TULA-2 dephosphorylation of Syk. Active TULA-2 was purified from Sf9 cells and incubated with phosphorylated Syk for the indicated times, Western blotted and probed for phosphotyrosine, pY525/6 Syk and total Syk. Blots are representative of 3 experiments. Ratios are equal to band intensity for the phosphorylated Syk blots divided by band intensity for the total Syk blot and normalized to the 0 minute timepoint.

63

Figure 14. Syk and c-Cbl associate with GST-TULA-2 H380A and GST-TULA- 2. (A) For each sample, 2ml of 8x10 8 cells/ml were lysed with NP-40 supplemented with 0.1% SDS, precleared and incubated with GST-TULA-2 (H380A) or GST. Pulldowns were Western blotted and probed for c-Cbl, Syk and TULA-2. All blots are representative of 3 experiments (B) Mean band densities for Syk from 3 experiments were normalized to TULA-2 15 second time point, plotted as a histogram and a one-way ANOVA performed (P< 0.05) (C) Mean band densities for c-Cbl from 3 experiments were normalized to TULA-2 15 second time point, plotted as a histogram and a one-way ANOVA performed (P< 0.05). (D) As part A except GST-TULA-2 was used.

64

Figure 15. Syk and c-Cbl can associate with TULA-2. (A) For each sample, 2ml of 8x10 8 cells/ml were lysed with NP-40 supplemented with 0.1% SDS, precleared and incubated with 4 g of anti-TULA-2 antibody. Immunoprecipitates were Western blotted and probed for pY525/6 Syk, Syk and c- Cbl. All blots are representative of 3 experiments. (B) Mean band densities for Syk from 3 experiments were normalized to TULA-2 15 second time point, plotted as a histogram and a one-way ANOVA performed (B) Mean band densities for c-Cbl from 3 experiments were normalized to TULA-2 15 second time point, plotted as a histogram and a one-way ANOVA performed (P< 0.0001).

65 Syk is hyperphosphorylated in TULA-dKO platelets in a time dependent manner

Deletion of both TULA family members causes the hyperphosphorylation of Zap70

following TCR stimulation (Carpino et al. 2004). Additionally, it has been shown

that Zap70 and Syk could be TULA-2 substrates as the catalytic domain of TULA-2

can dephosphorylate both proteins in vitro (Agrawal, Carpino, and Tsygankov 2008;

Mikhailik et al. 2007). We hypothesized that TULA-2 may therefore have a role in negatively regulating Syk activity in platelet GPVI signaling and proposed to study the effect TULA-2 ablation on Syk regulation in murine platelets as a means to achieve this. Therefore, we studied Syk phosphorylation in TULA-2 deficient platelets. These studies were first performed in TULA-dKO murine platelets. As

TULA does not appear to be expressed in platelets (figures 11A and 11B) the effects observed are most likely due to the lack of TULA-2 in these mice.

A time-course of Syk phosphorylation using convulxin as agonist is shown in figure

16A. In WT platelets an increase in Syk phosphorylation was observed following activation, peaking at thirty seconds, followed by a dephosphorylation of Syk.

However, in TULA-dKO platelets, a hyperphosphorylation of Syk was seen at all time points. In addition, the TULA-dKO platelets lacked an apparent dephosphorylation seen at later time points in the WT platelets. To quantitate these data, mean band intensity for pY525/6 Syk from 3 experiments was calculated and plotted against time (figure 16B). Analysis by two-way ANOVA indicates a statistically significant potentiation of Syk phosphorylation in TULA-dKO platelets

66 compared to WT, confirming an apparent lack of regulation of Syk phosphorylation in TULA-dKO platelets.

To more fully investigate this apparent lack of Syk regulation in TULA-dKO platelets, a time-course was also performed using CRP as the agonist. Similar to the observations of the convulxin time-course, a potentiation of Syk phosphorylation is apparent in TULA-dKO platelets compared to WT (figure 16C). Statistical analysis by two-way ANOVA of 3 replicate experiments reveals a statistically significant potentiation of Syk phosphorylation in TULA-dKO platelets (figure 16D).

A time-course was also performed using the physiological agonist collagen which causes an increase in Syk phosphorylation in both WT and TULA-dKO platelets up to 4 minutes. As observed with convulxin and CRP, a hyperphosphorylation of Syk was seen in the TULA-dKO platelets.

67

Figure 16. Syk is hyperphosphorylated in TULA-dKO platelets in a time dependent manner. (A) Isolated WT and TULA-dKO platelets were stimulated with 200ng/ml convulxin for 0, 30, 60 120 and 240 seconds, the protein precipitated and pY519/20 Syk and total Erk were probed for. (B) Mean relative Syk phosphorylation of 3 experiments for convulxin in WT ( λ) and TULA-dKO ( ) platelets was plotted against time and a Two-way ANOVA performed (P <0.0001 comparing WT v dKO). (C) As part A except platelets were stimulated with 1 g/ml CRP and total Syk was used as a loading control. (D) As part B except CRP used as agonist (P <0.0001 comparing WT v dKO). (E) As part A except platelets were stimulated with 50 g/ml collagen.

68 Syk is hyperphosphorylated in TULA-dKO platelets in a dose dependent manner

In addition to the time course of Syk phosphorylation we measured dose-response

relationships for convulxin, CRP and collagen-mediated Syk phosphorylation in WT

and TULA-dKO platelets. These were performed to evaluate whether the impaired

Syk dephopshorylation phenotype seen in the time-course experiment was both

concentration-dependent and whether higher concentrations of agonists would overcome the difference between WT and TULA-dKO platelets. Figure 17A shows the Syk phosphorylation dose-response for convulxin taken at a 2 minute time-point and indicates that Syk phosphorylation is potentiated across all concentrations of convulxin in the TULA-dKO murine platelets. Plotting the mean Syk phosphorylation from 3 experiments against agonist concentration and fitting this with a non-linear regression it was found that these curves fits were significantly different (figure 17A).

We also repeated the dose-response with CRP. As shown in figure 17C, Syk is also hyperphosphorylated at 2 minutes at all concentrations of CRP in the TULA-dKO platelets compared to WT. Quantitation of the mean Syk phosphorylation from 3 experiments and plotting against dose revealed a statistically significant increase in

Syk phosphorylation in TULA-dKO platelets compared to WT (figure 17D).

69 Performing a dose-response study with the physiological agonist collagen also showed a potentiation of Syk phosphorylation in TULA-dKO platelets indicating that the hyperphosphorylation occurs with the natural agonist and not just synthetic agonists to GPVI (figure 17E).

The time- and dose-dependent aberrant Syk phosphorylation observed here in TULA- dKO platelets suggests that TULA-2 could be the phosphatase responsible for dephosphorylating and regulating Syk in platelets. The potentiated Syk phosphorylation in the TULA-dKO platelets also suggests that only a minor role for

SHP-1 or SHP-2, if any, in Syk dephosphorylation is evident. If either SHP-1 or

SHP-2 were profoundly dephosphorylating Syk this would be evident even in the absence of TULA-2. However, we observed a substantial potentiation of Syk phosphorylation when TULA-2 is absent.

Syk kinase activity is enhanced in TULA-dKO platelets

While an increase in Syk phosphorylation at tyrosines 519 and 520 (525 and 526 in human Syk) is usually taken as evidence of increased kinase activity, we took this a step further by performing in vitro kinase assays using Syk immunoprecipitated from

WT and TULA-dKO platelets. This was to test whether Syk immunoprecipitated from TULA-dKO platelets exhibited increased kinase activity compared to Syk from

WT platelets. To maximize the sensitivity of the kinase assay, platelets were treated with convulxin for 4 minutes prior to lysis and immunoprecipitation. The activity of 70 Syk was assessed by the phosphorylation of a known substrate, tubulin (Dangelmaier et al. 2005). Figure 18 shows a representative time course from 5 independent experiments in which tubulin phosphorylation by Syk was monitored. An increased rate of tubulin phosphorylation by Syk immunoprecipitated from TULA-dKO platelets compared to WT platelet is evident. This suggests that TULA-2 negatively regulates Syk kinase activity in GPVI stimulated platelets and confirms that phosphorylation of Y519/520 reflects Syk activity in platelets.

71

Figure 17. Syk is hyperphosphorylated in TULA-dKO platelets in a dose dependent manner. (A) Isolated WT and TULA-dKO platelets were stimulated for 2 minutes with 50, 100, 200, 400 or 800ng/ml convulxin, processed for Western blotting and probed for pY519/20 Syk and total Syk. (B) Mean relative Syk phosphorylation of 3 experiments for convulxin in WT ( λ) and TULA-dKO ( ) platelets was plotted as a function of dose and curves fitted using a non-linear regression (P < 0.0001 comparing WT v dKO). (C) As part A except platelets were stimulated with 0.5, 1, 2, 5 or 10 g/ml CRP. (D) As part B except CRP was used as agonist (P = 0.0009 comparing WT v dKO). (E) As part A except platelets were stimulated for 1 minute with 25, 50 or 100 g/ml collagen.

72

Figure 18. Syk is more active in TULA-dKO platelets than WT platelets. Isolated WT and TULA-dKO platelets were left untreated or treated with 200ng/ml convulxin for 4 minutes. Kinase activity of the resulting immunoprecipitates was then measured and plotted as function of time. WT ( λ) and TULA-dKO () platelets. Shown is a representative plot from 5 independent experiments.

73 PLC γ2 is hyperphosphorylated in TULA-dKO platelets in a time dependent manner

PLC γ2 is the phospholipase C isoform downstream of Syk activation and responsible for mobilization of calcium and platelet activation downstream of GPVI (Blake et al.

1994; Daniel, Dangelmaier, and Smith 1994; Ozdener et al. 2002). Thus, we assessed the phosphorylation of PLC γ2 tyrosine 759, an important residue for its activation, in

WT and TULA-dKO platelets to see if increased Syk phosphorylation caused a concurrent increase in PLC γ2 phosphorylation (Ozdener et al. 2002; Watanabe et al.

2001).

A time-course of Syk phosphorylation was first performed using convulxin.

Similarly to the Syk phosphorylation profile observed with convulxin, PLC γ2 undergoes phosphorylation peaking at 30 seconds followed by a dephosphorylation at later time-points in WT platelets. However, in TULA-dKO platelets PLC γ2 is hyperphosphorylated at all time-points and does not undergo dephosphorylation at later time-points (figure 19A).

A time-course was next performed using CRP as the agonist and, as shown in figure

19B, a potentiation of PLC γ2 phosphorylation occurs in TULA-dKO platelets at all time-points compared to WT platelets.

74 Performing the same time-course except using the physiological agonist collagen, a

potentiation of PLC γ2 phosphorylation is evident in TULA-dKO platelets at all time-

points (figure 19C).

Figure 19. PLC γγγ2 is hyperphosphorylated in TULA-dKO platelets in a time dependent manner. (A) Isolated WT and TULA-dKO platelets were stimulated with 200ng/ml convulxin for 0, 30, 60, 120 and 240 seconds, the protein precipitated, Western blotted and pY759 PLC γ2 and total PLC γ2 probed for. (B) As part A except platelets were stimulated with 1 g/ml CRP (C) As part A except platelets were stimulated with 50 g/ml collagen. Convulxin and CRP blots are representative of 3 experiments. Ratios are equal to band intensity for the phosphorylated PLC γ2 blots divided by band intensity for the total PLC γ2 blot and normalized to the 0 minute timepoint in the WT platelets.

75

PLC γ2 is hyperphosphorylated in TULA-dKO platelets in a dose dependent manner

Concurrently with time-course studies of PLC γ2 phosphorylation, dose-response studies were also undertaken to more fully evaluate the effect of TULA-2 deletion on

PLC γ2 phosphorylation. Figure 20A shows the dose-response taken at a 2 minute time-point for PLC γ2 phosphorylation with convulxin and shows a potentiation of phosphorylation across all doses in TULA-dKO platelets.

A similar pattern of phosphorylation is seen with a CRP dose-response. PLC γ2

phosphorylation is potentiated across all doses of CRP in the TULA-dKO platelets

(figure 20B). This potentiation of PLC γ2 phosphorylation in TULA-dKO platelets was also observed when using the physiological agonist collagen (figure 20C).

The time-course and dose-response data for PLC γ2 phosphorylation shown above suggests that a possible lack of regulation of Syk phosphorylation by TULA-2 causes a potentiation of phosphorylation of downstream signaling proteins. This may cause enhanced GPVI signaling and GPVI-mediated platelet functional responses.

76

Figure 20. PLC γ2γ2γ2 is hyperphosphorylated in TULA-dKO platelets in a dose dependent manner. (A) Isolated WT and TULA-dKO platelets were stimulated for 2 minutes with 50, 100, 200, 400 or 800ng/ml convulxin, processed for Western blotting and probed for pY759 PLC γ2 and total PLC γ2 probed for. (B) As part A except platelets were stimulated with 0.5, 1, 2, 5 or 10 g/ml CRP. (C) As part A except platelets were stimulated with 25, 50 or 100 g/ml collagen. Convulxin and CRP blots are representative of 3 experiments. Ratios are equal to band intensity for the phosphorylated PLC γ2 blots divided by band intensity for the total PLC γ2 blot and normalized to the 0 minute timepoint in the WT platelets.

77 Ca 2+ mobilization is potentiated in TULA-dKO platelets

Ca 2+ mobilization is a critical event in the activation of platelets therefore

measurement of this event can provide insight into extent of platelet activation. As

we observed hyperphosphorylation of PLC γ2, which is thought to be a reflection of increased enzyme activity, we resolved to also measure Ca 2+ mobilization to see if it

was potentiated in TULA-dKO platelets. FURA-2-AM loaded WT and TULA-dKO

platelets were stimulated with multiple doses of CRP and the Ca2+ mobilization

measured in a fluorometer. A dose-response curve was then constructed from the

Ca 2+ measurements and the data fitted using a non-linear regression. Figure 21 shows

the constructed dose-response curve and indicates a potentiation of Ca 2+ mobilization

in TULA-dKO platelets compared to WT. Statistical analysis of the curve fits

showed a statistically significant difference between the groups (P = 0.0001).

Therefore, a suspected lack of regulation of Syk during GPVI signaling due to

TULA-2 deletion also causes a potentiation of Ca 2+ mobilization. This suggests that

TULA-2 is an important negative regulator of GPVI-mediated platelet activation.

78

Figure 21. Ca 2+ mobilization is potentiated in TULA-dKO platelets. FURA-2-AM loaded murine platelets were stimulated with varying doses of CRP in the presence of 100 M 2MeSAMP, 10 M MRS2719, 150nM Echistatin, 10 M Indomethacin and 2mM Probenecid and the Ca 2+ mobilization measured using a spectrofluorometer. The change in intracellular calcium was then measured, plotted against dose of CRP and fitted with a hyperbolic curve. Data points are constructed from 3 independent experiments.

79 GPVI-mediated platelet aggregation is enhanced in TULA-dKO platelets

Previous reports have shown that deletion of both TULA family members causes a

hyperproliferative functional response it T-cells, which can be rescued by the

reexpression of TULA-2 (Mikhailik et al. 2007). Additionally, having observed

enhanced GPVI signaling in TULA-dKO platelets we also sought to ascertain

whether the increased GPVI signaling lead to increased platelet function. The first

functional response measured was platelet aggregation. This measures the transition

of the αIIb β3 to its high affinity state allowing the cross-linking of platelets by fibrinogen, which is a measure of platelet activation.

Figure 22A shows the aggregation traces for convulxin stimulated WT and TULA- dKO platelets for 100ng/ml and 200ng/ml doses. At 100ng/ml full aggregation is seen in the TULA-dKO platelets while only shape change is observed the WT platelets. At 200ng/ml both WT and TULA-dKO platelets aggregate fully.

Therefore, this aggregation defect seen at lower doses is surmountable.

The same experiments were then performed using CRP to confirm the convulxin data.

Figure 22B shows the aggregation tracings and shows at lower doses of CRP TULA- dKO platelets aggregate fully but WT platelets only undergo shape change. At higher doses of CRP this proaggregatory response is overcome as observed with convulxin.

80 Taken together, the convulxin and CRP aggregation data suggest that TULA-dKO

platelets are hyperresponsive to GPVI stimulation. Therefore, the enhanced signaling

seen in TULA-dKO platelets appears to have functional consequences and implicates

TULA-2 as a negative regulator of GPVI signaling.

GPVI-mediated dense granule secretion is enhanced in TULA-dKO platelets

The second platelet functional response measured was dense granule secretion.

Platelets secrete dense granules following activation to amplify the number of

platelets activated in response to injury. In the dense granules platelets store ATP,

amongst other prothrombotic autocoids, and its release can be measured using a

luciferin-luciferase based reagent which fluoresces when ATP bound.

Dense granule secretion was first measured in convulxin stimulated WT and TULA-

dKO platelets and, as shown in figure 23A, it was potentiated in TULA-dKO platelets

at 50, 100 and 200ng/ml. Performing the same experiments except using CRP as the

agonist, dense granule secretion was potentiated in TULA-dKO platelets at the 3

doses used (figure 23B).

Consistent with the proaggregatory response seen with convulxin and CRP, a prosecretory phenotype is also observed, again suggesting that the enhanced signaling seen in TULA-dKO platelets leads to enhanced platelet functional responses and further implicates TULA-2 as a negative regulator of GPVI signaling. 81

Figure 22. GPVI-mediated platelet aggregation is potentiated in TULA-dKO platelets. (A) Isolated WT and TULA-dKO platelets were stimulated with 100ng/ml or 200ng/ml convulxin and the change in light transmittance observed. (B) As part A but platelets were stimulated with 1, 2 or 4 g/ml CRP. All traces are representative of 3 experiments.

82

Figure 23. GPVI-mediated platelet dense granule secretion is enhanced in TULA-dKO platelets. (A) Isolated WT and TULA-dKO platelets were stimulated with 50, 100, 200ng/ml convulxin and the dense granule secretion was measured by monitoring ATP release using a luciferin-luciferase reagent. (B) As part A but platelets were stimulated with 1, 2 or 4 g/ml CRP. All traces are representative of 3 experiments. 83 PAR4-mediated platelet aggregation is not affected in TULA-dKO platelets

To further validate the role TULA-2 in regulating Syk during GPVI-mediated platelet

functional responses platelet aggregation was measured following activation of the

thrombin receptor, PAR4. This served as a negative control as Gq mediated Ca 2+

mobilization is the primary mechanism of platelet activation induced by this GPCR

(Woulfe 2005). Stimulation of WT and TULA-dKO platelets with the 300 M or

1000 M of the PAR4 agonist AYPGKF revealed no difference in platelet aggregation as shown in figure 24. This suggests that TULA-2 mediated negative regulation of platelet signaling is specific to GPVI or signaling cascades in which Syk in critically involved.

PAR4-mediated dense granule secretion is not affected in TULA-dKO platelets

In addition to studying platelet aggregation downstream of the PAR4 receptor, dense granule secretion was also studied. As shown in figure 25, no difference was seen in dense granule secretion at 300 M and at 1000 M a small reduction in secretion was

observed. These data suggest that TULA-2 has no role or may suppress dense

granule secretion downstream of the PAR4 receptor.

The lack of any real PAR4-mediated platelet functional phenotype in TULA-dKO

platelets suggests that TULA-2 plays no role in regulating PAR4 signaling, as

expected.

84

Figure 24. PAR4-mediated platelet aggregation is not affected in TULA-dKO platelets. Isolated WT and TULA-dKO platelets were stimulated with 300 M or 1000 M AYPGKF and the change in light transmittance observed.

85

Figure 25. PAR4-,mediated dense granule secretion is not affected in TULA- dKO platelets. Isolated WT and TULA-dKO platelets were stimulated with 300 or 1000M AYPGKF and the dense granule secretion was measured by monitoring ATP release using a luciferin-luciferase reagent.

86 TULA-dKO mice are prothrombotic

As functional responses in the platelets isolated from TULA-dKO are enhanced in

response to GPVI agonists, we evaluated the in vivo physiological implications of

these enhanced responses. To evaluate this, the well-established FeCl 3–induced

thrombosis injury model was employed (Wang and Xu 2005). Damage to the

endothelial layer by FeCl 3 leads to the exposure of sub-endothelial collagen so is an appropriate model to use to test enhanced GPVI signaling (Dubois et al. 2006).

Isolated carotid arteries from WT and TULA-dKO mice were injured with 7.5%

FeCl 3 for two minutes and the time to occlusion and the thrombus stability measured.

The time to occlusion for WT and TULA-dKO mice is shown in figure 26A and the results show a statistically significant shorter time to occlusion in the TULA-dKO mice (P < 0.05). Figure 26B is a plot of thrombus stability and shows an enhanced thrombus stability in TULA-dKO mice compared to WT, with 87% of mice forming stable thrombi in the TULA-dKO mice and only 33% in the WT. These data indicate that TULA-2 has a physiologically important function in vivo to negatively regulate thrombus formation and stability.

87

Figure 26. TULA-dKO mice are prothrombotic. WT and TULA-dKO mice were subjected to injury of the left carotid artery by 7.5% FeCl 3 for 2 minutes and the time to occlusion and thrombus stability noted. (A) is a plot of time to occlusion for each group of mice. n=12 for each group. A mean of 16.5 minutes +/- 3 minutes for WT and a mean of 7.37 minutes +/- 0.76 minutes for TULA-dKO was calculated. Statistical analysis was performed using an unpaired T- test. (B) is a plot of thrombus stability, defined as a complete occlusion for at least five minutes.

88 Syk is hyperphosphorylated in TULA-2 deficient platelets in a time dependent manner

Although we detected little or no TULA in human or murine platelets we cannot rule

out the possible role of an undetectable amount of TULA that may be affecting the

phenotype of the TULA-dKO murine platelets. Also, Carpino et al. showed that

deletion of both family members was required to see a phenotype downstream of the

TCR. Therefore, we performed similar Syk phosphorylation studies to those outlined

for the TULA-dKO platelets in TULA (-/-) or TULA-2(-/-) platelets.

A time-course using convulxin as the agonist was first performed in WT, TULA (-/-) and TULA-2(-/-) platelets and is shown in figure 27A. WT platelets show a phosphorylation of Syk followed by a dephosphorylation at later time-points. The

TULA-2(-/-) platelets however show a potentiation of Syk phosphorylation compared to WT. The phosphorylation profile in TULA (-/-) platelets is similar to that of WT platelets.

A time-course was also performed using CRP in all three sets of mice. As shown in figure 27B, Syk is hyperphosphorylated in TULA-2(-/-) platelets compared to WT.

Additionally, no difference between Syk phosphorylation is apparent between WT and TULA (-/-) platelets. Taken together with the convulxin data, these data confirm that TULA-2 is the TULA family member responsible for the negative regulation of

Syk in GPVI signaling.

89

Figure 27. Syk is hyperphosphorylated in TULA-2 deficient platelets in a time dependent manner. (A) Isolated WT, TULA (-/-) and TULA-2(-/-) platelets were stimulated with 100ng/ml convulxin for 0, 30, 60 120 and 240 seconds, the protein precipitated and pY519/20 Syk and total Syk were probed for. (B) As part A except platelets were stimulated with 10 g/ml CRP. All blots are representative of 2 experiments.

90 Syk is hyperphosphorylated in TULA-2 deficient platelets in a dose dependent manner

To further confirm the time-course data above, dose-response experiments were also

performed with convulxin and CRP. Figure 28A shows the dose-response for

convulxin and demonstrates that Syk is hyperphosphorylated in the TULA-2(-/-) platelets whereas the TULA (-/-) Syk phosphorylation is similar to that of WT. This phenomenon was also apparent when the experiment was performed using CRP

(figure 28B). These data and the time-course data recapitulate the studies performed in the TULA-dKO mice and further suggest that TULA-2 is the TULA family member responsible for the negative regulation of Syk in GPVI-mediated platelet activation. As discussed above, they also suggest little or no role for SHP-1 or SHP-2 in Syk dephosphorylation as substantial Syk phosphorylation is also observed TULA-

2(-/-) platelets.

91

Figure 28. Syk is hyperphosphorylated in TULA-2 deficient platelets in a dose dependent manner. (A) Isolated WT, TULA (-/-) and TULA-2(-/-) platelets were stimulated for 2 minutes with 50, 100, 200, 400 or 800ng/ml convulxin, processed for Western blotting and probed for pY519/20 Syk and total Syk. (B) As part A except platelets were stimulated with 1, 2, 10 or 20 mg/ml CRP.

92 GPVI-mediated platelet aggregation is enhanced in TULA-2 deficient platelets

In addition to performing Syk phosphorylation studies in TULA-2 and TULA

deficient platelets we performed platelet functional studies to confirm the

observations made in the TULA-dKO mice. Figures 29A shows that aggregation is

enhanced in TULA-2 deficient platelets in response to convulxin at 100 and 200ng/ml

but is a surmountable defect as no difference is seen at 400ng/ml. TULA (-/-) platelet

aggregation was no different to that of WT platelets. Performing similar experiments

with CRP as the agonist shows a similar pattern of a potentiation of aggregation in

TULA-2(-/-) platelets that is surmountable at higher doses of agonist (figure 29B).

Additionally, no effect on aggregation is observed in TULA (-/-) platelets compared to

WT.

GPVI-mediated dense granule secretion is enhanced in TULA-2 deficient platelets

Platelet dense granule secretion was also studied in the three groups of platelets to

further validate the functional experiments performed in TULA-dKO mice. As

shown in figure 30A, dense granule secretion is potentiated in TULA-2(-/-) platelets

but not TULA (-/-) platelets compared to WT. As observed for the aggregation studies,

a similar phenotype was seen with CRP (figure 30B). Taken together with the

aggregation experiments, these studies utilizing TULA single knockout mice indicate

that TULA-2 is the TULA family member responsible for the negative regulation of

GPVI-mediated platelet activation and TULA does not appear to play a role.

93

Figure 29. GPVI-mediated platelet aggregation is enhanced in TULA-2 deficient platelets. (A) Isolated WT and TULA-dKO platelets were stimulated with 100ng/ml, 200ng/ml or 400ng/ml convulxin and the change in light transmittance observed. (B) As part A but platelets were stimulated with 10 or 20 g/ml CRP.

94

Figure 30. GPVI-mediated dense granule secretion is enhanced in TULA-2 deficient platelets. (A) Simultaneous to platelet aggregation studies dense granule secretion was measured by monitoring ATP release using a luciferin-luciferase reagent. 100, 200 or 400ng/ml convulxin was used. (B) As part A but platelets were stimulated with 10 or 20 g/ml CRP.

95 Phosphatases role in the c-Cbl-mediated negative regulation of Syk

Auger et al. and Dangelmaier et al. have previously reported that c-Cbl is a negative regulator of GPVI signaling and that Syk is hyperphosphorylated in c-Cbl deficient platelets. In addition, c-Cbl is able to function as an adapter protein to regulate cell function and has been reported to bind Syk, TULA family proteins, SHP-2 and indirectly to SHP-1 (Chernock, Cherla, and Ganju 2001; Kowanetz et al. 2004;

Tsygankov et al. 2001; Uddin et al. 1996). c-Cbl then may be able to colocalize Syk and a phosphatase to allow dephosphorylation of Syk and may serve to explain the phenotype in c-Cbl deficient platelets.

The data presented in figures 7 and 10 suggest that SHP-1 and SHP-2 do not associate with c-Cbl, however TULA-2 can associate with c-Cbl in platelets (Figure 14 and

15). We therefore investigated the possible role of TULA-2 in mediating the c-Cbl- dependent negative regulation of Syk in GPVI signaling.

Syk is hyperphosphorylated in c-Cbl deficient platelets

As previously reported, deletion of c-Cbl causes a hyperphosphorylation of Syk in platelets stimulated with GPVI agonists (Auger et al. 2003; Dangelmaier et al. 2005).

We sought to confirm these data by performing studies of Syk phosphorylation in c-

Cbl deficient platelets and comparing this to WT. Figure 31A shows a time-course of

Syk phosphorylation following stimulation of WT or c-Cbl deficient platelets with convulxin. Here we see that Syk is hyperphosphorylated compared to WT platelets. 96 Additionally, while Syk undergoes dephosphorylation at later time-points in WT

platelets this dephosphorylation does not occur in c-Cbl deficient platelets.

Comparing this to convulxin-mediated Syk phosphorylation in TULA-dKO platelets

(Figure 16A) we see a similar pattern of impaired Syk dephosphorylation. Figure

31B shows a dose response of Syk phosphorylation in CRP treated WT and c-Cbl

deficient platelets and shows a hyperphosphorylation of Syk when c-Cbl is absent.

Comparing these data to the same experiment performed in TULA-dKO platelets we

see a similar pattern of Syk phosphorylation (Figure 17B). These data confirm the

data presented by Auger et al. and previously published data from this laboratory.

Furthermore, when comparing the Syk phosphorylation profiles from TULA-dKO

and c-Cbl deficient platelets, the similarities suggests there could be a relationship

between the mechanism of negative regulation of Syk phosphorylation.

PLC γ2 is hyperphosphorylated in c-Cbl deficient platelets

PLC γ2 is hyperphosphorylated in platelets deficient in TULA-2, most likely due to a lack of regulation of Syk causing enhanced downstream signaling. Therefore, concomitantly with Syk phosphorylation studies in c-Cbl deficient platelets we also performed studies of PLC γ2 phosphorylation to see if this phenomenon also occurred

in c-Cbl deficient platelets. Figure 32A shows a time-course of PLC γ2

phosphorylation following convulxin stimulation and shows a potentiation of PLCγ2

phosphorylation in c-Cbl deficient platelets. A similar phenomenon was observed in

a CRP dose-response experiment (Figure 32B). A similar profile of PLC γ2

97 phosphorylation is seen in TULA-dKO platelets (Figures 19A and 20B) again suggesting there may be a link between c-Cbl and TULA-2 mediated negative regulation of Syk and GPVI signaling.

Figure 31. Syk is hyperphosphorylated in c-Cbl deficient platelets. (A) Isolated WT and c-Cbl deficient platelets were stimulated with 200ng/ml convulxin for 0, 30, 60 120 and 240 seconds, the protein precipitated and pY519/20 Syk and total Syk were probed. (B) As part A except platelets were stimulated with 0.5, 1, 2, 5 or 10 g/ml CRP for 2 minutes.

98

Figure 32. PLC γγγ2 is hyperphosphorylated in c-Cbl deficient platelets. (A) Isolated WT and c-Cbl deficient platelets were stimulated with 200ng/ml convulxin for 0, 30, 60, 120 and 240 seconds, the protein precipitated, Western blotted and pY759 PLC γ2 and total PLC γ2 probed for. (B) As part A except platelets were stimulated with 0.5, 1, 2, 5 or 10 g/ml CRP for 2 minutes.

99 Ca 2+ mobilization is not potentiated in c-Cbl deficient platelets

To more fully evaluate the GPVI signaling cascade in c-Cbl deficient platelets and to

see if the aberrant Syk and PLC γ2 phosphorylation caused a potentiation of Ca 2+ mobilization, a dose-response of Ca 2+ mobilization in WT and c-Cbl (-/-) was performed. FURA-2-AM loaded WT and c-Cbl (-/-) platelets were stimulated with

CRP and the fluorescence due to Ca 2+ mobilization recorded. Figure 33 shows a plot of this Ca 2+ mobilization from 3 separate experiments fitted with a non-linear regression and, unlike TULA-dKO platelets, shows no difference between WT and c-

Cbl (-/-). These data suggest there may be differences between the TULA-2 and c-Cbl mediated negative regulation of GPVI signaling.

GPVI-mediated platelet aggregation is enhanced in c-Cbl deficient platelets

In addition to biochemical studies of GPVI signaling in c-Cbl deficient platelets, functional studies were performed to see if they exhibited the same phenotype as

TULA-dKO and TULA-2(-/-) platelets. Figure 34 shows aggregation traces from convulxin stimulated WT and c-Cbl deficient platelets and indicates that c-Cbl deficient platelets are hyperactive in response to convulxin compared to WT platelets, an effect that is mirrored in the TULA-dKO and TULA-2(-/-) platelets.

100

Figure 33. Ca 2+ mobilization is not potentiated in c-Cbl deficient platelets. FURA-2-AM loaded murine platelets were stimulated with varying doses of CRP in the presence of 100 M 2MeSAMP, 10 M MRS2719, 150nM Echistatin, 10 M Indomethacin and 2mM Probenecid and the Ca 2+ mobilization measured using a spectrofluorometer. The change in intracellular calcium was then measured, plotted against dose of CRP and fitted with a hyperbolic curve. Data points are constructed from 3 independent experiments. Closed circles are WT values and open circles are c- Cbl deficient values.

101

Figure 34. GPVI-mediated platelet aggregation is enhanced in c-Cbl deficient platelets. Isolated WT and c-Cbl deficient platelets were stimulated with 100ng/ml convulxin and the change in light transmittance observed.

102 TULA-2 does not require c-Cbl to bind Syk

Although Syk and c-Cbl are present in TULA-2 pulldowns and immunoprecipitates

this does not necessarily mean that they form a trimolecular complex. Syk may be

binding to one TULA-2 molecule and c-Cbl a different TULA-2 molecule. Also, the

similarities between TULA-dKO and c-Cbl deficient Syk and PLC γ2 phosphorylation profiles is not proof of an interaction between Syk, c-Cbl and TULA-2. Therefore, an in vitro binding experiment was performed using agarose-conjugated TULA-2 and

TULA-2 H380A, Syk and c-Cbl to ascertain whether c-Cbl was required for Syk to bind TULA-2. Figure 35A shows a Western blot of the association assay and indicates that while Syk and c-Cbl can bind to TULA-2 and TULA-2 H380A simultaneously, c-Cbl is not required for Syk to bind to TULA-2. Quantitation of the simultaneous c-Cbl and Syk binding suggests that Syk and c-Cbl compete for binding to TULA-2 and TULA-2 H380A however not enough data was acquired to calculate whether this competition was significant (figure 35B and C). These data suggest that

TULA-2 does not have a role in c-Cbl mediated negative regulation of Syk phosphorylation and GPVI signaling and therefore c-Cbl may negatively regulate Syk and GPVI signaling in a TULA-2 independent manner. This is discussed more fully in the discussion section.

103

Figure 35. TULA-2 does not require c-Cbl to bind to Syk. (A) Agarose-conjugated GST, GST-TULA-2 and GST-TULA-2 H380A were incubated for 15 minutes with Syk, c-Cbl or both together, Western blotted and probed for TULA-2, Syk and c-Cbl. (B) Mean band intensity of Syk for 2 experiments was plotted as a histogram for Syk alone binding to TULA-2 or Syk binding to TULA-2 in the presence of c-Cbl and a t-test performed. (C) As part B except binding to TULA-2 H380A quantitated.

104 CHAPTER 4

DISCUSSION

The platelet GPVI receptor is important for the initiation of platelet activation and hemostasis following damage to the vascular endothelium (Watson et al. 2005). The signaling cascade is a complex one, which involves numerous kinases and adapter proteins that undergo phosphorylation to converge on PLC γ2 to cause Ca 2+ mobilization and platelet activation (Watson et al. 2005). To date, the identity of a substantial number of these signaling proteins has been realized. One such protein is the tyrosine kinase Syk, whose kinase activity and activation are indispensable for collagen-mediated platelet activation (Poole et al. 1997). However, substantially less is known about the identity of the proteins that negatively regulate GPVI signaling and the mechanisms by which this negative regulation occurs. This thesis has focused on defining, in part, the mechanisms leading to negative regulation of Syk and more specifically, the identity and regulation of the protein that dephosphorylates

Syk. The study of the negative regulation of GPVI signaling is of critical importance as negative regulation of platelet activation serves to prevent unnecessary platelet activation and potentially harmful thrombus formation. Based on previously published data, we hypothesized that the phosphatases SHP-1, SHP-2 and TULA-2 may have a role in negatively regulating Syk by dephosphorylation, thus negatively regulating GPVI signaling and platelet activation. To test this hypothesis, we posed four questions: Is Syk a substrate for a specific phosphatase expressed in platelets?

Do Syk and a specific phosphatase associate in GPVI stimulated platelets? Do

105 phosphatase-null murine platelets display a phenotype in which Syk

dephosphorylation is impaired? Do phosphatases play a role in the c-Cbl-mediated

negative regulation of Syk?

Is Syk a Substrate For a Specific Phosphatase Expressed In Platelets?

In order to evaluate whether Syk was a substrate for various phosphatases, in vitro phosphatase assays were performed. Candidate phosphatases were incubated with phosphorylated Syk and its dephosphorylation measured using phosphospecific antibodies in a Western blot.

SHP-1

SHP-1’s expression in platelets is well documented. Furthermore, its reported association with Syk in platelets suggest Syk could be a substrate for SHP-1 in platelets (Pasquet et al. 2000). Additionally, work by Plas et al. showed that the Syk homologue, Zap70, could be dephosphorylated by SHP-1. Therefore, we performed an in vitro phosphatase assay looking for a dephosphorylation of Syk by SHP-1.

Figure 2 indicates that, at least in vitro , Syk appears to be a substrate for SHP-1.

Therefore, these data are in agreement with the indirect evidence presented by

Pasquet et al. regarding the association of SHP-1 and Syk in platelets, as well as T-

cell data presented by Plas et al.

106 SHP-2

A negative regulatory role for SHP-2 in GPVI-dependent platelet function has been postulated by Patil et al. In addition, some reports have implicated SHP-2 in the negative regulation of the, GPVI-like, TCR (Kwon et al. 2005). Also, SHP-2 has been shown to associate with Syk (Xu and Pecht 2001). Therefore, its ability to dephosphorylate Syk was evaluated. As shown in figure 8, SHP-2 did not dephosphorylate Syk as rapidly as SHP-1 suggesting that SHP-2 may not be the phosphatase responsible for dephosphorylating Syk in platelets. However, dephosphorylation of a potential substrate in vitro does not necessarily correlate with

the natural substrates for a phosphatase within the cell. Spatial regulatory

mechanisms such as recruitment of SHP-2 via its SH2 domains to tyrosine

phosphorylated residues can cause the localization of the phosphatase to a particular

compartment of cell. If a specific substrate, in this case Syk, is concentrated in this

compartment then apparent lack of dephosphorylation of an in vitro substrate may be

overcome (Tiganis and Bennett 2007). Indeed, recruitment of SHP-2 via its dual SH2

domains to an immunoreceptor tyrosine inhibition motif (ITIM) found in PECAM-1

is thought to be the mechanism by which PECAM-1 negatively regulates GPVI-

mediated platelet activation (Jackson et al. 1997; Patil, Newman, and Newman 2001).

Therefore SHP-2 may still have a role in regulating Syk phosphorylation that is

masked in a simplistic in vitro assay. Thus, we further investigated the possible

interaction between SHP-2 and Syk as discussed below.

107 TULA-2

The expression of SHP-1 and SHP-2 in platelets is well documented. However, the presence of TULA family proteins in platelets had not been described. Performing a

Western blot of total platelet protein, we show that TULA-2 is expressed in human and murine platelets with little or no TULA being present (figure 11). TULA-2 is expressed in a large number of tissues and thus it is not surprising that it is present in platelets (Tsygankov 2008). The data for TULA are consistent with previous reports indicating that TULA expression is limited to lymphoid cells (Tsygankov 2008).

While the lowest level of detection for Western blotting is in the picogram range there still may be a femtogram amount of TULA expressed in platelets that we could not detect. A more sensitive method of protein detection such as mass spectrometry could be utilized to address this question. We did not directly address this question but instead determined the biochemical and functional responses of platelets deficient in each TULA family member, singly, as discussed further below.

Having established that TULA-2 is expressed in platelets, we utilized an in vitro phosphatase assay to evaluate the ability of TULA-2 to dephosphorylate Syk (Figure

13). We show that full-length TULA-2 can dephosphorylate Syk. This is consistent with previous findings which show Syk dephosphorylation by the phosphatase domain of TULA-2 in vitro as well as similar experiments using Zap70 as a substrate

(Agrawal, Carpino, and Tsygankov 2008; Mikhailik et al. 2007).

108 Do Syk And a Specific Phosphatase Associate In GPVI activated Platelets?

The second approach was to determine whether a specific association of Syk with

each of the three phosphatases can occur. Association of Syk with a phosphatase

suggests that the phosphatase may be able to dephosphorylate Syk.

SHP-1

Association of Syk with SHP-1 has previously been reported utilizing multiple methodologies (Pasquet et al. 2000). Our coimmunoprecipitation studies revealed that SHP-1 does not associate with Syk suggesting that in the intact cell, Syk may not be a substrate for SHP-1 (figure 3). This is in obvious disagreement with previous reports. A possible explanation for this is the different time-point used by us and

Pasquet et al. We performed the immunoprecipitation at 15 seconds whereas Pasquet et al. performed the immunoprecipitation at 2 minutes. It may be that Syk only associates with SHP-1 at later time-points. Hence, we would not Syk and SHP-1 association at 15 seconds.

SHP-2

Although SHP-2 did not dephosphorylate Syk as rapidly as SHP-1, its possible association with Syk in platelets was still evaluated since SHP-2 can associate with

Syk in RBL cells, is a documented negative regulator of platelet activation and, in

109 some reports, a negative regulator of the TCR (Kwon et al. 2005; Xu and Pecht

2001). The data shown in figure 9 indicates that, in platelets, Syk does not coimmunoprecipitate with SHP-2. This datum taken together with the in vitro phosphatase assay (figure 8) suggests that SHP-2 is probably not the phosphatase responsible for dephosphorylating Syk in the platelet. Not observing an association of Syk with SHP-2 in platelets does not agree with the data from RBL -2H3 cells (Xu and Pecht 2001). The association between Syk and SHP-2 may therefore be a mast cell specific phenomenon.

SHP-2’s binding to the ITIM domain containing inhibitory receptor, PECAM-1, and

PECAM-1’s implication as a negative regulator of GPVI-signaling suggest that SHP-

2 does have an important role in negatively regulating GPVI signaling (Jackson et al.

1997; Patil, Newman, and Newman 2001). As our data suggests Syk is not a SHP-2 substrate then SHP-2 must have other substrates within the GPVI signaling cascade.

ITIM containing receptors have been put forward as possible negative regulators of

ITAM receptor signaling (Neel 1997). Binding of phosphatases to the ITIM allows it to then dephosphorylate the ITAM or tyrosine kinases involved in signal transduction(Neel 1997). Thus, SHP-2 may dephosphorylate the FcR γ chain ITAM or maybe Fyn and Lyn to negatively regulate the GPVI signaling cascade. Having a lack of strong evidence that SHP-2 may be negatively regulating Syk in platelets and studying other possible substrates being beyond the scope of this work, SHP-2 was not pursued further.

110 TULA-2

Having established that TULA-2 is expressed in platelets and the full-length protein can dephosphorylate Syk, its ability to form a complex with Syk and c-Cbl was measured. Figure 14 and 15 reflect the two approaches used and show that in both

GST-pulldowns and coimmunoprecipitations TULA-2 can bind to Syk and c-Cbl.

The binding of TULA-2 to Syk is consistent with the work of Agrawal et al. who showed TULA-2 and Syk could interact in HEK293T cells expressing both proteins.

Additionally, the GST-pulldown studies suggest a stimulation dependent association of Syk with TULA-2. A logical explanation for this is that Syk must undergo phosphorylation and activation, as is seen in GPVI stimulation, before TULA-2 binding and dephosphorylation to regulate its kinase activity. Alternatively, recruitment of Syk to the phosphorylated ITAM may bring it in closer proximity to

TULA-2 to allow a greater extent of association between TULA-2 and Syk. Little is known about TULA-2 subcellular localization other than it is found in the cytosol and also membrane bound, but if TULA-2 is enriched in the areas surrounding the

GPVI/FcR γ then an increased level of TULA-2 and Syk association on stimulation is plausible (Tsygankov 2008). Phosphorylation data from the coimmunoprecipitation studies indicates that at least a portion of the Syk associating with TULA-2 in unstimulated platelets is phosphorylated at Y525/6. It is not possible to say whether all the associating Syk is phosphorylated but if it was then this would fit with the stimulation dependent increase in association between Syk and TULA-2, as, in this instance, Syk phosphorylation is required for TULA-2 binding. If not all the Syk bound to TULA-2 is phosphorylated then it is possible that TULA-2 can bind both 111 phosphorylated and non-phosphorylated Syk but preferentially binds phosphorylated

Syk in order to dephosphorylate it. Clearly a more detailed investigation of the interaction of Syk and TULA-2 and the factors affecting this association is required.

Analysis of the Syk band in the immunoprecipitates by mass spectrometry looking for phosphorylated and non-phosphorylated species of Syk would be one approach that could be used to ascertain whether TULA-2 binds phosphorylated Syk exclusively or can also bind non-phosphorylated Syk.

Furthermore, it is interesting to note that a fraction of Syk appears to be phosphorylated in unstimulated platelets, as this is not clearly observed in studies of lysates or protein precipitates from unstimulated platelets (figure 16 and 17). It is possible that the coimmunoprecipitation studies enrich for the phosphorylated form of

Syk so it can more easily be seen via Western blotting if TULA-2 binds only phosphorylated Syk. This datum also suggests that TULA-2 may constitutively bind

Syk to dephosphorylate it to prevent platelet activation in the absence of collagen.

Thus a tonic basal activity of TULA-2 may be required in the platelet to help keep it in its quiescent discoid state. To date, little is known about the mechanisms regulating TULA-2 activity but studying the catalytic activity in unstimulated and across a time-course of GPVI-stimulated platelets would provide new insight into

TULA-2’s function in general terms and in platelets.

112 The in vitro phosphatase data, taken together with the GST-pulldown and

coimmunoprecipitation data, suggest that TULA-2 could be the phosphatase that

negatively regulates Syk during GPVI-mediated platelet activation. The implications

of c-Cbl binding to TULA-2 are discussed further below.

Do Phosphatase-null Murine Platelets Display a Phenotype In Which Syk

Dephosphorylation Is Impaired?

The third approach used to investigate the possible role of SHP-1 and TULA-2 in the

negative regulation of Syk activity in GPVI-mediated platelet activation was to use

murine platelets deficient in each phosphatase. GPVI signaling and GPVI-mediated

platelet function were studied in these platelets as a means to evaluate the possible

role of the candidate phosphatases in regulating Syk in GPVI signaling. SHP-2 null

murine platelets were not evaluated due to the lack of strong evidence for its

involvement in Syk regulation and SHP-2 deficiency being embryonic lethal.

SHP-1

To study the effect of SHP-1 deficiency on Syk and GPVI signaling the spontaneously mutated SHP-1 strain of mice, mev, was used. Pasquet et al. have successfully utilized this strain to characterize SHP-1 deficiency in GPVI signaling.

In terms of biochemical analysis, we have shown that following stimulation of GPVI,

113 Syk is hypophosphorylated compared to WT platelets (figure 6). This is in agreement

with Pasquet et al. who showed the same Syk phosphorylation phenotype. This result

does not fit with SHP-1 being a negative regulator of Syk phosphorylation in GPVI

signaling however, as it can be reasoned that deletion of a phosphatase regulating Syk

phosphorylation would lead to an increase in Syk phosphorylation when the

phosphatase is absent. The opposite is observed here. The reason for the

hypophosphorylation of Syk in mev mice is unclear. It is possible that SHP-1 may

dephosphorylate the inhibitory phosphorylation site on src-family kinases thus

positively regulating Fyn and/or Lyn in the GPVI signaling cascade. Thus, when

SHP-1 is absent Fyn and/or Lyn are retained in their inactive form leading to

decreased GPVI signaling and Syk phosphorylation. Senis et al. have proposed that

the receptor protein tyrosine phosphatase, CD148, may be performing this function in

platelets. Therefore working in concert, SHP-1 and CD148 may regulate src-family

kinase activation (Senis et al. 2009). This possible positive regulatory role in GPVI

signaling for SHP-1 is also evident in the platelet functional data presented by

Pasquet et al. and in some of the functional data presented here. Pasquet et al

reported a suppression of α-granule secretion in mev platelets and we show a slight suppression in dense granule secretion (figure 5). We did not, however, observe a suppression of aggregation in mev platelets but it is possible that the dose of convulxin was too high to discern differences in aggregation between mev and WT platelets (figure 4). It therefore appears that SHP-1 is a positive regulator of GPVI signaling and does not appear to be a negative regulator of Syk phosphorylation in

114 platelets despite it being able to dephosphorylate SHP-1 in vitro. The mechanism by which SHP-1 positively regulates GPVI signaling remains to be investigated.

TULA-2

To study the effect of TULA-2 on GPVI signaling and platelet function we primarily

used TULA-dKO murine platelets. These results were confirmed with studies in

murine platelets deficient in either TULA family member.

Consistent with the proposed phosphatase function of TULA-2, we observed a

hyperphosphorylation of Syk in a time and dose-dependent manner with convulxin,

CRP and collagen in TULA-dKO platelets (figure 16 and 17). This also caused an

increase in Syk kinase activity (figure 18). Hyperphosphorylation was also observed

in TULA-2(-/-) platelets, thus confirming that it is indeed TULA-2 that is responsible for the phenotype seen in TULA-dKO platelets (figure 27 and 28). The single knockout data also confirms that TULA does not appear to have a role in regulating

Syk phosphorylation in platelets. This is not surprising as we did not detect any measurable amount of TULA in platelets. While these data are consistent with a negative regulatory role of TULA-2, in T-cells hyperphosphorylation of the Syk homologue ZAP-70 is only observed in cells deficient in both TULA family members and not TULA-2(-/-) cells (Carpino et al. 2004; Mikhailik et al. 2007). This suggests that the negative regulation of Zap-70 following T-cell receptor engagement relies on

115 both TULA and TULA-2. Thus, the TULA-2-mediated negative regulation of signaling differs between platelets and T-cells.

From the lack of Syk dephosphorylation in TULA-2 deficient platelets we can also conclude that SHP-1 and SHP-2 have little or no role in dephosphorylating Syk. If either phosphatase was working in combination with TULA-2 we would expect less hyperphosphorylation of Syk in the TULA-2 deficient platelets, as residual dephosphorylation would be occurring due to SHP-1 or SHP-2. Instead, we see a substantial hyperphosphorylation of Syk. To conclusively prove SHP-1 or SHP-2 has little or no role, mice lacking TULA-2 along with SHP-1 or SHP-2 could be generated. The Syk phosphorylation phenotype between these mice and TULA-2(-/-) mice could then be compared to ascertain whether SHP-1 or SHP-2 have minor roles in dephosphorylation.

While the enhanced Syk phosphorylation observed here happens at all time-points the most marked differences are present at later time-points. However, Ca 2+ mobilization and platelet activation occur rapidly, most notably with convulxin. Thus, it is unclear whether the phosphorylation of Syk at later time-points and the subsequent potentiation seen in TULA-2 deficient platelets is of physiological importance or just residual phosphorylation that remains following Ca 2+ mobilization. Tomlinson et al. have shown that weak but sustained GPVI signaling by collagen is required to maintain spreading of the platelet on collagen over a time-course of several hours.

116 Therefore continual phosphorylation of Syk is probably an important factor to help

maintain the spreading of the platelet on collagen (Tomlinson et al. 2007).

Furthermore, in terms of maintenance of a thrombus in vivo, the constant GPVI

signaling and thus Syk phosphorylation may play an important role. While convulxin

is a powerful experimental tool that induces robust tyrosine phosphorylation and

platelet activation, the time-course of collagen-mediated platelet activation is slower

than that of convulxin. Therefore, the spike in Ca 2+ mobilization and Syk

phosphorylation at earlier time-points observed with convulxin may not fully reflect

the temporal kinetics of collagen-mediated Ca 2+ mobilization in vivo.

In addition to enhanced Syk phosphorylation in murine platelets deficient in TULA-2,

we observed enhanced PLC γ2 phosphorylation in a time and dose-dependent manner

(figure 19 and 20). These data are significant as they show that not only does TULA-

2 regulate Syk phosphorylation but this regulation is also important in controlling the

formation of a signalosome as a platform for the activation of PLC γ2, as enhanced

Syk phosphorylation causes enhanced PLC γ2 phosphorylation, not just an increase in

the fraction of phosphorylated Syk. Syk is unable to phosphorylate PLC γ2 directly, at

least in vitro , therefore it is the enhanced signaling of the whole GPVI signaling cascade when TULA-2 is absent, due to a lack of regulation of Syk, that causes the enhanced PLC γ2 phosphorylation (Rodriguez et al. 2001).

117 Given the observed hyperphosphorylation of PLC γ2 it is possible to postulate that

PLC γ2 could be a substrate for TULA-2, thus a lack of TULA-2 leads to PLC γ2

hyperphosphorylation and enhanced platelet function. This possibility remains to be

investigated. However based on the dephosphorylation of Syk by TULA-2 and the

association between Syk and TULA-2 seen in figure 14 and, we propose that the

hyperphosphorylation of PLC γ2 observed is due to lack of Syk regulation. The

previously published data suggesting Syk and Zap70 are TULA-2 substrates also

support this proposition (Agrawal, Carpino, and Tsygankov 2008; Mikhailik et al.

2007).

Calcium mobilization is a critical event in the activation of platelets. Thus, we tested

whether the lack of Syk regulation by TULA-2 in TULA-2 deficient platelets had a

concomitant effect on calcium mobilization. We show that there is a statistically

significant potentiation of Ca 2+ mobilization in CRP-treated TULA-dKO platelets compared to WT (figure 21). This potentiation is consistent with the negative regulatory function of TULA-2 in GPVI signaling and correlates with the hyperphosphorylation of Syk and PLC γ2 presented here. This result is also consistent

with previous published work in which enhanced PLC γ2 activity leads to potentiated

Ca 2+ mobilization (Elvers et al. 2010). An interesting difference between the PLC γ2

phoshorylation and the Ca 2+ mobilization is the dose dependence. At low doses or

CRP only a small difference in PLC γ2 is seen but the largest difference in Ca2+

mobilization is seen and at higher doses the reverse is observed. A possible

118 explanation for this that at higher doses of CRP a maximal amont of PLC γ2 has been activated to mobilize Ca 2+ and the additional phosphorylation is now an excess and has no functional significance. Alternatively, the PLC γ2 substrate, PIP2, may have become exhausted and so is rate limiting at higher agonist concentrations.

To ensure that deletion of TULA-2 caused a platelet functional phenotype and not just a biochemical phenotype, platelet functional responses were measured in TULA- dKO and TULA-2(-/-) platelets . Ex vivo , we show that platelet aggregation and dense granule secretion are potentiated in platelets deficient in TULA-2 (figure 22, 23, 29 and 30). We also confirm that TULA plays no role in regulating GPVI platelet functional responses as no phenotype was observed in platelets deficient in TULA

(figure 29 and 30). The negative regulatory function of TULA-2 is consistent with data published in T-cells following T-cell receptor engagement where TULA-dKO T- cells exhibit a hyper-responsive phenotype to T-cell receptor engagement. But, as discussed above, deletion of both TULA family members was required to observe a phenotype in T-cells whereas in platelets it appears that only deletion of TULA-2 is required, possible because only TULA-2 is expressed in platelets. The negative regulatory function of TULA-2 we show here is also in agreement with Raguz et al. who showed the EGFR is negatively regulated by TULA-2 dephosphorylation of phosphorylated tyrosines (Raguz et al. 2007).

119 Our functional data also suggests that TULA-2 specifically regulates GPVI signaling,

or at least is specific to signaling cascades where Syk has a predominant role, as no

difference between platelet function was observed between WT and TULA-dKO

platelets stimulated with the PAR-4 agonist, AYPGKF (figure 22 and 23).

Other signaling cascades that TULA-2 may regulate in platelets

Two other receptors downstream of which TULA-2 may play a role are C-type lectin-

like type II (CLEC-2) and the fibrinogen receptor, αIIb β3, as Syk is an important kinase in the signaling cascades downstream of these receptors (Gao et al. 1997;

Spalton et al. 2009; Woodside et al. 2001; Woodside et al. 2002). If TULA-2 is specifically regulating Syk downstream of GPVI then we would not expect to see a phenotype downstream of these receptors in TULA-2 deficient platelets. In CLEC-2 signaling, Syk is recruited to a single phosphorylated tyrosine on the cytoplasmic tail of CLEC-2 to initiate signaling and leads to platelet activation via PLC γ2 (Spalton et al. 2009). Thus, stimulating CLEC-2 in WT and TULA-2 deficient platelets and performing similar experiments to those outlined here may reveal whether TULA-2 is also involved in CLEC-2 signaling. For αIIb β3, Syk is recruited to the β3 cytoplasmic tail during fibrinogen binding (Woodside et al. 2001; Woodside et al. 2002).

Therefore, studying Syk phosphorylation in WT and TULA-2 deficient platelets spread on fibrinogen may indicate whether TULA-2 is involved in regulating Syk phosphorylation downstream of αIIb β3.

120 In addition to ex vivo platelet functional studies, in vivo thrombosis injury model studies were performed using FeCl 3 to injure the vascular endothelium and expose subendothelial collagen. Our data shows that there is an enhanced time to occlusion in TULA-dKO mice and the thrombi formed by 7.5% FeCl 3 injury are more stable in

TULA-dKO mice when compared to WT (figure 26). This is consistent with the hypothesis that TULA-2 is a negative regulator of GPVI signaling and correlates with the enhanced phosphorylation of GPVI signaling proteins, Ca 2+ mobilization and

enhanced GPVI-mediated platelet functional responses shown here. These data are

also consistent with in vivo thrombosis model data published for other reported

negative regulators of platelet function. Falati et al. reported that PECAM-1 deficient

mice are prothrombotic, indicated by a shorter time to occlusion of the carotid artery

in the FeCl 3 thrombosis model (Falati et al. 2006). Additionally, carcinoembryonic

antigen cell adhesion molecule-1 (CEACAM-1) deficient mice also exhibit a

prothrombotic phenotype, when mesenteric arterioles are subject to FeCl 3 injury

(Wong et al. 2009).

The data from murine platelets deficient in SHP-1 and TULA-2 presented here

suggest that TULA-2, and not SHP-1, is the phosphatase that negatively regulates Syk

in GPVI-mediated platelet activation.

121 Do Phosphatases Play a Role in the c-Cbl-mediated Negative Regulation of Syk?

Deletion of c-Cbl has been shown to potentiate platelet functional responses as well

as Syk phosphorylation in GPVI-stimulated platelets (Auger et al. 2003; Dangelmaier

et al. 2005). The mechanism for the c-Cbl–mediated negative regulation remains

unknown. Based on the Syk hyperphosphorylation observed in c-Cbl (-/-) platelets and the reports that Syk and a number of phosphatases can bind to c-Cbl, we hypothesized that the adapter function of c-Cbl may be responsible. This was therefore investigated.

SHP-1

SHP-1 has been shown, indirectly via Tyk-2, to bind to c-Cbl therefore we looked for an association between SHP-1 and c-Cbl in platelets (Uddin et al. 1996; Yetter et al.

1995). We show that SHP-1 does not coimmunoprecipitate with c-Cbl (figure 7), suggesting that SHP-1 may not be the phosphatase responsible for mediating the phenotype in c-Cbl deficient platelets. This result is in disagreement with previously published reports in hematopoietic cell lines. Tyk-2 has been shown to be expressed in platelets, so all the components required for the interaction between SHP-1 and c-

Cbl are present in platelets (Rodriguez-Linares and Watson 1994). We therefore conclude that this interaction may not occur in platelets.

122 SHP-2

Unlike SHP-1, SHP-2’s interaction with c-Cbl has been shown to be direct

(Chernock, Cherla, and Ganju 2001). To investigate its possible association with c-

Cbl in platelets we used a similar approach to that of SHP-1, namely coimmunoprecipitation. Using this approach we did not see a specific interaction between SHP-2 and c-Cbl (figure 10). This result is in disagreement with Chernock et al. thus the interaction between SHP-2 and c-Cbl may be confined to Jurkat T-cells or is an interaction that does not occur in platelets.

TULA-2

As discussed above, we show that TULA-2 can bind to Syk. In addition to binding

Syk, we have shown that TULA-2 can also bind c-Cbl. These data are in agreement

with Kowanetz et al. who observed an association between TULA-2 and c-Cbl in

HEK293T cells and Agrawal et al, who showed an association between Syk and

TULA-2 in HEK293T cells (as discussed above). These data therefore suggest that

TULA-2 could be the phosphatase responsible for the Syk hyperphosphorylation

phenotype observed in c-Cbl deficient platelets.

To more fully investigate the possible role of TULA-2 in mediating c-Cbl negative

regulation of GPVI-signaling, a more detailed study of c-Cbl (-/-) platelets was

performed. We confirm the reports of Auger et al. and Dangelmaier et al. who

observed a potentiation of Syk phosphorylation and platelet functional responses in c- 123 Cbl deficient platelets (figure 31 and 34). Additionally, we report a potentiation of

PLC γ2 phosphorylation in c-Cbl deficient platelets, which is previously unreported

(figure 32). Comparing these data with data from the TULA-dKO and TULA-2(-/-) a similar pattern of Syk phosphorylation in the mutant platelets is evident.

Furthermore, the platelet aggregation phenotype is the same in the mutant platelets

(figure 22 and 34). Taken together with the GST-pulldown and coimmunoprecipitation data, these comparisons provide corollary evidence that

TULA-2 may be responsible for c-Cbl negative regulation of GPVI signaling.

One aspect of the GPVI signaling cascade that differed between the TULA-dKO and c-Cbl (-/-) platelets was Ca 2+ mobilization. TULA-dKO platelets exhibited a potentiation of Ca 2+ mobilization in response to CRP but c-Cbl (-/-) platelets showed no

difference compared to WT platelets (figure 33). This was somewhat unexpected,

however c-Cbl can bind a multitude of signaling proteins some of which may

positively regulate Ca 2+ mobilization in the GPV signaling cascade (Tsygankov et al.

2001). Therefore, the potential positive and negative effects on c-Cbl negate each other when c-Cbl is absent, so no Ca 2+ mobilization phenotype is seen. Alternatively, the differences observed are possible evidence that c-Cbl- and TULA-2-mediated negative regulation of GPVI signaling are not related.

Although Syk and c-Cbl appeared in TULA-2 pulldowns and coimmunoprecipitations, a trimolecular complex is not necessarily forming between

124 the three proteins. It may be that Syk is binding to one TULA-2 molecule and c-Cbl

another. Therefore, an in vitro binding assay was performed to see if Syk could bind to TULA-2 in the absence of c-Cbl (figure 35A). We found that Syk could bind to

TULA-2 in the absence of c-Cbl suggesting that TULA-2 may not be mediating the c-

Cbl negative regulation of GPVI signaling. While these data suggest that Syk can bind to TULA-2 in the absence of c-Cbl a more detailed study of the interaction between the three proteins may be of value. This would conclusively show that c-Cbl is not required, as the in vitro conditions may not fully reflect the associations occurring inside the cell. To achieve this, a number of approaches could be used.

First, various point mutations in the three proteins could be made at the reported binding sites between Syk, c-Cbl and TULA-2. For Syk, a mutation at Y323 to phenylalanine which would prevent its binding to c-Cbl. A tryptophan to leucine mutation in the SH3 of TULA-2 to render its SH3 domain unable to bind proline rich domains thus should not be able to bind to TULA-2. For c-Cbl, a Glycine (G) 306

Glutamic acid (E) would render it unable to bind Syk. Cotransfecting various combinations of these proteins with WT proteins and performing coimmunoprecipitations may provide further evidence regarding the nature of the interaction between TULA-2 and Syk and the domains that are required. Depending on the results from the first approach, a second approach would be to produce transgenic mouse models containing the same mutated proteins. Studying GPVI- mediated platelet function and signaling and association between the three proteins may provide more insight into the Syk-TULA-2 interaction in a more physiological setting.

125

In addition to observing TULA-2 binding to Syk in a c-Cbl independent manner, we observed that c-Cbl and Syk appeared to compete for binding with TULA-2, although this result was not significant (figure 35B and C). This suggests that Syk and c-Cbl may bind at the same site on TULA-2. While Syk can bind to c-Cbl in its tyrosine kinase binding domain it has also been reported that Syk can bind to c-Cbl via c-Cbl’s proline rich domain (Ota et al. 2000). As TULA-2 binds to c-Cbl’s proline rich domain it is possible that the two proteins may compete for c-Cbl binding in the platelet. The experiments outlined above regarding studying the three proteins interactions with mutated proteins would also provide evidence for Syk and TULA-2 competitive binding with c-Cbl.

In summary, the GST-pulldown and coimmunoprecipitation data suggest TULA-2 could be mediating the c-Cbl negative regulation of Syk, by binding to c-Cbl and c-

Cbl to Syk. In addition, the similarities between the phenotypes of the TULA-2 and c-Cbl deficient platelets suggest the same. However, binding studies suggest c-Cbl is not required for TULA-2 to bind Syk. Therefore, the mechanism of c-Cbl negative regulation of GPVI remains refractory and more detailed studies are required as outlined in Future Experiments.

126 Summary and Conclusions

Collagen dependent activation of the platelet via GPVI is important for hemostasis following damage to the vascular endothelium. Therefore, negative regulation of

GPVI is of critical importance to prevent unnecessary platelet activation and potentially harmful thrombus formation. This work set out to identify the phosphatase responsible for the negative regulation of Syk during platelet GPVI signaling and whether these phosphatases mediated the c-Cbl negative regulation of

GPVI signaling. This was achieved by studying three phosphatases which could potentially be responsible for the negative regulation of Syk. These were SHP-1,

SHP-2 and TULA-2. We show that SHP-1 nor SHP-2 are the phosphatases responsible for the negative regulation of Syk. However, our studies indicate that

TULA-2 is the phosphatase responsible for the negative regulation of Syk in GPVI signaling by virtue of its ability to bind and dephosphorylate Syk to negatively regulate GPVI functional responses ex vivo and in vivo . In addition, we show that

TULA-2 may not mediate c-Cbl negative regulation of GPVI signaling, despite the similarities between phenotypes of TULA-2 deficient and c-Cbl deficient platelets.

Further investigation is required into the possible role of TULA-2 in c-Cbl negative regulation of GPVI signaling.

127 Future Experiments

1. Our data regarding the role of TULA-2 in mediating c-Cbl negative regulation

of GPVI signaling is equivocal. Binding studies suggest that c-Cbl is not

required for TULA-2 to bind Syk. To more fully investigate the relationship

between TULA-2 and c-Cbl mediated negative regulation of Syk we propose a

series of coimmunoprecipitation studies that include WT and mutant forms of

Syk, c-Cbl and TULA-2 cotransfected in HEK293T. Syk Y323F mutant and c-

Cbl G306E would render Syk and c-Cbl unable bind each other. TULA-2 W/L

mutant would inactivate the SH3 domain and render it unable to bind c-Cbl and

the truncated form of c-Cbl that comprises only the tyrosine kinase binding

domain of c-Cbl would render c-Cbl unable to bind TULA-2. Thus,

cotransfecting these mutants with WT proteins would provide more insight into

the possible role of TULA-2 in c-Cbl negative regulation of GPVI signaling.

Additionally, these experiments may provide more physiological data than the

in vitro binding assays we have performed in this study.

2. In the same cotransfected cells, localization studies could also be performed

using fluorescently labeled antibodies to Syk, c-Cbl and TULA-2. These

experiments would provide spatial data for Syk, c-Cbl and TULA-2. Overlay of

the three proteins would suggest the proteins can form a trimolecular complex.

However, the experiments may reveal a localization of perhaps two proteins to a

specific compartment of the cell.

128 3. To complement the experiments in HEK293T cells, transgenic mice could be

generated of the various Syk, c-Cbl and TULA-2 mutants. The mutation’s

effects on Syk phosphorylation, platelet functional responses and association

between the three proteins could then be evaluated.

4. In addition to studying the adapter function of c-Cbl in the above experiments

the E3 ligase activity of c-Cbl could be studied. Dangelmaier et al. have shown

that c-Cbl ubiquitinates Syk following GPVI stimulation but the purpose of this

is unknown. Using E3 ligase inactive c-Cbl mutant, C381A, the effect on Syk

phosphorylation, platelet functional responses and association between the three

proteins could then be evaluated. This may reveal the function of Syk

ubiquitination and any impact it has on Syk negative regulation by TULA-2.

5. While the above experiements focus of the the role of TULA-2 in c-Cbl

negative regulation of Syk an number of experiments to further study TULA-2

could also be performed. As discussed, little it known about the regulation of

TULA-2. We therefore propose to perform an immunoprecipitation of TULA-2

from unstimulated and GPVI-stimulated platelets and then measure the

hydrolysis of pNPP at the multiple time-points. This would provide insight into

whether there is a change in the overall phosphatase activity of TULA-2 during

GPVI stimulation and suggest that there are regulatory mechanisms controlling

TULA-2 in the platelet thus providing a new avenue of study for TULA-2.

129 6. To study in more detail the effect of TULA-2 on GPVI-mediated platelet

function Phosphatidylserine (PS) exposure could be measured as this is a

marker for activation of platelets and is Ca 2+ dependent. Thus TULA-2

deficient platelets would likely have an increased amount of PS exposure on the

platelet surface following GPVI-stimulation. This would be measure using a

FITC-labelled anti-Annexin V antibody and performing flow cytometry.

7. To study the effect of TULA-2 deletion in an almost entirely collagen

dependent setting and minimize the effect of generated thrombin the collagen

flow model could be used. Here, collagen coated flow chambers would be

perfused with WT and TULA-2 deficient platelets under flow and the binding to

collagen measured. We would expect to see an enhanced binding to collagen in

the TULA-2 deficient platelets. These data would provide more evidence for

TULA-2’s role in negatively regulating GPVI signaling.

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