Regulation of Endothelial Barrier Integrity Via Store-Operated Ca2+ Entry

Regulation of Endothelial Barrier Integrity via Store-Operated Ca2+ entry

DHEERAJ SONI BS, North Dakota State University, 2009

DISSERTATION

Submitted as partial fulfillment of the requirements for the degree of Doctor of Philosophy in Cellular and Molecular Pharmacology in the Graduate College of the University of Illinois at Chicago, 2017

Chicago, Illinois

Defense Committee:

Dr. Chinnaswamy Tiruppathi, Advisor and Chair Dr. Viswanathan Natarajan Dr. Stephen M. Vogel Dr. Youyang Zhao Dr. Pradeep K Dudeja, Physiology

I dedicate this dissertation to my wonderful parents,

Mr. Vijay Kumar Soni and Mrs. Manju Soni,

and my lovely sister Ms. Neelam Soni, whose support and affection knew no bounds and helped me achieve this goal.

ACKNOWLEDGMENTS

I am very grateful to my advisor Dr. Chinnaswamy Tiruppathi, an outstanding mentor, for his guidance, patience and support. He kept me motivated throughout my graduate study to achieve nothing short of excellence. His ability to think critically about any given research problem has inspired and influenced me over the years and still continues to do so. It was a real privilege to work under his directions for the past five years; I have learnt much more than a graduate student could hope for.

Next, I want to thank my committee members for their guidance and support. Specially, I am indebted to Dr. Vogel and Dr. Zhao who met with me every semester and gave their valuable inputs and suggestions to help me progress. I also want to thank Dr. Zhao for teaching me how to generate cell- specific knockout mice and providing the Cre strains. I am also thankful to Dr. Malik for encouraging me since the early years of my PhD.

I cannot thank enough the office members of the Department of Pharmacology for going out of their way to support me and help me resolve issues. Most importantly, I want to thank several members of the Tiruppathi lab; past and present, for all their insightful suggestions and discussions.

Special thanks to Dr. Auditi DebRoy, who taught me calcium measurement and helped me generate preliminary data for my project. I am grateful to Mr. Yubin Wu for teaching me cell culture, western blotting and mice handling techniques. I want to thank Dr. Dong-Mei Wang and Dr. Sushil C. Regmi who did excellent work in the lab and helped me to advance my research projects significantly. Last but not least,

I am grateful to the current lab members Dr. Manish Mittal, Dr. Saroj Nepal, Dr. Dagmara Grzych and Dr.

Luiza Grzych for their support and friendship and for making the Tiruppathi lab a great place to work.

I want to convey my heartfelt thanks to my friends and family. A special thanks to my cousin Dilip, and sister-in-law Neetu. Thank you for always supporting me since the early days of my studies in US. I thank Ankit and Nikhil, for being wonderful brothers and creating a home environment here in Chicago. I want to thank my overseas friends, Priyanka, Tripti and Nishant, for always being available to provide support and fall-back cushion during the long patience-testing journey of acquiring a

PhD. Thank you for cheering me through bad days, pushing me through difficult times and most

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ACKNOWLEDGMENTS (continued) importantly, always bringing the brighter side to the fore-front. Finally, I thank my parents for being my greatest strength, biggest inspiration, and loudest cheerleader, all throughout my life! Their love, support, motivation and philosophies had the most impact on my life and are the reason for what I am today.

I hope I make you proud.

TABLE OF CONTENTS

CHAPTER PAGE 1 LITERATURE REVIEW ...... 1 1.1 Endothelial cells ...... 1 1.1.1 Endothelial structure and function ...... 1 1.1.2 Endothelial barrier ...... 2 1.1.3 Endothelial dysfunction and inflammation ...... 6 1.2 Sepsis ...... 6 1.2.1 Pathogenesis ...... 6 1.3 Thrombin ...... 7 1.3.1 Structure and function ...... 7 1.3.2 Coagulation ...... 8 1.4 Protease-activated receptors (PAR) ...... 9 1.5 Calcium Signaling ...... 10 1.5.1 Types & function ...... 10 1.5.2 Store-operated Ca2+ entry ...... 11 1.6 Store-operated Ca2+ channel [SOC] ...... 13 1.6.1 Stromal Interacting Molecular proteins (STIMs) ...... 13 1.7 SOCE-mediated disruption of AJs ...... 16 1.8 Termination of Store operated calcium entry and resealing of AJs...... 16 1.9 TAK1 ...... 18 1.10 Endothelial barrier repair and restoration ...... 21 2 OBJECTIVES ...... 22 2.1 To determine the role of STIM1-activated Ca2+ entry [SOCE] in mediating vascular leak through disassembly of AJs ...... 22 2.2 To determine the role of TAK1 activation secondary to SOCE in restoring lung vascular barrier integrity after injury, via termination of SOCE and stabilization of β-catenin ...... 22 3 MATERIALS AND METHODS ...... 23 3.1 Materials ...... 23 3.2 Animals...... 24 3.2.1 Generation of endothelial-cell restricted STIM1 knockout (STIM1∆EC) mice ...... 24 3.2.2 Generation of inducible endothelial-cell restricted TAK1 knockout (TAK1i∆EC) mice ...... 24 3.3 Primary endothelial cell culture ...... 24 3.4 Genotyping ...... 26 3.5 Cell Culture ...... 26 3.6 Immunoprecipitation ...... 26 3.7 Immunoblotting ...... 27 3.8 Cytosolic Ca2+ measurement ...... 27 3.9 Transendothelial Electrical Resistance Measurement ...... 28 3.10 Immunostaining ...... 28 3.11 Paraffin-Embedded Tissue section staining ...... 28 3.12 Subcellular fractionation ...... Error! Bookmark not defined. 3.13 siRNA transfection ...... 29 3.14 In vivo siRNA delivery in mouse lungs ...... 29 3.15 Mouse Lung Capillary Filtration Coefficient Measurement ...... 29 3.16 Assessment of Mouse lung Microvessel permeability In Vivo ...... 30 3.17 In vitro deletion of TAK1 in LECs ...... 30 3.18 Statistical Analysis ...... 30 4 RESULTS I...... 31

TABLE OF CONTENTS (continued)

CHAPTER PAGE

4.1 Hypothesis: STIM1-induced SOCE activates Pyk2 to mediate tyrosine phosphorylation of VE- PTP and thus cause vascular leak through disassembly of AJs ...... 31 4.1.1 SOCE is required for tyrosine phosphorylation of VE-PTP and disassembly of VE-cadherin junctions ...... 31 4.1.2 EC specific STIM1 deletion inhibits phosphorylation of VE-PTP and VE-cadherin and increases in lung vascular permeability ...... 34 4.1.3 SOCE-induced Pyk2 activation phosphorylates VE-PTP to induce dissociation of VE-PTP from VE-cadherin ...... 37 4.1.4 In vivo silencing of Pyk2 in mouse lung microvascular ECs prevents VE-PTP tyrosine phosphorylation, VE-cadherin phosphorylation, and increased endothelial permeability ...... 41 4.1.5 VE-PTP C-terminal phospho-Y1981 binds to and activates Src to increase vascular permeability ...... 45 5 DISCUSSION I ...... 52 6 RESULTS II ...... 56 6.1 Hypothesis 1: TAK1 activation secondary to STIM1-mediated SOCE induces STIM1 phosphorylation which in turn terminates SOCE and thereby dampens the vascular permeability response ...... 56 6.1.1 Pharmacological inhibition of TAK1 augments Ca2+ entry as well as endothelial permeability ...... 56 6.1.2 TAK1 is activated downstream of PAR-1-induced SOCE ...... 58 6.1.3 Characterization of inducible EC-specific TAK1 knockout mice ...... 58 6.1.4 Loss of Endothelial-TAK1 augments PAR1-induced lung vascular leak and polymicrobial sepsis induced death ...... 61 6.1.5 TAK1 deficiency augments PAR-1-induced Ca2+ entry in ECs ...... 61 6.1.6 TAK1 kinase activity is essential for terminating SOCE...... 64 6.1.7 PAR-1-SOCE-CaMKKβ axis activates TAK1 in ECs ...... 64 6.1.8 Intermediate role of TAK1 in mediating STIM1 phosphorylation in ECs ...... 67 6.2 Hypothesis 2: TAK1 activation downstream of SOCE inactivates GSK-3β via p38 MAPK, which in turn enhances β-catenin expression at endothelial AJs and thus restores vascular barrier function ...... 72 6.2.1 Decreased expression of β-catenin and VE-cadherin induced by TAK1 deficiency in ECs 72 6.2.2 GSK-3β regulates β-catenin expression ...... 72 6.2.3 Inducible EC-restricted GSK-3β deletion promotes restoration of lung vascular integrity .. 76 7 DISCUSSION II ...... 78 8 LITERATURE CITED ...... 82 9 VITAE ...... 95 10 APPENDIX ...... 98

LIST OF FIGURES

FIGURE PAGE

1. Schematic representation for endothelial adherens junction assembly ...... 5 2. PAR-1 induced ER store Ca2+ depletion and SOCE ...... 12 3. Domain Structure of STIM1 ...... 15 4. Signaling Mechanism for termination of SOCE ...... 17 5. Schematic representation depicting domain structure and post-translational modifications for TAK1 and its binding partners (TAB1, TAB2, TAB3) ...... 20 6. STIM1 knockdown in HLMVECs prevents thrombin-induced SOCE, VE-PTP phosphorylation...... 32 7. STIM1 knockdown in HLMVECs prevents thrombin-induced permeability increase, and disassembly of VE-cadherin (VE-cad) at endothelial AJs ...... 33 8. PAR-1-induced Ca2+ entry, phosphorylation of VE-PTP, and phosphorylation of VE-cad are impaired in Stim1∆EC mice ...... 35 9. PAR-1-induced increase in lung vascular permeability impaired in Stim1∆EC mice ...... 36 10. STIM1-dependent Pyk2 activation induces tyrosine phosphorylation of Pyk2 in ECs...... 38 11. STIM1-dependent Pyk2 activation induces tyrosine phosphorylation of VE-PTP and VE- cad in ECs...... 39 12. Thrombin as well as VEGF promotes VE-cad dissociation from VE-PTP in ECs ...... 40 13. SOCE-induced Pyk2 activation promotes tyrosine phosphorylation of VE-PTP and dissociation from VE-cad ...... 42 14. In vivo silencing of Pyk2 in mouse lung microvascular ECs abrogates PAR-1-induced phosphorylation of VE-PTP and VE-cad ...... 43 15. In vivo silencing of Pyk2 in mouse lung microvascular ECs abrogates PAR-1-induced lung vascular leak ...... 44 16. SOCE-induced Pyk2 activation triggers VE-PTP mediated Src phosphorylation in ECs .... 46 17. VE-PTP’s C-terminal tyrosine phosphorylation triggers Src activation in ECs ...... 48 18. Peptide derived from the Pyk2 phosphorylation site on VE-PTP prevents PAR-1-induced disassembly of VE-cad at endothelial AJs ...... 49 19. Peptide derived from the Pyk2 phosphorylation site on VE-PTP prevents PAR-1-induced permeability increase ...... 50 20. Model for induction of VE-PTP-dependent Src activation in ECs via PAR-1-SOCE-Pyk2 pathway ...... 51 21. PAR-1-STIM1-SOCE axis activates TAK1 in ECs ...... 59 22.TAK1 inhibition augments PAR-1-induced Ca2+entry and permeability ...... 57 23. Generation of TAK1iΔEC mice ...... 60 24. Persistent PAR-1-induced lung vascular leak and increased mortality in TAK1iΔEC mice ... 62 25. TAK1 deficiency augments PAR-1-induced Ca2+ entry in ECs ...... 63 26. TAK1 kinase activity is essential for terminating SOCE ...... 65 27. CaMKKβ signaling downstream of SOCE required for PAR-1 induced TAK1 activation and SOCE inhibition in ECs ...... 66 28. TAK1 activation downstream of SOCE is required for AMPKα activation in ECs ...... 68 29. SOCE-induced TAK1 activation selectively triggers p38β MAPK signaling in ECs ...... 69 30. TAK1 activation downstream of SOCE is required for phosphorylation of STIM1 ...... 70 31. Schematic representation for SOCE-mediated TAK1 activation responsible for STIM1 phosphorylation and thus inhibition of SOCE ...... 71

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LIST OF FIGURES (continued)

FIGURE PAGE

32. EC-restricted TAK1 deletion in mice downregulates AJs expression ...... 73 33. STIM1 deletion blocks thrombin induced GSK-3β inactivation ...... 74 34 TAK1 deletion blocks thrombin-induced GSK-3β inactivation and augments β-catenin ubiquitination ...... 75 35. Generation of tamoxifen-inducible EC-restricted GSK-3β knockout (GSK-3βi∆EC) mice .. 77 36. Proposed model for role of STIM1-mediated SOCE signaling in TAK1 activation which terminates SOCE and resolves lung vascular hyperpermeability ...... 81

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LIST OF ABBREVIATIONS

AJs Adherens Junctions AMPK 5' adenosine monophosphate-activated protein kinase ARDS Acute Respiratory Distress Syndrome CLP Cecal ligation puncture EBA Evans blue conjugated Albumin ECs Endothelial Cells ER Endoplasmic Reticulum GPCR G protein-coupled receptor GSK-3β Glycogen-Synthase Kinase-3 Beta HLMVECs Human Lung Microvascular Endothelial Cells HUVECs Human Umbilical Vein Endothelial Cells ICU Intensive Care Unit MAPK Mitogen Activated Protein Kinase NO Nitric Oxide PAR-1 Protease-activated receptor-1 PI3K Phosphatidyl Inositol 3-Kinase PKC Protein Kinase C PLC Phospholipase C PYK2 Protein Tyrosine Kinase 2 Beta ROCs Receptor-operated calcium channels RTP Receptor-Type Protein Tyrosine Phosphatase SCL Stem Cell Leukemia Sc-siRNA Scrambled-small interfering RNA SFK Src family kinases SOAR STIM1-Orai activating region SOCE Store-Operated Calcium Entry SOCs Store-Operated Calcium Channels STIM1 Stromal Interacting Molecule 1 STIM1∆EC EC-restricted Stim1 knockout TAK1 Transforming Growth Factor-β-Activated Kinase 1 TAK1i∆EC Inducible EC-restricted TAK1 knockout TER Trans-endothelial monolayer resistance TFPI Tissue Factor Pathway Inhibitor Thr Thrombin TM Thrombomodulin VE-cad Vascular Endothelial-cadherin VEGF Vascular Endothelial Growth Factor VE-PTP Vascular Endothelial Protein Tyrosine Phosphatase vWF von Willebrand factor

SUMMARY

The lung endothelium is a monolayer of cells which are strategically positioned in between the

blood and the airspace. The endothelium functions as a semi-permeable barrier which regulates the

exchanges of gas, fluids and nutrients between circulation and the underlying tissue. Endothelial barrier

dysfunction leads to unchecked fluid extravasation and lung edema formation, the hallmarks of sepsis.

Sepsis is a systemic inflammatory response syndrome to bacterial infection, and a common cause of death in hospitalized patients. ALI is in large part the result of lung vascular leak. There is currently no effective drug targeting the key molecular players that treats this devastating disease. Although sepsis- mediated ALI induced activation of pro-inflammatory cytokines and coagulation cascade is well studied; mechanisms causing aberrant endothelial cell “activation” and leading to lung vascular leak and injury still remains an ill-defined concept. Thus, understanding of the innate mechanisms regulating lung vascular EC behavior is critical to the development of novel therapeutics that can more precisely target

ALI.

Previous studies from our laboratory have shown that thrombin, an inflammatory pro-coagulant mediates lung vascular leak by activating the G protein-coupled receptor (GPCR), protease-activated receptor-1 (PAR-1) on the EC surface. Further, studies have showed that PAR-1-induced Ca2+-entry in

lung ECs mediates lung vascular leak. Particularly, studies from various groups have elucidated the

mechanism of the endoplasmic reticulum (ER) localized Ca2+ sensor protein, stromal interacting molecule

1 (STIM1) in activation of store-operated calcium entry (SOCE) via store-operated calcium channels

(SOCs). Thrombin-induced ER-store Ca2+ depletion results in clustering of STIM1 at the ER/plasma

membrane interface, which then binds to and activates SOCs present in the plasma membrane. Since

thrombin-induced disruption of endothelial barrier is transient process regulated by SOCE, I hypothesized

in the present study that STIM1-dependent SOCE activates two different pathways for regulation of

endothelial barrier function. Firstly, it triggers protein tyrosine kinase 2 beta (Pyk2) activation which results

in VE-PTP-phosphorylation mediated VE-cadherin disassembly at endothelial AJs, and thus increases

vascular permeability response to thrombin. Secondly, it leads to Transforming growth factor-β-activated

kinase 1 (TAK1) activation which inhibits SOCE via phosphorylation of STIM1 and also increases

expression of adherens junction proteins, thus restoring lung vascular barrier integrity after injury.

In first hypothesis of my thesis work, I predicted that STIM1-mediated SOCE signaling promotes tyrosine phosphorylation of VE-cadherin, which is known to cause endothelial barrier destabilization. VE-

PTP, an endothelial cell-specific tyrosine phosphatase that associates with VE-cadherin at endothelial

Adherens Junctions (AJs), stabilizes VE-cadherin through dephosphorylation of its tyrosine residues.

Here I show that STIM1-mediated Ca2+ entry in endothelial cells (ECs), induces tyrosine phosphorylation

of VE-PTP activating the Ca2+-dependent protein tyrosine kinase Pyk2, which is required for Src kinase-

mediated tyrosine phosphorylation of VE-cadherin to increase permeability. In human lung microvascular

ECs, protease-activated receptor 1 (PAR-1) agonist thrombin induced Pyk2 phosphorylation at Y402

(activation) in a STIM1-dependent manner, which caused tyrosine phosphorylation of VE-PTP leading to increased VE-cadherin phosphorylation at Y685 and Y731 and subsequent endothelial hypermeability response. In agreement with these results, in EC-restricted Stim1 knockout (Stim1∆EC) mice, these PAR-

1-induced responses were abolished. Interestingly, silencing of Pyk2 had no effect on PAR-1-induced

Ca2+ entry, but blocked PAR-1-induced tyrosine phosphorylation of VE-PTP, Src binding to VE-PTP, Src activation, and VE-cadherin phosphorylation. Moreover, in vivo silencing of Pyk2 in mouse lung

microvascular ECs by liposome-mediated delivery of siRNA, prevented PAR-1-induced tyrosine phosphorylation of VE-PTP, Src activation, and VE-cadherin phosphorylation, and also markedly reduced

PAR-1-induced lung vascular permeability increase. Importantly, a cell permeable peptide

(1977LFPIYENVNPEY1988) derived from the C-terminal of VE-PTP prevented thrombin-induced Src activation in ECs. These findings demonstrate that 1) PAR-1-induced Pyk2 activation secondary to

STIM1-activated Ca2+ entry promotes tyrosine-phosphorylation of VE-PTP, and 2) that tyrosine- phosphorylated VE-PTP binds to and activates Src, which in turn phosphorylates VE-cadherin at endothelial AJs to increase lung vascular permeability.

In second hypothesis of my thesis work, I predicted that in ECs SOCE-mediated TAK1 activation

triggers the repair of leaky endothelial barrier. Here, I show that TAK1 activation downstream of PAR-1 inhibits SOCE and signals the reassembly of endothelial AJs. I observed that thrombin induced a time- xi

dependent phosphorylation of TAK1 at Thr-187 (which is essential for activation) in lung endothelial cells

(ECs). In addition, silencing of STIM1 suppressed thrombin-induced TAK1 phosphorylation in ECs, indicating that SOCE is essential for PAR-1-mediated TAK1 activation. Further, I used a cell-permeable specific inhibitor for irreversibly inhibiting TAK1 and observed augmented thrombin-induced permeability increase both in vitro and in intact lung microvessels. Consistent with these findings, inhibition of TAK1 impaired reassembly of VE-cadherin at AJs after thrombin treatment. I also observed that thrombin- induced SOCE was markedly increased in TAK1 deficient ECs. Since global as well as EC-restricted

TAK1 knockout mice are embryonically lethal, to study the in vivo role of EC-expressed TAK1, I generated tamoxifen-inducible EC-restricted TAK1 knockout (TAK1iΔEC) mice. Surprisingly, I observed markedly

reduced expression of VE-cadherin and its associated protein β-catenin in lungs of TAK1iΔEC mice

compared with their wild type littermates. I also observed enhanced lung vascular permeability as

assessed by Evans blue dye-conjugated albumin uptake in TAK1iΔEC mice. These findings support the

notion that TAK1 signaling in endothelial cells is required to maintain lung vascular barrier integrity.

To support second hypothesis of my thesis, I generated endothelial cell-specific inducible knockout mice for glycogen-synthase kinase-3 beta (GSK-3β) to address the notion that GSK-3β inactivation is a critical mechanism for the re-annealing of endothelial barrier after PAR-1-induced endothelial barrier disruption. Thus, I performed preliminary studies using GSK-3β mice to study whether

GSK-3β modulates lung vascular barrier integrity through control of β-catenin expression at AJs.

Interestingly, I observed augmented expression of β-catenin in LECs of GSK-3βi∆EC mice compared to

WT littermates. Next, I determined whether augmented β-catenin expression enhances the endothelial barrier. Here, I observed that PAR-1-induced lung vascular leak was substantially reduced in lungs of

GSK-3βi∆EC mice as compared with WT mice. These findings support the notion that TAK1 signaling in

endothelial cells is required to maintain lung vascular barrier integrity.

Thus, targeting the Pyk2 and TAK1 activation pathways may be potentially important strategies

to combat vascular leak associated pulmonary edema in sepsis.

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1 LITERATURE REVIEW

1.1 Endothelial cells

Endothelial structure and function

In the past, the endothelium was considered to be an inert “cellophane wrapper” lining the

entire vascular tree as a selectively-permeable monolayer membrane(1). However, enormous advances

since the mid-nineteenth century has led to the current view of the endothelium as a dynamic,

disseminated and heterogeneous organ of paramount importance with vital synthetic, secretory, immunologic and metabolic functions. In adults, approximately ten trillion (1013) endothelial cells cover a

surface area of approximately 1-7 m2 to form an almost 1 kg ‘organ’. Endothelial cells (ECs) are

essentially modified squamous epithelial cells; that regulate the flow of diverse nutrient and biologically active molecules and blood cells. The role of the ECs as a gate-keeper is effected through the presence

of various membrane-bound receptors for numerous molecules including proteins, lipid-transporting

particle, hormones, metabolites, as well as through specific junctional proteins and receptors that regulate cell-cell and cell-matrix interactions (2). Endothelial cells originate from embryonic precursor cells known as hemangioblasts. Further, ECs can also transdifferentiate into mesenchymal and intimal smooth muscle cells. Interestingly, there is distinct phenotypic variation between ECs from different parts of the vascular system, moreover they express different markers depending upon the location and generate different responses to the same stimulus (3). Based upon their direct contact with blood or lymph, ECs are known as vascular or lymphatic endothelial cells, respectively.

Vascular endothelial cells: The vascular endothelium, strategically positioned at the interface

between the tissue and blood, plays a vital role in maintaining the integrity of the blood vessels, blood

fluidity and blood flow (4,5). The vascular ECs are also critically involved for preventing inappropriate thrombogenesis, through expression of inhibitors of coagulation pathway including tissue factor pathway

inhibitor (TFPIs), thrombomodulin (TM), protein S, von Willebrand factor (vWF), heparin-like molecules.

In addition, vascular ECs also maintain the blood flow through production of mediators such as Nitric

Oxide (NO) which are responsible for regulating the tone of the surrounding smooth muscle cells (6,7).

1 2

The smallest of the body’s blood vessels are called capillaries, which are classified into three main

groups: Continuous capillaries (create an uninterrupted lining by forming tight and adherens junctions

[AJs] with adjacent ECs, which only allow smaller molecules such as water and ions to pass through intercellular clefts), fenestrated capillaries (ECs forming discontinuous junctions with pores that allow small molecules and limited amount of protein to diffuse), and sinusoid capillaries (similar to fenestrated capillaries but have larger pores with allow red and white blood cell, and various serum proteins to pass through). The continuous capillaries are found in the skin, lungs, skeletal muscles and the central nervous system, however discontinued capillaries are found in the liver, kidney, spleen and pancreas. Vascular

ECs are responsible for trafficking of essential nutrients, macromolecules, plasma proteins and cells such as monocyte, leukocyte, lymphocyte, in a highly specific and regulated manner. Hence, they are multifunctional system that regulates immune responses, inflammation and coagulation and fluid balance; and endothelial dysfunction can give rise to myriad complications leading to complex pathological states

(7).

Endothelial barrier

Endothelial cell-to-cell junctions are the essential units of endothelial monolayer. In addition to maintaining intercellular adhesion, they also transfer intracellular signals that can modulate inhibition of cell polarity, cell growth, lumen formation, and interactions with pericytes and smooth muscle

cells (8). Thus, any condition that can disrupt the endothelial junctions might not only elevate endothelial

permeability but also alter their responses to the surrounding cells and environment (8). Endothelial

adherens junctions (AJs) have been extensively studied by numerous groups (8-11) and it has been established that the vascular endothelial cadherin (VE-cad) serves as the “cornerstone” for sealing the endothelium.

VE-cad belongs to a classical superfamily of cadherins, and forms Ca2+-dependent homophilic

cis and trans dimers at AJs (12). The cytoplasmic domain of VE-cad interacts with intracellular proteins, including β-catenin, plakoglobin and p120-catenin, which transfer intracellular signals and modulate interaction with the actin cytoskeleton (Fig. 1) (13). Expression of VE-cad is endothelial-specific and

extremely important such that VE-cad knockout in mouse embryos is lethal, owing to severe angiogenic

defects (14,15). Further, arresting VE-cad expression in embryos and adult mice perturbs vascular

integrity (16,17). Most importantly, various in vitro studies utilizing silencing of VE-cad expression and

blocking its adhesive function, show that VE-cad is essential for AJs formation and maintenance of

endothelial barrier (18-21). Thus, VE-cad is regarded as the architect of the endothelial cell-cell junctions,

as it dictates the expression level and/or localization of other junctional molecules, including N-cadherin and claudin-5 (20,22,23).

Phosphorylation of VE-cad and its associated proteins (β-catenin, p120 catenin) have been shown to destabilize endothelial AJs (24,25). Published evidence supports the concept that inflammatory mediators (e.g., VEGF, bradykinin, TNF-α, and LPS) destabilize endothelial AJs by inducing phosphorylation of VE-cad at tyrosine residues (Y568, Y685, and Y731) to increase vascular permeability and leukocyte extravasation (26-29). Recently, Wessel et al (30) have shown that knock-in mice expressing a Y685F mutant VE-cad exhibited impaired vascular permeability responses whereas mice expressing the Y731F mutant showed decreased neutrophil-extravasation, suggesting that phosphorylation of VE-cad at different tyrosine residues differentially regulate vascular permeability and leukocyte extravasation. Inflammation triggering mediators have been shown to induce endothelial barrier dysfunction by disassembling endothelial AJs via c-Src-dependent phosphorylation of VE-cad and its binding partner p120 catenin in ECs (24,25). Several protein tyrosine phosphatases have been shown to regulate endothelial barrier function by controlling the phosphorylation of adherens junctional proteins

(26). A large amount of emerging evidence now supports the concept that vascular endothelial protein tyrosine phosphatase (VE-PTP), exclusively expressed in endothelial cells, interacts with VE-cad at endothelial AJs to maintain the endothelial barrier integrity in a phosphatase dependent manner

(26,31,32). Besides its role in tightly controlling the VE-cad at endothelial AJs, VE-PTP also interacts

with Tie-2, an endothelial cell-expressed tyrosine kinase receptor which regulates angiogenesis and

endothelial barrier integrity (31,32).

VE-PTP (also known as PTPRB or PTPβ) is a receptor-type protein tyrosine phosphatase (RPTP) of the R3 subtype (33). The extracellular region of VE-PTP contains multiple fibronectin type III-like domains and the cytoplasmic region has a single catalytic domain (34). VE-PTP binds to VE-cadherin via VE-PTP’s extracellular domains (35). It has been suggested that binding of a phosphorylated VE-

PTP substrate to the cytosolic domain of VE-PTP could lead to detachment of the extracellular domain of VE-PTP from VE-cadherin (36). All R3 subtype RPTPs contain a conserved amino acid motif YxNФ

(where Ф is a hydrophobic amino acid and x, any amino acid) in the C-terminus (34). This YxNФ motif undergoes tyrosine phosphorylation which in turn binds to the SH2 domain of Src family kinases (SFKs).

This event activates SFK (34,37). Both human (h) and mouse (m) VE-PTP’s deduced sequence contains the amino acid motif “YENV” (h residues 1981-1984; m residues 1982-1985) in the C-terminus. However, whether inflammatory mediators induce VE-PTP phosphorylation at Y1981 in the cytosolic domain is unknown as is the tyrosine kinase inducing this phosphorylation. Also, whether phosphorylated VE-PTP

binds to and activates Src, and whether Src in turn induces VE-cadherin phosphorylation to promote endothelial barrier destabilization is also unknown.

VE-PTP

β-cat α-cat VE-cad α-cat α-cat Actin p120 Intact Ca2+ EC AJs

EC AJs Permeability ↑↑ Mediators: Disassembly Transmigration ↑↑ Thrombin, VEGF, LPS etc.

Figure 1. Schematic representation for endothelial adherens junction assembly. VE-cadherin forms Ca2+-dependent homophilic dimers which mediates adhesion with adjacent endothelial cells. The cytoplasmic domain of VE-cadherin binds to β-catenin which in turn recruits α-catenin to the AJs. α- catenin directly interacts with actin and mediates formation of cortical actin bundles. On the contrary, VE protein tyrosine phosphatase (VE-PTP) is an endothelial-specific receptor phosphatase which associates with VE-cadherin through extracellular protein domains. Under basal conditions, this association enhances VE-cadherin-mediated adhesion in mammalian cells in a phosphatase- dependent manner. However, upon endothelial activation with mediators (such as Thrombin, VEGF, LPS etc.) certain structural and conformational changes that occur across the membrane, leads to detachment of VE-PTP from the VE-cadherin AJs complex. This promotes phosphorylation of components in the VE-cadherin-catenin complex and thus destabilization of endothelial cell contacts. Disassembly of endothelial AJs is the foundation for increased endothelial permeability and leukocyte transmigration across the vascular barrier.

Endothelial dysfunction and inflammation

Inflammation is a defensive mechanism that is triggered by noxious stimuli and conditions, such as

pathogens and tissue injury (38). At the site of infection, macrophages are responsible for phagocytosing

the bacteria and release of various kinds of proinflammatory mediators such as cytokines (eg. TNF-α

and interleukins) and chemokines. Activation of ECs occurs in two distinct phases, type I (immediate response) and type II (slower response) (7). Both types of activation phases induce the fundamental signs of inflammation: rubor (enhanced blood flow resulting in red color), calor (warmth as a result of inflammation of the tissues), tumor (swelling caused by endothelial contraction leading to the leakage of the interstitial fluid) and dolor (pain due to the release of additional inflammatory mediators by the activated leukocytes recruited in the damaged tissues) (4,7).

1.2 Sepsis

Pathogenesis

Sepsis, a systemic inflammatory response syndrome, is one of the oldest and most elusive disorder in

medicine (39). The term “Sepsis” was originally introduced by Hippocrates (ca. 460-370 BC), derived

from the Greek word sipsi ("make rotten"). However, in 1989 US-American ICU specialist Roger C. Bone

(1941-1997) offered a sepsis definition that is still valid until today: "Sepsis is defined as an invasion of

microorganisms and/or their toxins into the bloodstream, along with the organism's reaction against this

invasion". In the United States, severe sepsis is documented in two percent of patients admitted to the

hospital, of which half are treated in the intensive care unit (ICU). The total number of reported cases in

the US exceeds 750,000 per year and is increasing every year (40). Sepsis is usually characterized by

two or more of these criteria, changes in body temperature, respiratory rate, heart rate and the white

blood cell count (39). It also causes inappropriate regulation of a multitude of cell types, proinflammatory

mediators and pro-coagulation factors. Depending upon its severity, sepsis can result in multi-organ

failure. Sepsis has been shown to be primarily induced by gram-negative bacterial infection (62% gram-

negative infection, 47% gram-positive infection and 19% fungal infection) (40). Sepsis commonly affects

the respiratory and cardiovascular systems, and its classical manifestations are acute respiratory distress

syndrome (ARDS) and myodcardial dysfunction (40).

The mortality rates due to sepsis (regardless of the type of infection triggering the onset of sepsis) continue to rise (41). In addition, antibiotic therapies directed against the endotoxin have largely failed during clinical trial suggesting that host response plays a crucial role in determining the severity of sepsis.

Pathogens are recognized by host cells (monocytes, macrophages and endothelial cells) through the pattern recognition receptors (such as Toll-like receptor or TLRs) for triggering a host response or its first line of defense. The host response then leads to release of soluble mediators which are responsible for enhancing the effects of the inflammatory and the coagulation cascades. However, during the host’s attempt to eliminate pathogens, unchecked inflammatory response can inflict collateral damage on normal tissues causing focal distribution of the disease state (41).

1.3 Thrombin

Structure and function

Thrombin is a trypsin-like serine protease which plays a critical role in homeostasis(42). It is involved in regulation of various physiological and patho-physiological mechanisms such as inflammation, coagulation, anti-coagulation and atherogenesis (43). It cleaves substrates in a highly selective and specific manner; guided by the “classical” active site which is enveloped with critical features, loops and charged patches. In addition, there are two other recognition sites, anion-binding exosite 1 (a.k.a. fibrinogen recognition site) and anion-binding site 2 (a.k.a heparin-binding site), which interact with complementary sites of specific substrates, receptors and inhibitors and thus implement crucial regulatory functions (43). Allosteric regulation of thrombin function occurs through another loop with Na+ binding site

(42,43). There are two Na+ binding sites present. The occupancy of the site leads to conformational changes in thrombin (viz. “slow” [Na+-free] or a “fast” [Na+-bound] form) (43). The fast form has higher

affinity and catalytic activity toward fibrinogen, promoting pro-coagulant activities. However, the slow form selectively actives Protein C, resulting in anti-coagulant activities of Thrombin (43). Thus, allosteric

regulation is substantial for thrombin function. Ultimately, generation of thrombin activates diverse

downstream signaling pathways resulting in calcium influx, cytoskeletal reorganization, release of growth

factors, soluble mediators, metalloproteinases and enhanced expression of genes involved in cell

proliferation, leukocyte adhesion, vasomotor tone, inflammation and hemostasis (44).

Coagulation

The activation of coagulation cascade can occur through two distinct pathways which are dependent upon its mode of initiation, the extrinsic (tissue factor) pathway and the intrinsic pathway (surface contact)

(45). Both pathways assemble at the thrombin production step, where factor Xa proteolytically cleaves its substrate prothrombin. The thrombin generated then acts on fibrinogen, converting it into fibrin (45).

Coagulation relies on multitude of soluble factors produced in the liver, circulating as inactive zymogen forms in the plasma. These inactive forms are denoted with roman numbers and the active forms with a lower case “a” following the Roman number (45). These factors have been classified accordingly: Factor

I [Fibrinogen], Factor II [Prothrombin], Factor III [Tissue Factor], Factor IV [Calcium], Factor V

[Proaccelerin], Factor VII [Proconvertin], Factor VIII [Antihemophilic factor], Factor IX [Christmas factor],

Factor X [Stuart-Prower factor], Factor XI [Plasma thromboplastin antecedent], Factor XII [Hageman factor] and Factor XIII [Fibrin stabilizing factor] (45). Most of the factors (except Factor XIII) are serine proteases in their active form and are similar to the digestive enzyme trypsin. Other factors including

Factor V, Factor VIII, tissue factor and molecular weight kininogen (HK) act as co-factors.

a) Extrinsic pathway: Extrinsic pathway is the primary mechanism of thrombinogenesis in the

blood vessels which is triggered upon exposure of the tissue factor on the surface of the endothelial cells

and monocytes (43). Tissue factors are glycoproteins composed of cytosolic, transmembrane and

extracellular domains; they are expressed on the sub-endothelium of blood vessels and thus unexposed

to circulating blood. Activation of tissue factor requires phospholipid which ensures coagulation complex

formation at the injury site. Factor VII consists of 2 chains and circulates in the blood as an active

protease; VIIa is cleaved at Arg152, a conformation which restricts plasma activation (43,45). However,

upon binding to tissue factors, the conformation changes into an active form resulting in activation of factors IX and X.

b) Common pathway: This is the convergence point of both intrinsic and extrinsic pathways.

Factor II, V and Xa make up the “prothrombinase” complex that plays a central role in generating thrombin

(45). Factor Xa cleaves prothrombin to produce thrombin, which in turn cleaves fibrinogen to expose the

center that enables bulbous ends of other fibrinogen to tether together. Fibrins comprised of β and γ

chains are produced which form a meshwork surrounding the site of injury and also capturing about 40%

of the available thrombin to protect it from innate inhibitors (45). To ensure formation of fibrin at the desired location and not being washed off by the circulating blood, two methods are mainly employed.

Firstly, platelets bind and stabilize the fibrin strands through their phospholipids, and an array of inhibitors constrain the platelet hemostatic plug to the injured site (45). After activation, a platelet-specific enzyme flippase leads to reversal of charged phospholipids on the platelets to direct the negatively charged phosphatidylserines to face outward. On the contrary, the Vitamin K dependent coagulation factors carry negatively charged glutamic acids at their N-terminal regions. Ca2+ ions counteract the negatively charged

acid phospholipids and proteins and induce conformational change on the latter stabilizing the platelet

plug (45). Secondly, natural inhibitors such as, heparin cofactor II (HCII), antithrombin III (ATIII), thrombomodulin, protein C and tissue factor pathway inhibitor (TFPI) in the blood restrain the platelet

plug at the site of injury by various mechanisms (45).

1.4 Protease-activated receptors (PAR)

The PARs belong to a G-protein coupled receptor (GPCR) super family which consists of, PAR-1, PAR-

2, PAR-3 and PAR-4. Signal transduction via PARs are sustained intracellularly through heterotrimeric

G-proteins that bind to the intracellular domain of the receptor (46). Upon activation, G-proteins couple to intermediates including phospholipase C (PLC), protein kinase C (PKC), phosphatidyl inositol 3-kinase

(PI3K), mitogen activated protein kinase (MAPK) and AKT (44). Mammalian PAR activation occurs via a number of trypsin-like serine proteases such as thrombin, prothrombin intermediates, Factor Xa, trypsin

IV, granzyme A and activated protein C. Thrombin is known to specifically activate PAR-1, PAR-2 and

PAR-4 receptors which are coupled to different G-proteins including Gα (Gαq/11, Gα12, Gα13, G0 and Gi), Gβ

and Gγ, thus in a stimulus and tissue dependent manner various intracellular signaling cascades get activated (43). PAR-2 is activated by the coagulation proteases VIIa and Xa; instead of thrombin (47).

Studies have shown that human umbilical vein endothelial cells (HUVECs) express predominantly PAR-

1 receptors with a lesser extent of PAR-2, PAR-3 and PAR-4 (44). In addition, various genetic studies performed with single knockout (PAR-1/PAR-2/PAR-4) or double knockouts (PAR-1/PAR-2; PAR-2/PAR-

4) support PAR-1 being the main thrombin receptor on the microvascular endothelial cells (47). A direct co-relation among PAR-1 and Ca2+ concentration has been established in the human microvascular

endothelial cells as well by utilizing PAR-1 knockout mice (48,49).

a) Activation: Thrombin binds to a hirudin-like domain on the extracellular sequence of PAR-1 and

cleaves it between Arg41 and Ser42. Thereafter, the new N-terminus is exposed (44,45). The tethered

ligand (SFLLRN) then interacts with the extracellular loop 2 (aa 248 to 268) of the receptor, leading to

its activation. Following agonist binding, PAR-1 binds to Gαq/11, resulting in phospholipase C (PLC) β

(43) which then cleaves phosphatidylinositol-4, 5-bisphosphate (PIP2) to yield secondary messengers

(diacylglycerol (DAG) and inositol 1,4,5-triphosphate (IP3)). IP3 induces the downstream signaling which

leads to Ca2+ mobilization from the endoplasmic reticulum (ER) into the cytosol (46). On the contrary,

DAG activates PKCα which causes phosphorylation of VE-cadherin junctional proteins to disassemble

VE-cadherin junctions. Moreover, PKCα can activate MLC to generate contractile forces and thus further

enhance endothelial retraction by activating Rho-kinase (50).

1.5 Calcium Signaling

Types & function

Calcium acts as an essential second messenger in almost all eukaryotic cells. The significance of extracellular Ca2+ influx inside the cell in maintaining contraction of isolated hearts was first initially recognized by Ringer (51). The cellular Ca2+ concentration is strictly sustained at ~100 nM and variation

in this concentration triggers a plethora of cellular responses such as muscle contraction, neurotransmitter release, cell growth, cell proliferation, mitochondrial metabolism and gene expression

(50,52). Unrestrained Ca2+ release can result in major pathophysiological conditions including apoptosis,

necrosis, inflammation and sepsis (52). Extensive research from our laboratory and other groups have

shown a direct link between elevated calcium levels and endothelial hyper-permeability (50,53-55).

Calcium entry inside cells occurs via three distinct mechanisms: Voltage-dependent calcium channels

(VDCCs), Receptor-operated calcium channels (ROCs) and store-operated channels (SOCs) (50,56).

The VDCCs are located in the membranes of excitable cells including neurons, muscles and glial cells.

Their activation occurs upon depolarization of the membrane potential, therefore called “voltage- dependent” (55). Calcium measurements unveiled that there are periodic rises of cellular calcium

concentration originating from a fixed point and propagated with waves. This phenomenon is referred to

as calcium oscillations (57). However, in non-excitable cells such as endothelial cells, numerous studies

have established store-operated Ca2+ entry as the major calcium influx pathway (58,59).

Store-operated Ca2+ entry

In 1986, Putney proposed the idea of capacitative Ca2+ entry whereby he theorized that agonist-

stimulated production of second messengers could deplete the ER-stored Ca2+ (60). This spike in cellular

Ca2+ concentration could elicit sustained increase in Ca2+ influx in the cell from the extracellular medium in non-excitable cells including endothelial cells, mast cells, t cells and thymocytes (61). In 1993, Hoth

and Penner identified Ca2+ release activated Ca2+ current (CRAC current) in mast cell, which is a highly

Ca2+ selective, non-voltage gated and inwardly rectifying (62). SOCE consists of two distinct phases, activation of EC membrane-localized receptors including GPCRs (PAR-1), receptor protein tyrosine kinase (RPTK) triggers the transient release of ER-stored Ca2+, followed by sustained calcium influx from the extracellular medium to the cytosol. After ECs stimulation, activated phospholipase C (PLC) generates a second messenger, diacylglycerol (DAG) which is a known activator of the receptor-operated calcium channels (55) [Fig. 2 ]. Studies from our laboratory and other groups have established a crucial role between the SOCs and endothelial permeability associated with inflammation(55). An inflammatory

mediator such as thrombin, bradykinin, histamine and oxidants can stimulate endothelial cells resulting

in downstream signaling which leads to IP3 generation by PLC. IP3 binds and activates IP3 receptor (IP3R),

located on the ER membrane (55). There are different IP3Rs (IP3RI, IP3RII and IP3RIII) approximately

300 kDa each and belonging to the ryanodine channel family. They have cell-specific expression besides

exhibiting varying sensitivity to IP3; IP3RII being the most sensitive and IP3RIII the least (55). IP3R consists of tetrameric structures and are controlled by both concentration of its ligand IP3

2+ Thr Ca PAR-1

SOCs

PLC-β Ca2+↑↑ STIM1 IP3 Ca2+

IP3R ER

Figure 2. PAR-1 induced ER store Ca2+ depletion and SOCE. PAR-1 activation initiates downstream signaling leading to PLC activation and inositol 1,4,5-triphosphate (IP3) generation. IP3 binds to IP3 receptors on ER and result in ER store Ca2+ depletion. This is followed by STIM1 assembly into “puncta” at the ER/plasma membrane interfaces to interact with SOCs and to activate Ca2+ in ECs.

along with the intracellular concentration of Ca2+. Each isoform has consensus phosphorylation site for

kinases including, Ca2+/dependent kinase II (CamKII), AMP-dependent kinase II (PKA) and PKC (55).

2+ Activation of IP3R induces Ca release from ER-store, which in turn activates an ER-membrane localized

protein, Stromal Interacting Molecule 1 (STIM1). STIM1 then oligomerizes and directly interacts with the

cell-membrane localized channels eliciting SOCE (63). This results in Ca2+ influx into the cells and a

2+ sustained increase in [Ca ]i (intracellular calcium concentration).

1.6 Store-operated Ca2+ channel [SOC]

Stromal Interacting Molecular proteins (STIMs)

In the past, various theories were proposed to define the mechanism of stored calcium release from the

ERs which leads to opening of the channels on the plasma membrane. In 2005, research from two independent laboratories Roos et al (64)and Liou et al (65) established STIM proteins to be the pivotal in the ER-plasma membrane junctional coupling model, which was suggested more than a decade ago by

Putney (66,67) and Berridge (66). Roos et al., through his RNA interference (RNAi) studies in Drosophila melanogaster S2 cells, identified a single STIM1 protein (64). Moreover, Liou et also identified a pair of

STIM1 proteins by monitoring calcium signaling in HeLa cells (65). These proteins were characterized as tumor-suppressor gene product (68,69), located on the stromal cell surface. Interestingly, its original function was deduced to be a mediator of interaction among Stromal and haematopoietic cells, therefore the acronym, STIM1. It has two isoforms: STIM1 and STIM2 (70,71), both have structural similarities except in the amino- and the carboxy-terminals. In addition, both are known to be located in the ER- membrane, however certain studies suggest the presence of STIM1 in the plasma membrane to some extent. They are ubiquitously expressed in various cell types of vertebrates. Although STIM1 expression

level is much higher than STIM2 in most tissues, brain and dendritic cells are known to express the latter

predominantly (70). STIM proteins are known to induce Ca2+ influx via interaction with SOC [TRPC family] and ICRAC [Orai] present on the plasma membrane (70).

a) STIM1: Using deletion studies in STIM1 to identify the functions of each domain, STIM1 has been identified as a single pass trans-membrane protein which has amino terminal Ca2+ binding motif in the ER lumen (72,73). The N-terminus or the luminal domain consists of clusters of short α-helices

containing two EF-hand domains [a Ca2+-binding canonical EF-hand(cEF) domain and non-Ca2+ binding

hidden EF-hand (hEF) domain) and a sterile α-motif (SAM) domain with Asn-linked glcosylation sites

[Fig. 3]. The EF-hand domain senses slight short variations of Ca2+ concentration in the ER-lumen. ER- store Ca2+ depletion results in Ca2+ dissociation from the EF-hand, thereby unfolding and destabilizing the

N-terminal domain. This in turn results in STIM1 activation, which triggers oligomerization through the

SAM domain and stabilizing the exposed hydrophobic residues (73). The C-terminus is cytosol-facing

containing the Ezrin-Radixin-Moesin (ERM) domain, which consists of C-C (coiled coil domain), SOAR

(STIM1-Orai activation region). It is followed by inhibitory domain (ID), S/P (serine-proline) rich domain,

a microtubule interacting domain (TRIP) and the lysine (K) rich domain (73). The CC domain contains

CC1, CC2 and CC3 domains and the CC1 is further subdivided into three α-helices termed Cα1, Cα2 and Cα3. The Cα3 helix consist of acidic sequence EEELE, which inhibits SOAR function. The CC2

(363-389) and CC3 (399-423) regions extend into the SOAR domain.

The STIM1-STIM1 interactions are mediated by CC regions, in resting as well during activation phase.

The SOAR domain is the kernel for activation of STIM1 and is crucial for Orai1 activation. SOAR contains

KIKKKR (polybasic active site), which plays crucial role for interaction between STIM1 and Orai1 (73).

There are four types of α-helices found in SOAR: Sα1, Sα2, Sα3 and Sα4. Following SOAR another

inhibitory domain (ID) is present that inactivates Orai1. Next, a conserved sequence of lysine-rich domain

(K-domain) is present which is required for activation of SOCs such as TRPC channels. The K-domain has been shown to be facilitating channel activation via interaction of its positively charged lysine residues with the anionic phospholipids found in the membrane. Moreover, the K-domain supports junctional assembly during STIM1 redistribution at close proximities to the plasma membrane (73). The ERM and

K-domain have been shown to be indispensable and adequate to bind and activate the TRPCs. All

TRPCs are known to interact with the SOAR domain of STIM1, except TRPC7. The specific STIM1-

TRPC1 electrostatic interaction has been shown to occur between the basic residues at C-terminus

(K684 and K685) and acidic residues at the C-terminus (D639 and D640), respectively (72-75). TRPC4 also contains acidic amino acid residues in the C-terminus which can interact with STIM1. This biochemical interaction during SOCE was further supported with immunostaining and immune-

precipitation studies by our laboratory (76). Upon ER-stored Ca2+ depletion, STIM1 proteins undergo oligomerization and accumulation in the ER/plasma membrane interface, this results in discrete punctae

formation which is known to interact with the SOCs (TRPCs, Orai1) and thus activate SOCE (63,73).

Punta formation is a reversible and systematic process which is facilitated by microtubule formation. The sites for puncta are predetermined by scaffolding and structural molecules which act as place markers

(57).

1-29 67 – 96 132 – 200 215-234 251 - 43 383 389 475-483 535 600-629 672 -685

c/h EF SAMSAM TM CCC-C SOAR S/PS/P KK ERM

1 685

TM S486 S575 S600 S492 S628 S608 S621 S668 S618 T626 2+ Figure 3. Domain Structure of STIM1 is shown in A. STIM1 N-terminus contains Ca binding EF hands (67-96) which mediate STIM1 oligomerization and clustering upon ER store Ca2+ release. The transmembrane domain (215-234) anchors STIM1 in the ER. The Ezrin- Radixin-Moesin (ERM; 344- 442) domain has 2 coiled-coil domains (251- 343 and 383-389). The second coiled-coil domain is localized in the SOAR domain (344-442). The SOAR domain interacts with TRPCs. The K-domain gates TRPC channels by electrostatic interactions with conserved acidic residues (DD or EE) present in the TRPCs C termini. Phosphorylation of STIM1 at the C-terminus has been shown to inhibit Ca2+ entry during mitosis in HeLa cells. The cytosolic domain contains 21 Ser/Thr residues. Using software that predicts phosphorylation sites, we discovered the presence of 10 consensus phosphorylation sites (Ser/Thr residues) for p38 MAPK in the C-terminus of STIM1 (primarily in S/P domain) (B). In a recent study, we have shown that thrombin-induced p38 MAPK activation phosphorylates STIM1 which in turn inhibits Ca2+ entry through TRPC channels.

1.7 SOCE-mediated disruption of AJs

I have previously shown that thrombin induced vascular leak requires store-operated Ca2+ entry (SOCE)

through store-operated Ca2+ channels (SOCs) in endothelial cells (77) . In addition, we (76,77) and others

(78) have shown that the endoplasmic reticulum (ER) localized Ca2+ sensor protein, STIM1, is essential for activating SOCE in ECs. Importantly, LPS-induced lung vascular permeability increase and leukocyte

infiltration in lungs were abrogated in EC-restricted STIM1 knockout (Stim1∆EC) mice (79). These findings

support the notion that STIM1-activated Ca2+ entry (SOCE) in ECs is a critical determinant of endothelial

barrier dysfunction. However, the signaling pathways activated downstream of SOCE to cause

endothelial barrier dysfunction are not studied in the context of VE-PTP/VE-cad interaction mediated

endothelial barrier stability.

1.8 Termination of Store operated calcium entry and resealing of AJs

SOCE inhibition: Although SOCE activation is well studied, termination of SOCE to prevent calcium overload in cells remains a poorly understood area. However, studies done in the past decade have attempted to address the intricate mechanisms of SOCE inhibition.

STIM1 was initially recognized as a phosphoprotein with multiple serine (Ser) phosphorylation sites (80).

Research done in Hela cells by Smyth et al., demonstrated that STIM1-mediated Ca2+ entry was “turned

off” via phosphorylation of Ser-486 and Ser-668 residues at the C-terminus during mitosis (81). Moreover, they also unveiled that phosphorylation of STIM1 prevents store depletion-induced STIM1 puncta at ER- plasma membrane junctions, an event pivotal for SOCE activation. In addition, another study done in

HEK293 cells showed that STIM1 phosphorylation at ERK1/2 target sites modulates SOCE (82). Most importantly, recently published studies from our laboratory have established that SOCE induced by thrombin activates AMPKα1-p38β MAPK axis, which mediates STIM1 phosphorylation to serve as an “off switch” for SOCE in EC (Fig 4) (83).

Figure 4. Signaling Mechanism for termination of SOCE: PAR-1-induced ER-stored Ca2+-release activated SOCE. SOCE signal then activates CaMKKβ-AMPKα1-p38β MAPK signal axis, which leads to STIM1 phosphorylation for termination of SOCE in endothelial cells. Reprinted with permission from (appendix at end) Journal of Biological Chemistry, Sundivakkam, P. C., Natarajan, V., Malik, A. B., and Tiruppathi, C. (2013) Store-operated Ca2+ entry (SOCE) induced by protease-activated receptor-1 mediates STIM1 protein phosphorylation to inhibit SOCE in endothelial cells through AMP-activated protein kinase and p38beta mitogen-activated protein kinase. J Biol Chem 288, 17030-17041.

AMPK is an “energy-sensor” molecule which is activated in times of cellular energy deprivation, as a result of elevated levels of AMP. AMPK activation triggers ATP-generating processes such as glycolysis and lipolysis, and simultaneously shuts down ATP-depleting processes such as gluconeogenesis and lipogenesis. Therefore, AMPK is a key player for maintaining cellular homeostasis. In a screening study utilizing yeast-mutants deficient in upstream kinase of Snf1 (the yeast ortholog of mammalian AMPK),

TAK1 was identified as an upstream kinase of AMPK (84). In addition, TAK1 has also been shown to

activate AMPK in vitro (85). This study was further confirmed by Xie et al. who used a transgenic mice expressing a dominant-negative TAK1 and observed cardiac abnormalities resembling the Wolff-

Parkinson-White syndrome, which is known to be caused by mutations in AMPK (86). Although regulation of AMPK through TAK1 is unanticipated since there is no obvious connection between them. However,

TAK1 is known to be activated by certain stresses such as hypoxia and osmotic shock, which have also been shown to activate AMPK. Another study, showed that TRAIL can activated AMPK via TAK1 to regulate autophagy (mechanism for proteolysis to maintain cellular protein homeostasis (87).

AMPK is also a serine/threonine (Ser/Thr) protein kinase composed of a catalytic α-subunit and regulatory β- and γ-subunits (88,89). Thr-172 phosphorylation in the α-subunit is essential for catalytic function of AMPK (88,89). Two isoforms of the catalytic subunits (α1 and α2) are expressed in ECs and we have previously shown that SOCE induced the activation of AMPKα1 (83). We also observed in these studies that AMPKα1 activation in turn mediated the activation of p38β to induce STIM1 phosphorylation, and thus in negative feedback had the potential to inhibit SOCE (83). In addition, silencing of p38β enhanced thrombin-induced SOCE(83) and prevented re-annealing of endothelial barrier.

1.9 TAK1

Transforming growth factor-beta-activated kinase-1 (TAK1) is a protein kinase initially characterized as a MAPK kinase kinase (MAP3K7) activated by TGF-β and bone morphogenetic protein (BMP). It is well established as a crucial kinase in innate immune signaling pathways activated by various stimuli,

including cytokines, B-cell, T-cell, and Toll-like receptor ligands (90). TAK1 functions as a nodal kinase by phosphorylating IκB kinase, p38 MAPK, JNK, and ERK; thereby causing activation of NF-κB and

MAPK signaling pathways (91,92). TAK1 forms ternal complexes with its adaptor proteins TAB1 or

TAB2/TAB3 (93-95) which are essential for TAK1 activity (96). TAB1 and TAB2/TAB3 are known to bind

to N- and C-terminal domains of TAK1, respectively. TAB1 has been shown to bind constitutively to TAK1

and activate TAK1 kinase activity, whereas TAB2/TAB3 bind to TAK1 only after stimulation. TAB2/ TAB3

physically interact with TRAF6 through K63-linked polyubiquitin chains and thereby cause activation of

TAK1. Global deletion of TAK1 leads to severe abnormalities in development of neural tube in embryos as early as day 9.0 (90,97). Similarly, deletion of TAB1 and TAB2 also results in embryonic lethality

(98,99). Several post-translational modifications of TAK1 and TABs have been demonstrated to be regulating TAK1 kinase activity; including phosphorylation, ubiquitination and methylation (91). This warrants for assessment of TAK1 kinase activity specific to each stimuli and the cell type. Although several phosphorylation sites have been identified in the TAK1 activation loop (Thr-178, 184, 187 and

Ser-192), phosphorylation at Thr-187 has been shown to be crucial for activation of TAK1 kinase activity

(100,101).

C TAK1 N Kinase Domain TAB2/3 BD 579

p38 TAK1 C TAB1 N Pseudophosphatase domain BD BD 504

C TAB2 N CUE TAK1 BD NZF 693

C TAB3 N CUE TAK1 BD NZF 712

Figure 5. Schematic representation depicting domain structure and post-translational modifications for TAK1 and its binding partners (TAB1, TAB2, TAB3). TAK1 has kinase domain in the N-terminus, which is essential for autophosphorylation and thus activation of TAK1. Several phosphorylation sites for TAK1 have been identified, out of which T187 has been shown to be crucial for TAK1 kinase activity. In addition, TAK1 has binding domain for TAB2/TAB3 in the C-terminus region where polyubiquitinated TAB2 or TAB3 bind to mediate TAK1 activation. TAB1 is an adaptor which lacks any enzymatic activity but is constituently bound to TAK1 in cells. Although TAB1 has psudophosphatase domain it lacks critical residues for any phosphatase activity. TAB1 also has p38 and TAK1 binding domains in the C-terminus regioin. TAB2 and TAB3 each contain an N-terminal coupling of ubiquitin conjugation to endoplasmic reticulum–associated degradation (CUE) ubiquitin-binding domain (UBD), a TAK1-binding domain and a C-terminal Npl4 zinc finger (NZF) UBD (96).

1.10 Endothelial barrier repair and restoration

PAR-1 activation is known to increase endothelial permeability through cell contraction and disassembly

of adherens junctions (AJs) (102). The AJs is a multi-protein complex, which consists of the

transmembrane vascular endothelial cadherin (VE-Cad) and cytoplasmic catenins (p120; α, β and γ-

catenin). β-catenin is an essential component of the endothelial AJs complex, and is involved in the regulation of barrier function through its direct linkage with VE-Cad and the cytoskeleton (103). Although

2+ the mechanisms regulating AJs as a result of increased [Ca ]i still remain unclear, phosphorylation of β- catenin has been shown to alter AJs and thereby regulate cell-cell adhesion (104). Interestingly, one such

crucial factor regulating β-catenin activity, which also acts as an initial responder and effector of endothelial permeability maintenance is glycogen synthase kinase-3 beta (GSK-3β) (105). GSK-3β is a constitutively active kinase which phosphorylates β-catenin for ubiquitination and subsequent degradation. Absence of GSK-3β kinase activity allows stabilization and accumulation of β-catenin in the cytosol and thus translocation to the membrane to maintain AJs integrity (106,107). Regulation of β- catenin stability through GSK-3β; therefore, could be a potential mechanism for strengthening of endothelial barrier through AJs stability. Phosphorylation of GSK-3β by p38 MAPK has been shown to inactivate GSK-3β (108). Since p38 MAPK is a known target for TAK1 (90,109), suggesting the possibility that TAK1 acts upstream of p38 MAPK to regulate GSK-3β and thus stabilize junctional complex after

PAR-1 induced Ca2+ entry. Therefore, this warrants for elucidation of TAK1 function in stabilization of β-

catenin after PAR-1-induced disassembly of endothelial AJs.

2 OBJECTIVES

2.1 To determine the role of STIM1-activated Ca2+ entry [SOCE] in mediating vascular

leak through disassembly of AJs

2.2 To determine the role of TAK1 activation secondary to SOCE in restoring lung

vascular barrier integrity after injury, via termination of SOCE and stabilization of

β-catenin

3 MATERIALS AND METHODS

3.1 Materials

Hanks’ balanced salt solution (HBSS), L-glutamine, trypsin and Fura-2AM were purchased from

Invitrogen (Carlsbad, CA). Fetal bovine serum (FBS) was from Hyclone (Logan, UT). Human α-thrombin

was obtained from Enzyme Research Laboratories (South Bend, IN). All the peptides used for

experiments were synthesized by Genscript (Piscataway, NJ). Peptide purity and amino acid sequences

were determined by high-performance liquid chromatography and MS, respectively. PAR-1-activating

peptide (TFLLRNPNDK-NH2) was synthesized as the C-terminal amide with a purity of > 95%. VE-PTP’s

C-terminal peptides (h residues, 1977 – 1988; Wild type [WT]-LFPIYENVNPEY; mutant [mut]-

LFPIFENVNPEY) were synthesized as a fusion peptide to the C terminus of the antennapedia

(RQIKIWFQNRRMKWKK) internalization sequence with > 91% pure (110). Scrambled-siRNA (Sc-

siRNA), human (h)-specific siRNA against STIM1 (ON-TARGETplus SMARTpool siRNA containing sequences: GGUGGUGUCUAUCGUUAUU, UACAGUGGCUGAUCACAUA,

CAAUUACCAUGACCCAACA, UCUCUUGACUCGCCAUAAU), and mouse (m)-specific siRNA against

Pyk2 (ON-TARGETplus SMARTpool siRNA containing sequences: GACACUACCUGGAACGAAA,

UCAUGGAACUGUAUCCUUA, UAAGGGCUCUCUCAUCAUG, GAAGUUGGCUCAGCAGAAC) were

purchased from Dharmacon (Lafayette, CO). Antibodies against VE-cadherin, VE-PTP, β-catenin,

CAMMβ, siRNA specific to human Pyk2, and transfection reagents were obtained from Santa Cruz

Biotechnology (Santa Cruz, CA). Anti-STIM1 pAb, Anti-HA tag was from Proteintech Group (Chicago,

IL). Anti-phospho-Pyk2 (Y-402) pAb, and anti-Pyk2 pAb, Anti-phospho-TAK1(T-187), Anti-phospho-

AMPKα (T-172), Anti-phospho-p38 MAPK, Anti-TAK1, Anti-AMPK, Anti-p38α MAPK, Anti-p38β MAPK,

Anti-p38γ MAPK, Anti-phospho-GSK-3β (Ser-9), Anti-GSK-3β, Anti-Ubiquitin, were purchased from Cell

Signaling (Danvers, MA). Anti-β-actin mAb, Pyk2 inhibitor (PF-431396 hydrate, 1 µM working solution),

TAK1 inhibitor (OZ; 5Z-7-Oxozezenol) was from Sigma (St. Louis, MO). Anti-phospho-tyrosine (P-Y20) mAb, anti-phospho (Y731)-VE-cadherin was from EMD Millipore (Billerica, MA). Anti-phospho (Y685)-

VE-cadherin antibody was from Abcam (Cambridge, MA). We obtained HA-tagged WT-TAK1, TAK1∆N

(constitutively active TAK1), TAK1 K63W (dominant negative mutant TAK1) expression constructs from

Dr. Jun-Ninomiya-Tsuji, NC State University, Raleigh, NC.

3.2 Animals

Generation of endothelial-cell restricted STIM1 knockout (STIM1∆EC) mice

Stim1 floxed (Stim1fl/fl) mice (111) were a gift from Dr. Masatsugu Oh-Hora (Tokyo Medical and Dental

University, Tokyo, Japan). C57BL/6 (WT) and B6.Cg-Tg(Cdh5-Cre)7Mlia/J (VE-cadh Cre+) mice were

obtained from Jackson Laboratories. Stim1fl/fl mice were crossed with VE-cadh Cre+ (VE-cadherin promoter driving the expression of Cre) mice to generate endothelial-restricted Stim1 knockout (Stim1ΔEC)

mice as described previously (79).

Generation of inducible endothelial-cell restricted TAK1 knockout (TAK1i∆EC) mice

MAP3K7fl/fl mice were crossed with Endothelial-SCL-Cre-ERT transgenic mice. Endothelial-SCL-Cre-

ERT is tamoxifen-inducible Cre-ER(T) recombinase driven by the 5' endothelial enhancer of the stem cell leukemia (SCL) locus. The resultant heterozygous TAK1fl/+ end-Cre and homozygous TAK1fl/fl end-Cre

mice are designated as TAK1fl/+Cre+ and TAK1fl/flCre+ respectively. Only after tamoxifen administration

TAK1 is deleted in TAK1fl/flCre+ mice and they are regarded as TAK1i∆EC mice. To determine the efficacy of the Cre-recombinase system and consequent TAK1 deletion in endothelial cells, TAK1fl/fl and

TAK1fl/flCre+ were administered tamoxifen (1 mg/mouse) i.p for 2 days. After a resting period of 5 days, lung endothelial cells (LECs) from these mice were isolated using PECAM-1 antibody and TAK1 expression was measured. There is complete loss of TAK1 expression in LECs of TAK1fl/flCre+ (TAK1i∆EC)

mice for 2 consecutive days of tamoxifen injection (at 8th day, LECs were isolated). Therefore, I will

perform experiments using the mice at the 8th day (2 consecutive days of tamoxifen injection).

3.3 Primary endothelial cell culture

Primary human lung microvascular endothelial cells (HLMVECs) were acquired from Lonza Walkersville,

Inc. (Walkersville, MD). They were cultured using EGM-2 MV medium supplemented with 10% FBS and

used between passages 2 to 4. “As per the guidelines set by University of Illinois Animal care committee, mice weighing 20 to 25 g were anesthetized using 3% halothane, and injected with heparin (50 U/mouse) through the jugular vein. Next, the abdominal cavity was opened and the pulmonary artery was cannulated. For blood removal, Krebs-Henseleit solution supplemented with bovine serum albumin (5 g/100 mL) was infused. Lungs were harvested and placed inside a culture hood. Lung tissue slices from atleast 3 mice were prepared, washed and suspended in HBSS. Excess HBSS was aspirated, and the tissue slices were minced and transferred to a 15-ml sterile tube. The minced tissues were suspended in

10 mL of collagenase A (1.0 mg/mL in HBSS) and digested for 60 min at 37˚C with gentle shaking. The released cells were centrifuged at 200g for 10 min. The pellet was suspended in 10 mL suspension buffer

(Ca2+ - and Mg2+ -free containing 0.5 g/100 mL bovine serum albumin, 2 mM EDTA, and 4.5 mg/ml D- glucose), and filtered through 200 μm mesh filter. The filtered cells were suspended in 10 mL of suspension buffer. To this cell suspension, 1.5 μg/mlL anti-mouse PECAM-1 antibody (BD Pharmingen)

was added and incubated at 4˚C for 30 min with gentle shaking. The cell suspension was centrifuged to remove unbound antibody and washed once with suspension buffer. The washed cells were then incubated with Dynabeads M-450 (Sheep anti-rat IgG) for 30 min at 4˚C. Following this, the cell

suspension was attached to a magnetic column and the unbound cells were aspirated. Cell bound with magnetic beads were washed once with HBSS and digested with trypsin for 3 min at 22˚C. The cells

released from the magnetic beads were separated, washed, suspended in growth medium (EGM-2

supplemented with 10% FBS). The cell suspension was plated on matrigel (BD Biosciences)-coated 35

mm culture dish and allowed to grow to confluence for 10 days. Cells were then harvested from the

matrigel plates with dispase (BD Bioscience) for 60 to 90 min. Cells were washed after dispase treatment

once with growth medium and plated on 0.1% gelatin coated culture dish. Cells passaged between 3 and

4 times were used in experiments. Mouse lung endothelial cells (MLECs) were characterized by their

cobblestone morphology, PECAM-1 (platelet/endothelial cell adhesion molecule-1) (or CD31)

expression, and Dil-Ac-LDL uptake.” (54)

3.4 Genotyping

To characterize genotype of the mice, tail snips from the mice were harvested in tubes and labeled

appropriately. The tails were then incubuted overnight at 55˚C with a cocktail of tail lysis buffer

(containing a solution of 120 μl of 0.5 M EDTA, 500 μl Nuclei lysis solution (Promega) and 17.5 μl of 20 mg/mL of Proteinase K). Following day, 200 μl of protein precipitation solution (Promega) was added to the mixture and the solution was vortexed at high speed for 20 seconds. Samples was then kept on ice for 5 min and centrifuged at 13000 g for 5 more min. The supernatant was collected in a new tube and

600 μl Isopropanol was to it, mixed by inversion. The solution was centrifuged at 13000 g for 5 min and to the pellet 600 μl of Ethanol was added. Samples were then finally centrifuged at 13000 g for 5 min and the supernatant was discarded without disturbing the pellet. The pellet was air-dried and 30 μl TE buffer was added to it. To suspend, it was incubated at 65˚C for 1 h.

3.5 Cell Culture

Human lung microvascular endothelial cells (HLMVECs) and Endothelial growth media-2 (EGM-2) were purchased from Lonza (Walkersville, MD). Mouse (C57BL/6) lung endothelial cells (mLECs) were isolated and cultured as described previously (54). HLMVECs were cultured in EGM-2MV supplemented with 10% FBS, and mLECs were grown in EGM-2 supplemented with 5% FBS. Both cell types were used between passages 3 and 6.

3.6 Immunoprecipitation

ECs grown to confluence treated with or without agonists were three times washed with phosphate- buffered saline (PBS) at 40C and lysed in lysis buffer (50 mM Tris-HCl, pH7.5, 150 mM NaCl, 1 mM

EGTA, 1% Triton X-100, 0.25% sodium deoxycholate, 0.1% SDS, 10 μM orthovanadate, and protease- inhibitor mixture as described (112). Mouse lungs were homogenized in lysis buffer (113). EC lysates or mouse lung homogenates were centrifuged (13,000 X g for 10 min) to remove insoluble materials.

Clear supernatant collected (300 µg protein) was subjected to immunoprecipitation. Each sample was

incubated overnight with 1 µg/ml of the indicated antibody at 4 °C. Next day, Protein A/G beads were

added to the sample and incubated for 1 h at 4 °C. Immunoprecipitates were then washed three times

with wash buffer (Tris-buffered saline containing 0.05% Triton X-100, 1 mM Na3VO4, 1 mM NaF, 2 µg/ml leupeptin, 2 µg/ml pepstatin A, 2 µg/ml aprotinin, and 44 µg/ml phenylmethylsulfonyl fluoride).

Immunoprecipitated proteins were used for immunoblotting.

3.7 Immunoblotting

EC lysates, lung tissue homogenates, or immunoprecipitated proteins were resolved by SDS-PAGE on a 4-15% gradient separating gel under reducing conditions and transferred to a Duralose membrane.

Membranes were blocked with 5% dry milk in TBST (10mM Tris-HCl pH7.5, 150 nM NaCl, and 0.05%

Tween-20) for 1 hr at RT. Membranes were then probed with the indicated primary antibody (diluted in blocking buffer) overnight at 4°C. Next, membranes were washed 3x and then incubated with appropriate

HRP-conjugated secondary antibody. Protein bands were detected by enhanced chemiluminescence.

3.8 Cytosolic Ca2+ measurement

2+ 2+ 2+ “The cytoplasmic Ca ([Ca ]i) in ECs was measured using the Ca -sensitive fluorescent dye Fura-2/AM.

Cells were grown to confluence on gelatin-coated glass coverslips and then washed 2x with HBSS and incubated for 2 h at 37°C in culture medium containing 1% FBS. Cells were washed once and loaded

with 3 µM FURA-2/AM for 30 min. After loading, cells were washed with HBSS and the coverslips were transferred on a perfusion chamber at 37°C and imaged using a semimotorized microscope (Axio

Observer D1; Carl Zeiss GmbH, Jena, Germany) equipped with an AxioCam HSm camera (Carl Zeiss) and a Fluar 40x oil immersion objective. Light was provided by the DG-4 wavelength switcher (Princeton

Scientific Instruments, Monmouth Junction, NJ). A dual excitation at 340 and 380 was used, and emission was collected at 520 nM. The AxioVision physiology software module was used to acquire images at 1

sec intervals, and the data were analyzed off-line. In each experiment, 20-30 cells were selected to

2+ measure change in [Ca ]i.”(76,83,114)

3.9 Transendothelial Electrical Resistance Measurement

Real time changes in transendothelial monolayer electrical resistance (TER) was measured to assess

endothelial barrier function (115). Confluent endothelial monolayer was serum starved for 2 h with 1%

FBS containing medium. Next, thrombin-induced real-time change in TER was measured. Data are

presented as resistance normalized to its starting value zero time.

3.10 Immunostaining

Confluent ECs grown on glass coverslips were subjected either to thrombin treatment or left untreated.

After treatment, cells were washed quickly with cold PBS and fixed with 2% PFA for 15 min at 4°C.

Following fixation, cells were permeabilized with 0.05% Triton X-100 for 1 min at 4°C. Next, cells were washed three times with PBS and then incubated with blocking buffer (PBS containing 5% horse serum and 1% BSA) for 1 h at RT. Cells were then incubated overnight with indicated primary antibody (in PBS containing 1% BSA) at 4°C. Next day cells were washed three times, and then incubated with specific

Alexa-fluor conjugated secondary antibody and DAPI for 1 h at 4°C. Finally, cells were washed three times and mounted on glass slides for viewing. Images were acquired with the Zeiss LSM 510 confocal

microscope.

3.11 Paraffin-Embedded Tissue section staining

Mice lungs were isolated and paraffin-embedded using routine procedures. Slides embedded with tissue

sections were then de-waxed/hydrated, followed by washing three times, 10 min each. Lung sections were outlined with liquid blocker pen followed by blocking for 4 h at room temperature [Blocking Buffer:

1X TBS + 0.02% tween + 5% serum +1% BSA]. After blocking, the sections were incubated with primary

antibody, overnight (at 4˚C) in the presence of 1X TBS +0.02% tween +1% BSA. The slides were washed three times, 10 min each and were further incubated with secondary antibody for 1 h. They were then washed two times for 5 min each and incubated with DAPI for 30min. Finally, the slides were washed three times and then loaded with oil and a coverslip. The slides were left to dry overnight and viewed

under the confocal microscope the following day.

3.12 siRNA transfection

ECs grown to 70-80% confluence on gelatin-coated culture dishes were transfected with target siRNAs or sc-siRNA as described. At 72 h after transfection, cells were used for following experiments.

3.13 In vivo siRNA delivery in mouse lungs

Cationic liposomes were prepared with dimethyldioctadecylammonium bromide and cholesterol (1:1) as described (113,116). The liposomes and siRNA (control-siRNA or mouse Pyk2-siRNA) were then mixed

(8 moles lipid:1 μg siRNA) and the mixture (1 μg siRNA per g body weight) was injected retro-orbitally into mice under anesthesia (2.5% isoflurane in room air). At 48 h after delivery, the mice were used for experiments.

3.14 Mouse Lung Capillary Filtration Coefficient Measurement

“Pulmonary capillary filtration coefficient (Kf,c) was measured to determine microvascular permeability to liquid as described previously (49,54). To measure PAR-1 induced changes in endothelial barrier

function, PAR-1 agonist peptide (30 µM) containing perfusion buffer was infused via a side-port at a rate

of 0.2 ml/min. Kf,c measurement were made at baseline and after 20 min exposure to PAR-1 agonist peptide. The values are expressed as the ratio of experimental-to-basal Kf,c values in the same lung

preparation.”(114)

3.15 Assessment of Mouse lung Microvessel permeability In Vivo

Mice were anesthetized (2.5% isoflurane in room air) for retro-orbital vein injections. Initially, mice were injected with 100 µl of EBA (20 mg/kg). After 15 min, mice received either saline (100 µl) or PAR-1 peptide 100 µl (1 mg/kg). At 45 min after EBA injection, mice were sacrificed and lungs were harvested.

The EBA concentration in lung tissue was measured as described previously (114).

3.16 In vitro deletion of TAK1 in LECs

LECs obtained from TAK1fl/fl mice were infected with Adeno-cre virus to delete TAK1 in LECs. After 48

hrs, cells were used for experiments. Similarly, LECs obtained from TAK1fl/fl mice and TAK1fl/fl.Cre+ mice

were grown to confluence and incubated with 2 µM tamoxifen for 72 h, then used for experiments.

3.17 Statistical Analysis

Western blot bands were quantified using NIH ImageJ software. ANOVA and Student’s t-test (two-tailed) were used to determine statistical significance with a p-value threshold set at <0.05.

4 RESULTS I

4.1 Hypothesis: STIM1-induced SOCE activates Pyk2 to mediate tyrosine

phosphorylation of VE-PTP and thus cause vascular leak through disassembly of

AJs

SOCE is required for tyrosine phosphorylation of VE-PTP and disassembly of VE- cadherin junctions

We investigated first whether SOCE in ECs induces tyrosine phosphorylation of VE-PTP, which in turn, promotes destabilization of VE-cad at endothelial AJs. Since ER-localized STIM1 is critical for activating SOCE, we suppressed STIM1 expression using siRNA in human lung microvascular ECs

(HLMVECs). STIM1 expression as well as thrombin-induced Ca2+ entry were substantially reduced in

STIM1-siRNA transfected cells as compared to that in cells transfected with control siRNA (Sc-siRNA) or untreated cells (Fig. 6A). We observed that thrombin-induced tyrosine phosphorylation of VE-PTP was

also blocked in STIM1 knock down HLMVECs (Fig. 6B) but not in control or Sc-siRNA-treated HLMVECs

(Fig. 6B), suggesting that PAR-1-induced SOCE is essential for tyrosine phosphorylation of VE-PTP in

ECs. To address the functional relevance of tyrosine phosphorylation of VE-PTP, we determined endothelial permeability responses by measuring transendothelial monolayer resistance (TER) in control and STIM1-siRNA transfected HLMVECs. In STIM1-siRNA treated HLMVECs, thrombin-induced decrease in TER was markedly reduced compared with control cells (Fig. 7A). We also measured VE- cadherin expression at endothelial AJs in control and STIM1 knock down cells by immunostaining. In Sc- siRNA transfected cells, we observed VE-cadherin depletion from endothelial AJs 30 min after thrombin challenge (Fig. 7B) and restoration of VE-cadherin expression at endothelial AJs to near control levels 3 h after thrombin challenge (Fig. 7B). Interestingly, in STIM1-siRNA transfected cells, thrombin challenge failed to deplete EC AJs of VE-cadherin (Fig. 7B) suggesting that SOCE-induced VE-PTP tyrosine

phosphorylation was essential for PAR-1-induced depletion of VE-cadherin from endothelial AJs.

kDa IB: STIM1 84

β-actin 38

0 mM Ca2+ 1.5 mM Ca2+ 1.4 Control 1.2 Sc-siRNA 1 0.8 Thr 0.6

340/380 nm ratio 0.4 0.2 STIM1-siRNA

0 0 200 400 Time (sec)

B * B IP: VE-PTP 8 *

PTP Control Sc-siRNA STIM1-siRNA - 6 VE Thr (min) 0 15 30 0 15 30 0 15 30 - kDa 4

IB: p-Tyr PTP/T 200 - 2 (arbitrary units) (arbitrary VE

- p VE-PTP 0 200 Thr (min) 0 15 30 0 15 30 0 15 30

Control Sc-siRNA STIM1-siRNA

Figure 6. STIM1 knockdown in HLMVECs prevents thrombin-induced SOCE, VE-PTP phosphorylation. A, HLMVECs transfected with 100 nM Sc-siRNA or STIM1-siRNA. 48 h after transfection, cells were used to determine STIM1 protein expression by immunoblot (IB) (top panel) or 2+ 2+ 2+ used to measure thrombin-induced ER-store Ca release and Ca release-activated Ca entry (bottom panel). B, Non-transfected HLMVECs, or HLMVECs transfected with Sc-siRNA or STIM1-siRNA were challenged with thrombin (25 nM). Cell lysate were immunoprecipitated with VE-PTP pAb and blotted with phosphotyrosine mAb. Quantified data from three experiments are presented as the mean ± SE ratio of phosphorylated to total protein. *, p < 0.05; transfected compared with non-transfected or Sc-siRNA- transfected cells.

14000 Thr Control Sc-siRNA (100 nM) 12000 ) Ω STIM1-siRNA (100 nM) + Thr 10000

8000

( Resistance 6000 Sc-siRNA (100 nM) + Thr

4000 Time (h) 0 0.5 1.0 1.5 2.0 2.5

DB Thr (min)

0 30 180

Control

120 Control

100 Sc-siRNA *** STIM1-siRNA 80 60 40 siRNA - cad cad at expression AJs) - Sc

20 VE RFI (normalized to RFI cellscontrol (normalized 0 Thr (min) 0 30 180

siRNA -

STIM1

Figure 7. STIM1 knockdown in HLMVECs prevents thrombin-induced permeability increase, and disassembly of VE-cadherin (VE-cad) at endothelial AJs. A, HLMVECs were transfected with Sc- siRNA or STIM1-siRNA. At 24 h after transfection, cells were plated on gold electrodes (see details under “Methods”). After 24 h, cells were washed and incubated in 1% FBS containing medium for 2 h, and then cells challenged with medium or thrombin (25 nM). Values from three experiments are presented as mean ± SE. B, HLMVECs transfected with 100 nM Sc-siRNA or STIM1-siRNA and then challenged with thrombin (25 nM) were stained with anti-VE-cad pAb. Experiment was repeated three times. The bar graph represents quantitative analysis of VE-cad staining at cell-cell junctions. Data shown are mean ± SE of relative fluorescent intensity (RFI) compared with untreated cells. n = 8-10 cells per group; ***, p < 0.001, transfected compared with non-transfected or Sc-siRNA-treated cells.

EC specific STIM1 deletion inhibits phosphorylation of VE-PTP and VE-cadherin and increases in lung vascular permeability

To further address the role of SOCE in inducing tyrosine phosphorylation of VE-PTP to

disassemble AJs, we generated endothelial cell-restricted STIM1 knockout (Stim1∆EC) mice (see details

in Methods). We observed a loss of STIM1 expression in lung ECs (LECs) obtained from Stim1∆EC mice

(Fig. 8A). Thrombin-induced Ca2+ entry (i.e., SOCE) was absent in LECs of Stim1∆EC mice (Fig. 8B).

Next, we measured PAR-1-induced tyrosine phosphorylation of VE-PTP in vivo in lungs of WT and

Stim1∆EC mice following i.v. injection of PAR-1 agonist peptide TFLLRNPNDK-NH2 into mice. At 20 min

after PAR-1 peptide injection, the lungs harvested from WT and Stim1∆EC mice were used to determine

tyrosine phosphorylation of VE-PTP. PAR-1 induced tyrosine phosphorylation of VE-PTP in lungs of WT

mice (Fig. 8C), whereas PAR-1-induced tyrosine phosphorylation of VE-PTP was significantly reduced

in lungs of Stim1∆EC mice (Fig. 8C). We also observed that PAR-1-induced phosphorylation of VE- cadherin at Y685 and Y731 was suppressed in lungs of Stim1∆EC mice (Fig. 8D), indicating that SOCE- dependent tyrosine phosphorylation of VE-PTP was responsible for VE-cadherin phosphorylation.

To address the in vivo role of STIM1-activated SOCE in mediating lung vascular leak through

disassembly of VE-cadherin at AJs, we measured PAR-1-induced lung microvascular liquid permeability

(Kf,c) in intact lungs. We observed that the PAR-1 agonist peptide induced a 3-fold increase in Kf,c over

fl/fl basal in Stim1 (WT) mice (Fig. 9A). PAR-1 agonist peptide however failed to significantly increase Kf,c

in Stim1∆EC mice (Fig. 9A). We next determined PAR-1-induced lung vascular leak in vivo by measuring uptake of Evans blue dye conjugated with albumin (EBA) into lungs of WT and Stim1∆EC mice. The PAR-

1 agonist peptide induced a 3-fold increase in lung EBA uptake over basal in WT mice (Fig. 9B) whereas

PAR-1 agonist peptide failed to induce EBA uptake in lungs of Stim1∆EC mice (Fig. 9B). These results

together demonstrate that STIM1-dependent VE-PTP tyrosine phosphorylation signaled VE-cadherin

phosphorylation, which in turn, destabilized VE-cadherin junctions.

A C C IP: VE-PTP

LT STIM1fl/fl STIM1∆EC LECs PAR-1-pep (min) kDa 0 0 20 20 0 0 20 20 kDa

IB: STIM1 84 IB: p-Tyr 200 38 β-actin VE- PTP 200

*** 2.5 0 mM Ca2+ 1.5 mM Ca2+ 10 B PTP 8

- 2 Stim1 fl/fl-LECs VE

- 6

1.5 4 PTP/T

- 2 (arbitrary units) (arbitrary

VE Thr -

1 p

340/380nm ratio 0 ∆EC Stim1 -LECs PAR-1-pep 0 20 0 20 (min) 0.5 STIM1fl/fl STIM1∆EC

0 Figure 8. PAR-1-induced Ca2+ entry, Time (sec) 0 200 400 600 phosphorylation of VE-PTP, and D phosphorylation of VE-cad are impaired in LT STIM1fl/fl STIM1∆EC ∆EC fl/fl Stim1 mice. A, LECs from Stim1 and PAR-1-pep (min) 0 0 20 20 0 0 20 20 kDa Stim1∆EC mice were used for immunoblot to 120

685 IB: p -VE-cad determine STIM1 expression. B, LECs from 120 p731-VE-cad Stim1fl/fl and Stim1∆EC mice were used to measure

120 VE-cad thrombin-induced ER-store Ca2+ release and Ca2+ 2+ β-actin 38 release-activated Ca entry. Data in A and B are

representative of three independent experiments. fl/fl ∆EC 7 * 5 * C, Stim1 and Stim1 mice were i.v. injected cad Cad - - 6 4 with PAR-1 agonist peptide (see details in VE VE 5 -

- 4 3 Methods). 0 and 20 min after administration of cad/T cad/T 3 - 2 - PAR-1 agonist peptide, lungs were harvested and

2 VE VE - (arbitrary units) (arbitrary -

(arbitrary units) (arbitrary 1

1 731

685 lung tissue extracts prepared for p p 0 0 PAR-1-pep 0 20 0 20 PAR-1-pep 0 20 0 20 immunoprecipitation (IP) with VE-PTP pAb and (min) (min) STIM1fl/fl STIM1∆EC STIM1fl/fl STIM1∆EC blotted with phosphotyrosine mAb. D, Lung tissue fl/fl ∆EC extracts from Stim1 and Stim1 mice were immunoblotted with the indicated phosphotyrosine specific VE-cad antibodies. Quantified results are presented as the ratio of

phosphorylated to total protein in C and D. n = 4 mice for each group. ***, p < 0.001; Stim1fl/fl versus Stim1∆EC.

** A FB *** 0.12 *** 35 ***

30 0.1 0/g) Lung)

2 25 0.08 µg/g 20 0.06 15

(ml/m/cm H 0.04 f,c 10 K

Evans blue ( 0.02 5

0 0 PAR - 1 - pep - + - + PAR-1-pep - + - +

fl/fl ∆EC fl/fl ∆EC STIM1 STIM1 STIM1 STIM1

Figure 9. PAR-1-induced increase in lung vascular permeability impaired in Stim1∆EC mice. A, Lungs from Stim1fl/fl and Stim1∆EC mice were isolated to determine lung vascular permeability (see details under “Methods”). PAR-1 agonist peptide (25 μM) was included in the perfusion buffer to determine PAR- 1-induced microvessel filtration coefficient (Kf,c). n = 4 mice per group; **, p < 0.01; ***, p < 0.001; control Stim1fl/fl versus PAR-1 peptide Stim1fl/fl or Stim1fl/fl PAR-1 peptide versus Stim1∆EC PAR-1 peptide. B, Stim1fl/fl and Stim1∆EC mice received PAR-1 agonist peptide intravenously and were subsequently used to measure EBA uptake in the lungs. Results are mean ± SE. n = 4 mice per group; ***, p < 0.001; control Stim1fl/fl versus PAR-1 peptide Stim1fl/fl or Stim1fl/fl PAR-1 peptide versus Stim1∆EC PAR-1 peptide.

SOCE-induced Pyk2 activation phosphorylates VE-PTP to induce dissociation of VE-PTP from VE-cadherin

Since Pyk2 is activated by Ca2+ signaling, we studied the possibility that thrombin-induced SOCE

activated Pyk2 in ECs. We measured thrombin-induced phosphorylation of Pyk2 at Y402 as a measure of Pyk2 activation (117,118). Thrombin induced phosphorylation of Pyk2 at Y402 in control and Sc-siRNA transfected HLMVECs (Fig. 10A). However, thrombin-induced Pyk2 phosphorylation at Y402 was blocked in STIM1-siRNA transfected HLMVECs (Fig. 10A). In other experiments, we measured PAR-1- induced Pyk2 activation in vivo in lungs of WT and Stim1∆EC mice as described above (Fig. 8). PAR-1 activation induced a 4-fold increase in phosphorylation of Pyk2 at Y402 over basal in lungs of WT mice

(Fig. 10B) whereas PAR-1-induced Pyk2 phosphorylation at Y402 was markedly reduced in Stim1∆EC

mice (Fig. 10B). We also observed that thrombin caused Pyk2 binding to VE-PTP in HLMVECs (Fig.

11A). Thus, SOCE signaling promoted Pyk2 activation and the association of activated Pyk2 to VE-PTP.

Next, we used the Pyk2 selective inhibitor PF431396 (36) to study effects of Pyk2 inhibition on

thrombin-induced VE-PTP tyrosine phosphorylation in HLMVECs. PF431396-treatment of ECs

prevented thrombin-induced tyrosine phosphorylation of VE-PTP as compared to control cells (Fig. 11B).

PF431396 treatment also prevented thrombin-induced phosphorylation of VE-cadherin at Y685 and Y731

(Fig. 11C). To address the relationship between tyrosine phosphorylation of VE-PTP and VE-PTP binding to VE-cadherin, we next performed immunoprecipitation. We observed thrombin as well as VEGF (known to induce dissociation of VE-PTP from VE-cadherin [48]) caused dissociation of VE-PTP from VE- cadherin in control cells (Fig. 12, left panel). This response was blocked in PF431396 treated cells, (Fig.

12, right panel), indicating that VE-PTP tyrosine phosphorylation is essential for the dissociation of VE-

PTP from VE-cadherin at the level of AJs.

* * A 12 * * A 10 Control Sc-siRNA STIM1-siRNA 8 Pyk2

Thr (min) - 0 15 30 0 15 30 0 15 30 kDa 6 IB: p-Pyk2 116 4 Pyk2/T

T-Pyk2 116 p (arbitrary units) (arbitrary 2

β-actin 38 0 Thr (min) 0 15 30 0 15 30 0 15 30 Control Sc-siRNA STIM1-siRNA BB 8 *

fl/fl ∆EC LT STIM1 STIM1 6

Pyk2 PAR-1-pep (min) - 0 0 20 20 0 0 20 20 kDa 4

IB: p-Pyk2 116 Pyk2/T - 2 p units) (arbitrary

T-Pyk2 116 0 β-actin 38 PAR -1-pep 0 20 0 20 (min) STIM1 fl/fl STIM1∆EC

, Figure 10. STIM1-dependent Pyk2 activation induces tyrosine phosphorylation of Pyk2 in ECs. A

HLMVECs transfected with Sc - siRNA or STIM1-siRNA as in Figure 6 were used to measure thrombin-

induced phosphorylation of Pyk2 at Y402 as a measure of Pyk2 activation. Quantified results are mean

± SE of three experiments and are presented as the ratio of phosphorylated to total protein. **, p < 0.01;

compared with control or Sc-siRNA treated cells. B, Stim1fl/fl and Stim1∆EC mice received PAR-1 agonist

peptide as described in Figure 8C. 0 and 20 min after administration of PAR-1 agonist peptide, the lungs were used for IB to determine tyrosine phosphorylation of Pyk2 at Y402. Quantified results are mean ±

SE and are shown as the ratio of phosphorylated to total protein. n = 4 mice for each group. *, p < 0.05; Stim1fl/fl versus Stim1∆EC.

D IP:VE-PTP CA IP: VE-PTP DB Vehicle PF431396 Thr (min) 0 15 30 60 kDa Thr (min) 0 15 0 15 kDa

IB: Pyk2 116 IB: p-Tyr 200

VE-PTP 200 VE-PTP 200

* * 10

PTP 15 -

* PTP * - * 8 VE - 10 6

5 PTP/T - (arbitrary units) (arbitrary (arbitrary units) (arbitrary 2 VE 0 - p Pyk2 bindingto VE Thr (min) 0 15 30 60 0 Thr (min) 0 15 0 15

Vehicle PF431396

EEC Vehicle PF431396 Figure 11. STIM1-dependent Pyk2 kDa Thr (min) 0 15 0 15 activation induces tyrosine 120 IB: p685-VE-cad phosphorylation of VE-PTP and VE-cad

120 in ECs. A, HLMVECs exposed to VE-cad thrombin were immunoprecipitated (IP-

120 p731-VE-cad ed) with VE-PTP pAb and blotted with anti-Pyk2 pAb. Quantified results are 120 VE-cad mean ± SE of three experiments. *, p < 0.05; control versus thrombin treated. B, 16 16 HLMVECs were pretreated with

cad * * * cad - -

* PF431396 (1 μM) or vehicle (DMSO) for VE VE

- 12

- 12 15 min and then challenged with thrombin 8 8 cad/T cad/T (25 nM). After thrombin treatment, cell - - VE VE lysates were IP-ed with anti-VE-PTP pAb - 4 - 4 (arbitrary units) (arbitrary (arbitrary units) (arbitrary 685 731 and immunoblotted with a phospho- p p 0 0 tyrosine specific mAb. The blot was re- Thr (min) 0 15 0 15 Thr (min) 0 15 0 15 probed with anti-VE-PTP pAb. C, Vehicle PF431396 Vehicle PF431396 HLMVECs challenged with thrombin as in B were immunoblotted with the indicated phospho-tyrosine specific VE-cad antibodies. Quantified results are mean ± SE of three experiments presented as the ratio of phosphorylated to total protein in B and C. *, p < 0.05; **, p < 0.01; ***, p < 0.001; control (vehicle) versus PF431396.

IP: VE-PTP IP: VE-PTP Thr VEGF Thr (min) 0 15 30 0 15 30

Time (min) 0 15 30 60 0 15 30 60 kDa PF431396 - - - + + + kDa 120 120 IB: VE-cad IB: VE-cad

VE-PTP VE-PTP 200 200

2.5

PTP * - 2.5 2 PTP

- * 2

1.5 1.5 1 1 0.5 (arbitrary units) (arbitrary

cad binding to VE 0.5 - 0 units) (arbitrary

cad binding to VE - VE Time 0 15 30 60 0 15 30 60 0

(min) VE Thr 0 15 30 0 15 30 Thr VEGF (min) Vehicle PF431396

Figure 12. Thrombin as well as VEGF promotes VE-cad dissociation from VE-PTP in ECs. HLMVECs challenged with thrombin (25 nM) or VEGF A (50 ng/ml) were IP-ed with anti-VE-PTP pAb and immunoblotted with anti-VE-cad pAb (left panel). Quantified results are mean ± SE of three experiments. *, p < 0.05; **, p < 0.01; control versus thrombin- or VEGF-treated. HLMVECs were pretreated with PF431396 (1 μM) or vehicle (DMSO) for 30 min and then exposed to thrombin; cells were IP-ed with anti-VE-PTP pAb and immunoblotted with anti-VE-cad pAb (right panel). Quantified results from three experiments are shown. *, p < 0.05; control versus PF431396 treated.

To address the role of Pyk2 in regulating AJ assembly, we suppressed Pyk2 expression with siRNA in lung ECs (LECs) obtained from wild type mice and studied PAR-1 responses. In Pyk2-siRNA transfected LECs, Pyk2 protein expression was markedly reduced (Fig. 13A, top panel). We measured

PAR-1-induced SOCE in control and Pyk2-siRNA transfected cells. Pyk2 knockdown had no significant effect on SOCE activated by thrombin (Fig. 13A, bottom panel). Next, we determined VE-PTP phosphorylation and VE-PTP binding to VE-cadherin in control and Pyk2-siRNA transfected cells.

Thrombin-induced tyrosine phosphorylation of VE-PTP and VE-PTP dissociation from VE-cadherin were prevented in Pyk2 knockdown LECs (Fig. 13B, and C), indicating that Pyk2 signaling downstream of

SOCE phosphorylated VE-PTP which in turn promoted the dissociation of VE-PTP from VE-cadherin.

In vivo silencing of Pyk2 in mouse lung microvascular ECs prevents VE-PTP tyrosine phosphorylation, VE-cadherin phosphorylation, and increased endothelial permeability

To address the in vivo role of Pyk2 in regulating endothelial barrier integrity secondary to VE-PTP tyrosine phosphorylation, we silenced Pyk2 expression in mouse lung microvascular ECs through the use of siRNA by liposome-mediated delivery, which targets lung ECs (113,116). At 48 h after siRNA delivery, Pyk2 protein expression was markedly reduced in murine lungs (Fig. 14A). Since both VE-PTP and VE-cadherin are exclusively expressed in ECs, we studied the effects of Pyk2 silencing on PAR-1 induced tyrosine phosphorylation of VE-PTP and VE-cadherin. PAR-1 agonist peptide administration resulted in tyrosine phosphorylation of VE-PTP and phosphorylation of VE-cadherin at Y685 and Y731 in

lungs of mice receiving control si-RNA (Fig. 14, B and C). In contrast, PAR-1 agonist peptide-induced

tyrosine phosphorylation of VE-PTP and VE-cadherin phosphorylation at Y685 and Y731 was suppressed in lungs of mice receiving Pyk2-siRNA (Fig. 14, B and C). PAR-1-induced lung vascular leak as assessed by EBA uptake was also prevented in lungs of Pyk2 silenced mice (Fig. 15). Thus, Pyk2 induced vascular leak through phosphorylation of VE-PTP and VE-cadherin in ECs.

HB IP:VE-PTP Control Sc-siRNA Pyk2-siRNA kDa GA Thr (min) 0 15 30 0 15 30 0 15 30 kDa

IB: Pyk2 116 IB: p-Tyr 200 β-actin 38

2+ VE-PTP 0 mM Ca 2+ 200 1 1.5 mM Ca * 0.8 Sc -siRNA * 6 Pyk2-siRNA PTP 0.6 - 5

VE Thr - 4 3 0.4

PTP/T

- 2

340/380 nm ratio

units) (arbitrary

VE 1

0.2 -

p 0 Thr (min) 0 15 30 0 15 30 0 15 30 0 Time (sec) 0 200 400 600 Control Sc - siRNA Pyk2 -siRNA

IP: VE-PTP C Figure 13. SOCE-induced Pyk2 activation

promotes tyrosine phosphorylation of VE-PTP and

dissociation from VE-cad. A, LECs from wild type Thr (min) 0 15 0 15 0 15 kDa (C57BL/6) mice transfected with 100 nM Sc-siRNA or

120 IB: VE-cad Pyk2-siRNA were immunoblotted with anti-Pyk2 pAb to

120 determine Pyk2 expression (top panel) or cells were

VE-PTP used to measure thrombin-induced ER-store Ca2+ * release and Ca2+ release-activated Ca2+ entry (bottom 1.2

PTP * - 1 panel). Values are mean ± SD. n = 15 cells per group. 0.8 B, control LECs, Sc-siRNA transfected LECs, and 0.6 Pyk2-siRNA transfected LECs were exposed to

thrombin for 0, 15, and 30 min. After thrombin 0.4 (arbitrary units) (arbitrary

cad binding to VE 0.2 treatment, cell lysates were IP-ed with anti-VE-PTP -

VE 0 pAb and immunoblotted with anti-phospho-tyrosine Thr (min) 0 15 0 15 0 15 mAb. The blot was re-probed with anti-VE-PTP pAb. Control Sc-siRNA Pyk2-siRNA Quantified data from three experiments are presented as the ratio of phosphorylated to total protein. *, p < 0.05; significantly different from control or Sc-siRNA transfected. C, control LECs, Sc-siRNA transfected LECs, and Pyk2-siRNA transfected LECs were exposed to thrombin for 0, and 15 min. After thrombin treatment, cell lysates were IP-ed with anti-VE- PTP pAb and immunoblotted with anti-VE-cad pAb. The blot was re-probed with anti-VE-PTP pAb. Quantified results from three experiments are shown. *, p < 0.05; significantly different from control or Sc-siRNA transfected.

B A Sc-siRNA Pyk2-siRNA IP:VE-PTP

1 2 3 1 2 3 kDa Sc-siRNA Pyk2-siRNA

kDa PAR-1-pep (min) 0 0 20 20 0 0 20 20 IB: Pyk2 116 IB: p-Tyr 200

β-actin 38 VE-PTP 200

4 *** *** 1.2 *** PTP

- 3 1

actin 0.8

- 2 β 0.6

PTP/T 0.4 - 1 Pyk2/ units) (arbitrary VE 0.2 - (arbitrary units) (arbitrary p 0 0 Sc-siRNA Pyk2-siRNA PAR-1-pep 0 20 0 20 (min) Sc-siRNA Pyk2-siRNA C Sc-siRNA Pyk2-siRNA Figure 14. In vivo silencing of Pyk2 in PAR-1-pep (min) kDa 0 0 20 20 0 0 20 20

685 mouse lung microvascular ECs 120

IB: p -VE-cad

abrogates PAR-1-induced p731-VE -cad 120

phosphorylation of VE-PTP and VE- VE-cad 120 cad. C57BL/6 mice injected with Sc-

siRNA or Pyk2-siRNA were used for *** 4 ** 3 *** experiments (see details under cad cad

- ** - “Materials and Methods”). A,

Immunoblot analysis of the lungs from 2

mice 48 h after injection of Sc-siRNA or

cad/Total VE cad/Total

cad/Total VE cad/Total Pyk2-siRNA to determine Pyk2 protein - - 1

(arbitrary units) (arbitrary VE (arbitrary units) (arbitrary

1 VE expression. Quantified data are - - 685 731

p presented in arbitrary units. n = 8-10 p 0 0 mice per group. ***, p < 0.001, Sc- PAR-1-pep 0 20 0 20 PAR-1-pep 0 20 0 20 (min) (min) siRNA versus Pyk2-siRNA. B, Mice Sc-siRNA Pyk2-siRNA Sc-siRNA Pyk2-siRNA injected with Sc-siRNA or Pyk2-siRNA were i.v. injected with PAR-1 agonist peptide (see details under “Materials and Methods”) for 0 or 20 min. The lungs harvested after PAR-1 agonist peptide injection were used for lung extracts preparation and then immunoprecipitated with anti-VE-PTP pAb. The immunoprecipitated samples were immunoblotted with anti-phosphotyrosine mAb. Blots were re-probed with anti-VE-PTP pAb. Quantified results are presented as the ratio of phosphorylated to total protein. n = 4-6 mice in each group. ***, p < 0.001; Sc- siRNA versus Pyk2-siRNA. C, Lung extracts from the above experiment were used for immunoblot analysis with anti-phospho-Y685-VE-cad or anti-phospho-Y731-VE-cad pAb. Quantified results are presented as the ratio of phosphorylated to total protein. n = 4-6 mice in each group. **, p < 0.01; ***, p < 0.001; Sc-siRNA versus Pyk2-siRNA.

35 *** *** 30

Lung) 25

µg/g

15 10

Evans blue ( 5

0 PAR-1-pep 0 30 0 30 (min) Sc-siRNA Pyk2-siRNA

Figure 15. In vivo silencing of Pyk2 in mouse lung microvascular ECs abrogates PAR-1-induced lung vascular leak. Mice injected with Sc-siRNA or Pyk2-siRNA were i.v. injected with PAR-1 agonist peptide (see details under “Materials and Methods”) to measure EBA uptake in the lungs. Results are mean ± S.E. of changes in lung EBA uptake. n = 4 mice per group; ***, p < 0.001; Sc-siRNA control versus Sc-siRNA PAR-1 pep or Sc-siRNA PAR-1 pep versus Pyk2-siRNA PAR-1 pep.

VE-PTP C-terminal phospho-Y1981 binds to and activates Src to increase vascular permeability

Our results above showed that Pyk2 signaling was essential for PAR-1-induced tyrosine phosphorylation of VE-PTP’s cytosolic region and VE-cadherin phosphorylation. However, the link between Pyk2-mediated tyrosine phosphorylation of VE-PTP and VE-cadherin phosphorylation was not established. VE-PTP contains the amino acid motif “1981YENV1984” in the C-terminus suggesting that VE-

PTP’s C-terminal Y1981 could be the target of Pyk2. Thus, we investigated whether VE-PTP’s C-terminal phospho-Y1981 binds the SH2-domain of Src, which in turn, activates Src to phosphorylate its substrate

VE-cadherin. To address this question, we first determined whether Src activation is required for VE- cadherin phosphorylation in response to thrombin. We observed that inhibition of Src with PP1 blocked thrombin-induced phosphorylation of VE-cadherin at Y685 and Y731 (Fig. 16A). Next, we determined whether PAR-1-induced Pyk2 activation promotes Src activation. Thrombin-induced Src phosphorylation at Y416 was indicative of Src activation in ECs (Fig. 16B). In Pyk2 knockdown ECs, thrombin-induced Src

phosphorylation was prevented (Fig. 16B). Next, we determined whether STIM1-dependent Pyk2

activation induced Src binding to VE-PTP’s C-terminal phospho-tyrosine in vivo (see above, Fig. 4). In

control mouse lungs, PAR-1 agonist peptide induced Src binding to VE-PTP (Fig. 16C), whereas Src

binding to VE-PTP was abolished in Pyk2-siRNA treated lungs (Fig. 16C). We also observed that Pyk2

inhibition prevented thrombin-induced binding of Src to VE-PTP (Fig. 16D).

To determine the relationship between Pyk2-mediated tyrosine phosphorylation of VE-PTP and

VE-PTP binding to and activation of Src, we used a synthetic cell-permeable peptide derived from VE-

PTP’s C-terminus (1977-LFPIYENVNPEY-1988), which contains the conserved amino acid motif “YENV”, to

block Src activation. It is noteworthy that the phospho-Y1981 of VE-PTP can bind to signaling molecules

containing the Src homology-2 (SH2) domain, including Src itself (33,34,37). We synthesized this sequence 1977LFPIYENVNPEY1988 (wild-type [WT]) as a

Sc-siRNA Pyk2-siRNA A Vehicle PP1 B Thr (min) 0 15 30 60 0 15 30 60 kDa Thr (min) 0 15 30 0 15 30 kDa

p-Src (Y416) 60 685 120

IB: p -VE-cad Src 120 60 p731-VE-cad Pyk2 120 116 VE-cad β-actin 38

16 ** *** 7 7 ** cad cad

- *** - 6

12 Src

5 - 5

units) 4 8 4

3 /Total cad/Total VE cad/Total cad/Total VE cad/Total - - 3 Src

2 - (arbitrary units) (arbitrary 4 units) (arbitrary VE VE - - 2 416 (arbitrary (arbitrary

1 p 685 731 p p 1 0 0 Thr (min) 0 15 30 0 15 30 Thr (min) 0 15 30 0 15 30 0 Thr (min) 0 15 30 60 0 15 30 60 Vehicle PP1 Vehicle PP1 Sc-siRNA Pyk2-siRNA

CC IP:VE-PTP DD IP:VE-PTP

Sc-siRNA Pyk2-siRNA Vehicle PF431396

PAR-1-pep (min) 0 20 60 0 20 60 Thr (min) kDa 0 15 30 60 0 15 30 60 kDa

IB: Src 60 IB: Src 60

VE-PTP VE-PTP 200 200

** * 3.5 ** 7 * 3

PTP 6 - PTP

2.5 -

2 4

1.5 3 1 2 binding to VE (arbitrary units) (arbitrary binding to VE (arbitrary units) (arbitrary 1

Src 0.5 Src 0 0 Thr (min) 0 20 60 0 20 60 Thr (min) 0 15 30 60 0 15 30 60 Sc-siRNA Pyk2-siRNA Vehicle PF431396 Figure 16. SOCE-induced Pyk2 activation triggers VE-PTP mediated Src phosphorylation in ECs. A, Confluent HLMVECs pretreated with Src inhibitor PP1 (10 μM) for 30 min were exposed to thrombin for different time intervals. Then cells were used to determine VE-cad phosphorylation at Y685 or Y731. Quantified results shown are from three experiments as the ratio of phosphorylated to total protein. **, p < 0.01; ***, p < 0.001; control versus PP1 treated. B, HLMVECs transfected with Sc-siRNA or Pyk2- siRNA were treated with thrombin and immunoblotted with an antibody specific to phospho-Y416-Src. Quantified results shown are from three experiments as the ratio of phosphorylated to total protein. **, p < 0.01; Sc-siRNA versus Pyk2-siRNA. C, C57BL/6 mice injected with Sc-siRNA or Pyk2-siRNA (see above in Fig. 4) were i.v. injected with PAR-1 agonist peptide for 0, 20, and 60 min. Lung extracts prepared after PAR-1 agonist peptide treatment were IP-ed with anti-VE-PTP pAb. The IP-ed samples were immunoblotted with anti-Src mAb. Quantified results shown are from three experiments. **, p < 0.01; Sc-siRNA versus Pyk2-siRNA. D, HLMVECs treated with Pyk2 inhibitor followed by thrombin exposure were immunoprecipitated with anti-VE-PTP pAb. The IP-ed samples were immunoblotted with anti-Src mAb. Quantified results shown are from three separate experiments. *, p < 0.05; vehicle versus PF431396.

fusion peptide to the C-terminus of the antennapedia (RQIKIWFQNRRMKWKK) cell-permeable sequence (110). We also designed a cell permeable mutant 1981Y/F peptide (1977LFPIFENVNPEY1988),

which served as a negative control. The WT-VE-PTP peptide but not the mutant (Mut)-VE-PTP peptide

blocked PAR-1-induced Src activation in ECs (Fig. 17A). Consistent with this observation, WT-VE-PTP

peptide prevented PAR-1-induced tyrosine phosphorylation of VE-PTP (Fig. 17B). These results

collectively demonstrate that Pyk2 induced phosphorylation of Y1981 in VE-PTP’s C-terminal region and

that phospho-Y1981 in turn bound to and activated Src to phosphorylate VE-cadherin.

We next investigated whether treatment with the cell permeable WT-VE-PTP peptide could inhibit the PAR-1-induced permeability response. We pretreated confluent EC monolayers with either WT-VE-

PTP or Mut-VE-PTP peptide and then exposed them to thrombin for different time intervals. After thrombin exposure, cells were used to measure the expression of VE-cadherin at cell-cell junctions by staining with an anti-VE-cadherin pAb (Fig. 18A). Thrombin addition caused profound disassembly of

VE-cadherin at cell-cell junctions in control and Mut-VE-PTP peptide treated cells at 15 min and 30 min

(Fig. 18A), which were restored to basal level 3 h after thrombin stimulation (Fig. 18A). In WT-VE-PTP

peptide treated ECs, thrombin failed to disassemble VE-cadherin at cell-cell junctions (Fig. 18A). We

observed by immunoprecipitation that WT-VE-PTP peptide pretreatment prevented thrombin-induced

dissociation of VE-PTP from VE-cadherin (Fig. 18B). In line with these results, pretreatment of the

monolayer with WT-VE-PTP but not Mut-VE-PTP peptide inhibited the thrombin-induced decrease in

TER (Fig. 19A), indicating that WT-VE-PTP peptide prevented thrombin-induced permeability increase.

We also measured the effects of these peptides on PAR-1-induced lung vascular leak in mice by

measuring EBA uptake. WT-VE-PTP peptide administration blocked PAR-1-induced lung vascular leak

in mice (Fig. 19B). Based on these key findings, we propose a model (Fig. 20) in which phosphorylation

of VE-PTP at Y1981 in the cytosolic C-terminal region via the STIM1/SOCE-Pyk2 axis regulate binding and activation of Src. Subsequently, Src induces VE-cadherin phosphorylation to disassemble AJs and increase endothelial permeability.

A 1977LFPIYENVNPEY1988 (WT-VE-PTP pep) 1977LFPIFENVNPEY1988 (Mut-VE-PTP pep)

1981

Antennnapedia (AP): RQIKIWFQNRRMKWKK

** 7 * BE

units) 6 Src 0 0 1 2.5 5 1 2.5 5 (μM) - 5 Thr (min) 0 30 30 30 30 30 30 30 kDa 4

/Total 3 IB: p-Src (Y416) Src 60 - 2 416

p Src 60 1

(at 30 min, arbitrary 0 Conc. (µM) 0 2.5 5 2.5 5

IP: VE-PTP

FC 10 units)

PTP 8 - Conc. (μM) - - 2.5 5 2.5 5 **

VE - 6 * Thr (min) 0 30 30 30 30 30 kDa 4 PTP/T IB: p-Tyr 200 -

VE 2 - p VE-PTP 200 (at 30 min, arbitrary 0 Conc. (µM) 0 2.5 5 2.5 5

Figure 17. VE-PTP’s C-terminal tyrosine phosphorylation triggers Src activation in ECs. A, Schematic design for VE-PTP B, Confluent HLMVECs were incubated for 60 min with the indicated concentrations of either cell permeable WT-VE-PTP or Mut-VE-PTP peptide and then exposed to thrombin for 30 min. Cell lysates were then immunoblotted with an antibody specific to phos-Y416-Src. Results are mean ± SE of three experiments quantified as the ratio of phosphorylated to total protein. *, p < 0.05; **, p < 0.01; control (vehicle-treated) versus WT-VE-PTP peptide treated. C, HLMVECs, pretreated with WT-VE-PTP or Mut-VE-PTP peptide and then exposed to thrombin, were IP-ed with anti- VE-PTP pAb. The IP-ed samples were immunoblotted with anti-phosphotyrosine mAb. Results are mean ± SE of three experiments quantified as the ratio of phosphorylated to total protein. *, p < 0.05; **, p < 0.01; control (vehicle-treated) versus WT-VE-PTP peptide treated.

A Thr (min) 0 15 30 60 180

Vehicle Vehicle 1.6 WT-VE-PTP Pep 1.4 Mut-VE-PTP Pep

1.2 ** **

1.0 ** 0.8 PTP Pep Pep PTP - 0.6 VE - 0.4

RFI (normalized to control) RFI (normalized 0.2 WT

0.0 Thr (min) 0 15 30 60 120 PTP Pep Pep PTP - VE -

Mut

IP: VE-PTP 1.2 B *** B PTP - units) 1 0.8 0.6 *** Conc. (μM) - - 2.5 5 2.5 5 0.4 Thr (min) 0 30 30 30 30 30 kDa

120 cad binding to VE 0.2 IB: VE-cad - (at 30 min, arbitrary VE 0 Conc. (µM) 0 2.5 5 2.5 5 VE-PTP 200

Figure 18. Peptide derived from the Pyk2 phosphorylation site on VE-PTP prevents PAR-1- induced disassembly of VE-cad at endothelial AJs. A, Confluent HLMVECs pretreated with vehicle

(DMSO), WT-VE-PTP peptide (5 μM), or mut -VE -PTP (5 μM) peptide for 60 min and then challenged with thrombin for different time intervals were immunostained with anti-VE-cad pAb. The bar graph represents quantitative analysis of VE-cad staining at cell-cell junctions. Data shown are mean ± SE relative fluorescent intensity (RFI) of compared with untreated control cells. n = 6-8 cells per group. **, p

< 0.01; compared with vehicle treated or Mut-VE-PTP peptide treated cells. B, Confluent HLMVECs pretreated with either 5 μM WT-VE-PTP or mut-VE-PTP peptide for 60 min were exposed to thrombin for 30 min. After thrombin treatment, cell lysates were IP-ed with anti-VE-PTP pAb and immunoblotted with anti-VE-cad pAb. Results are mean ± SE of three experiments quantified. ***, p < 0.001; control (vehicle - treated) versus WT-VE-PTP peptide treated.

Vehicle alone 45 ** A Vehicle + Thr B 40 *** WT-VE-PTP Pep 2.5 µM + Thr

WT-VE-PTP Pep 5.0 µM + Thr 35

Mut-VE-PTP Pep 2.5 µM + Thr 30 Mut-VE-PTP Pep 5.0 µM + Thr 14000 Thr 25 20

) 12000 15 Ω

Evans blue (µg/GLung) 10 10000 5

Resistance ( Resistance 0 8000 PAR-1-pep - + + + (min) 6000 Time (h) 0 1 2 3

Figure 19. Peptide derived from the Pyk2 phosphorylation site on VE-PTP prevents PAR-1- induced permeability increase. A, HLMVECs on gold electrodes (see details under “Methods”) were washed and incubated in medium containing 1% FBS for 2 h and the cells were incubated with the indicated concentrations of WT-VE-PTP or mut-VE-PTP peptide for 60 min. Then the cells were challenged with thrombin (25 nM). Values represent mean ± SD of three experiments. B, C57BL/6 mice weighing 25 g were i. v. injected with vehicle (100 μl), WT-VE-PTP peptide (100 μl of 0.1 mM per mouse), or mut-VE-PTP peptide (100 μl of 0.1 mM per mouse). 60 min after peptide injection, PAR-1 agonist peptide was injected intravenously to determine EBA uptake (see details under Methods). Results are mean ± SE changes in lung EBA uptake. n = 4 mice per group; **, p < 0.01; ***, p < 0.01; untreated versus PAR-1 pep treated, untreated versus Mut-VE-PTP pep + PAR-1 pep treated, or PAR-1 pep alone versus WT-VE-PTP pep + PAR-1 pep treated.

Ca2+ PAR-1 Thr SOCs

Src (inactive) Kinase STIM1 Ca2+ PLC-β SH3 SH2 P Y 527 IP P 3 IP3R VE-PTP P ER Pyk2 Intact // YENV EC AJs 1981 VE-cad

SH3 P VE-PTP P

// YENV Y Y 1981 Src (active) VE-cad Y527

VE-PTP P // YENV EC AJs Disassembly Permeability ↑↑ 1981 P P Y Y VE-cad 685 731

Figure 20. Model for induction of VE-PTP-dependent Src activation in ECs via PAR-1-SOCE-Pyk2 pathway. PAR-1-induced ER store Ca2+ depletion via phospholipase C (PLC)-inositol 1,4,5-triphosphate (IP3) activates SOCE, inducing auto-phosphorylation of Pyk2 at Y402 to activate Pyk2. The activated Pyk2 binds to and phosphorylates VE-PTP at Y1981 in the C-terminal region. The phospho-Y1981 of VE- PTP binds to the SH2 domain of Src (anchored to the membrane), which, in turn, promotes the auto- phosphorylation of Y416 in the kinase domain of Src to trigger Src activation. The activated Src can phosphorylate tyrosine residues in the cytosolic C-terminal region of VE-cad to promote endothelial barrier dysfunction.

5 DISCUSSION I

We previously showed that PAR-1-induced SOCE in ECs signaled increased vascular permeability by inducing endothelial cell retraction secondary to actin-stress fiber formation, and disassembly of AJs (54,119,120). An emerging concept is that VE-PTP plays a crucial role in mediating

VE-cadherin homotypic interaction at AJs, and thus has an essential role in regulating endothelial barrier

function. VE-PTP interacts with VE-cadherin at AJs, and maintains endothelial cell-cell contact in a

phosphatase-dependent manner (26,27,31,35,36). Here we addressed the mechanisms by which PAR-

1-induced SOCE regulated the activity of VE-PTP and hence mediated disassembly of AJs and increased endothelial permeability. We demonstrated that loss of STIM1-mediated Ca2+ entry prevented PAR-1-

induced increased endothelial permeability confirming our earlier findings (54,119,120). The novel point

in this study is that STIM1-activated Ca2+ entry was required for PAR-1-induced Pyk2 activation and Pyk2 activation in turn mediated tyrosine phosphorylation of VE-PTP’s C-terminal region. Importantly, tyrosine phosphorylation of VE-PTP’s C-terminal induced by Pyk2 facilitated by the binding and activation of Src tyrosine kinase and in turn phosphorylation of VE-cadherin at AJs to disassemble AJs and increase endothelial permeability. A cell permeable peptide derived from the Pyk2 phosphorylation site on VE-

PTP abrogated the SOCE-induced increase in endothelial permeability. These findings collectively support the model that while VE-PTP constitutively maintains endothelial barrier integrity through dephosphorylation of VE-cadherin at AJs, Pyk2-induced phosphorylation of VE-PTP enables VE-PTP to function as a scaffold for Src and facilitate Src activation, thereby dissembling AJs through the phosphorylation of VE-cadherin.

STIM1 is known to be required for PAR-1-induced Ca2+ entry, which occurs primarily via activation of SOCs in ECs (76,83,114,121). We showed that EC-restricted deletion of STIM1 (Stim1∆EC) in mice prevented PAR-1-induced increase in lung vascular permeability. This was the result of inhibition of VE- cadherin internalization from the AJs. A similar role of STIM1 has been shown for LPS-induced vascular permeability (79). The current paradigm is that Src-mediated tyrosine phosphorylation of VE-cadherin

and subsequent internalization of phosphorylated VE-cadherin increases endothelial permeability (26-

29). Since VE-PTP maintains the integrity of AJs by dephosphorylating VE-cadherin at Y658, Y685, and

Y731, we investigated whether the signaling cascade activated by SOCE disrupted VE-PTP/VE-cadherin interaction and was thus required for increasing endothelial permeability. We observed that STIM1 depletion in HLMVECs prevented PAR-1-induced tyrosine phosphorylation of VE-PTP as well as of VE- cadherin at Y-685 and Y-731. Since VE-PTP and VE-cadherin are exclusively expressed in ECs, we determined PAR-1-induced phosphorylation of VE-PTP and VE-cadherin in vivo in lungs of WT and

Stim1∆EC mice. As with results in HLMVECs, the PAR-1-induced tyrosine phosphorylation of VE-PTP

and VE-cadherin was seen in WT lungs but not in Stim1∆EC mouse lungs. These results thus show that

STIM1-activated SOCE is essential for tyrosine phosphorylation of both VE-PTP and VE-cadherin, and thereby for the mechanism of increased endothelial permeability.

Pyk2 is a non-receptor tyrosine kinase, which does not contain SH2 or SH3 domains, but has a central catalytic domain flanked by an N-terminal FERM domain (118). Ca2+ and calmodulin binding to the N-terminal FERM domain of Pk2 induce Pyk2 dimerization, which triggers autophosphorylation of

Pyk2 at Y402 required for its catalytic activity (117,122). Ca2+-dependent Pyk2 activation in monocytes

(123) and ECs (124) induced inflammation through NF-κB activation. Reactive oxygen species (ROS)

are also known to activate Pyk2 (36,118). van Buul et al. (125) showed that anti-VE-cadherin antibody

induced disassembly of AJs via the ROS-Pyk2 axis mediating β-catenin tyrosine phosphorylation.

Another study showed that TNF-α-induced VE-cadherin phosphorylation in HUVECs was activated by

Pyk2 via the p110α isoform of PI3K (126). Vockel and Vestweber reported that lymphocyte binding to

VCAM-1 on the murine brain EC surface or stimulation of murine ECs with VEGF induced the dissociation

of VE-PTP from VE-cadherin, which involved ROS-dependent Pyk2 activation (36). This study proposed

a model in which Pyk2 induced the binding of phosphorylated VE-PTP substrate to the cytosolic domain

of VE-PTP, thus inducing conformational changes across the membrane resulting in detachment of the

extracellular domain of VE-PTP from VE-cadherin.

Since Pyk2 can be activated by SOCE (117), we investigated whether PAR-1-induced SOCE

signaling was capable of phosphorylating Pyk2 at Y402, thereby activating Pyk2. First, on depleting

STIM1 in HLMVECs using siRNA, we observed that the PAR-1-induced Pyk2 activation was impaired in

STIM1 depleted HLMVECs. Next, using the Stim1∆EC mouse model, we showed that STIM-mediated

SOCE was essential for PAR-1-induced Pyk2 activation. Thus, it appears that PAR-1-induced SOCE signaling mediated Pyk2 activation in ECs. To address whether SOCE-dependent Pyk2 activation was in fact responsible for VE-PTP tyrosine phosphorylation, we inhibited Pyk2 by treating ECs with

PF431396. Pyk2 inhibition blocked thrombin-induced tyrosine phosphorylation of VE-PTP. Additionally, thrombin-induced tyrosine phosphorylation of VE-PTP was blocked in Pyk2-depleted ECs. To address the in vivo role of Pyk2, we silenced Pyk2 expression in mouse lung microvascular ECs through liposome- mediated delivery of siRNAs (113). Depletion of Pyk2 in mouse lung ECs in this manner abolished PAR-

1-induced tyrosine phosphorylation of VE-PTP and VE-cadherin, and increased lung vascular permeability. These results thus indicate the central role of the SOCE-Pyk2 axis in mediating VE-PTP and VE-cadherin phosphorylation, which in turn was responsible for increasing lung vascular permeability.

It is unclear whether Pyk2-induced tyrosine phosphorylation of VE-PTP can alter the phosphatase activity of VE-PTP and thus affect the phosphorylation level of VE-cadherin. This possibility has to be explored further to dissect the intricate mechanisms of VE-PTP regulated control of VE-cadherin AJs. In my study, I focused on how Pyk2-induced tyrosine phosphorylation of VE-PTP at Y1981 promotes Src activation and thus increased phosphorylation of VE-cadherin to disassemble the endothelial AJs.

It is now known that Src family tyrosine kinases induce VE-cadherin tyrosine phosphorylation to promote disassembly of endothelial AJs (26-29). We observed that either pharmacological inhibition of

Pyk2 or Pyk2 depletion in ECs prevented thrombin-induced Src activation, suggesting that Pyk2 is upstream of Src in the pathway mediating VE-cadherin tyrosine phosphorylation. In addition, we observed that thrombin-induced Src binding to VE-PTP and Src activation were dependent on Pyk2 activation.

However, whether Pyk2-mediated tyrosine phosphorylation of VE-PTP is essential for thrombin-induced

Src activation remains unknown. hVE-PTP’s cytosolic C-terminus contains the putative tyrosine residue

(Y1981), which can be phosphorylated by tyrosine kinases and bind and activate Src (33,34,37). Further, it was recently shown that in HEK293 cells expressing wild-type mVE-PTP, treatment with the tyrosine phosphatase inhibitor pervanadate (PV) induced VE-PTP tyrosine phosphorylation, which induced the

binding of the Src family kinase Fyn to VE-PTP (33). Interestingly, in HEK293 cells expressing the mutant

Y1982F–mVE-PTP, tyrosine phosphorylation of VE-PTP and Fyn binding to VE-PTP were abolished by

PV pretreatment, indicating that phosphorylation of Y-1982 in mVE-PTP’s C-terminus plays an essential

role in Src activation (33). To address whether Pyk2-mediated tyrosine phosphorylation of VE-PTP was

essential for Src activation downstream of PAR-1-induced SOCE, we used a synthetic cell-permeable peptide derived from hVE-PTP’s C-terminus (1977-LFPIYENVNPEY-1988) to block thrombin-induced Src activation. Here we observed that the wild-type peptide, but not the Y1981F mutant-VE-PTP (control) peptide, prevented thrombin-induced Src activation, VE-PTP tyrosine phosphorylation, VE-PTP dissociation from VE-cadherin, and increased vascular permeability. These results together show that

Pyk2-mediated Y1981-phosphorylation of VE-PTP induced VE-cadherin phosphorylation via Src activation to disassemble the AJs and increase endothelial permeability.

6 RESULTS II

Objective: SOCE-induced TAK1 activation in endothelial cells triggers the repair of the leaky

endothelial barrier

6.1 Hypothesis 1: TAK1 activation secondary to STIM1-mediated SOCE induces STIM1

phosphorylation which in turn terminates SOCE and thereby dampens the vascular

permeability response

Pharmacological inhibition of TAK1 augments Ca2+ entry as well as endothelial permeability

To study whether TAK1 kinase activity regulates PAR-1-induced Ca2+ entry, I measured thrombin- induced ER stored-Ca2+ release and Ca2+ release-activated Ca2+ entry (a.k.a. store operated Ca2+ entry;

SOCE) in HLMVECs in the presence and absence of TAK1 inhibitor. I observed that thrombin-induced

Ca2+ entry was dramatically augmented in TAK1-inhibitor treated cells when compared to the control untreated cells (Fig. 21A) indicating that TAK1 kinase activity plays a crucial role in regulating SOCE in

HLMVECs. Transendothelial electrical resistance (TER) of HLMVECs was determined as a measure of endothelial permeability (see methods). Rapid decrease in TER was observed in control cells upon thrombin challenge and this decrease in TER was recovered to the basal within 2 h (Fig. 21B). However, in TAK1-inhibitor treated cells decrease in TER was prolonged and did not recover after thrombin challenge. Thus, indicating that TAK1 activity in HLMVECs is crucial for regulating vascular endothelial permeability and its loss augments the permeability response as indicated by the prolonged decrease in

TER. To study whether TAK1 signaling modulates the re-annealing of AJs, HLMVECs were pretreated with and without OZ. After thrombin treatment, cells were immunostained with anti-VE-cadherin Ab.

Confocal analysis showed in control cells, thrombin-induced disruption of AJs was restored whereas in

OZ-treated cells, AJs were severely disrupted (Fig. 21C). To further support the in vitro observations, we

measured effect of TAK1 inhibition on PAR-1 induced increase in intact lung vascular permeability in

collaboration with Dr. Vogel (see methods). Here, I observed that augmented liquid permeability in the

lungs perfused with both TAK1 inhibitor and PAR-1 peptide (TFLLRNPNDK -NH 2) compared to control lung perfused with PAR-1 peptide alone (Fig. 22D).

Thr (min) 0 60 180 A 2+ 2+ C 0 mM Ca 2 mM Ca

3.5 3 DMSO 2.5 OZ 2 1.5 Thr control 1 340/380nm ratio 0.5 TAK1 inhibitor 0 0 200 400 600 Time (sec)

*** D HLMVECs 0.18 ** B Thr 1.2 0.16 )

O/g 0.14

2 Ω OZ 1.0 H 0.12 ** 0.1 0.8 0.08 Control 0.06 ml/min/cm ml/min/cm Resistance ( 0.6 0.04 Filtration Coefficient 0.02 0.4 OZ+T hr 0

0.2 Normalized

0.0 0 1.0 2.0 3.0 Time (h) Figure 21.TAK1 inhibition augments PAR-1-induced Ca2+entry and permeability. A, HLMVECs were incubated with DMSO (control) or TAK1 inhibitor OZ (1µM, 30 min). Following pretreatment, cells were used to measure thrombin (50 nM) induced store Ca2+-release and Ca2+-entry. Data are representative of three experiments. B, HLMVECs grown on gold-microelectrodes were pretreated with DMSO or TAK1 inhibitor OZ (1 μM, 30 min) and thrombin-induced decrease in TER was measured to assess endothelial permeability. C, HLMVECs pretreated with or without TAK1 inhibitor OZ (1 μM, 30 min) and exposed to different time intervals were immunostained with anti-VE-cadherin Ab and confocal images were obtained. D, Intact lungs were infused with Hanks buffer or TAK1 inhibitor OZ (1 μM, 30 min). After pretreatment perfused lungs were used to measure PAR-1 peptide (5 µM)-induced increase in capillary filtration coefficient (Kfc). N=4, in each group; **, p < 0.01; ***, p < 0.001

TAK1 is activated downstream of PAR-1-induced SOCE

HLMVECs pretreated with or without TAK1 inhibitor (OZ; 1 µM for 30 min) were challenged with thrombin

(50 nM). I observed that TAK1 inhibitor blocked thrombin-induced phosphorylation of TAK1 at T-187 (see

Fig. 22A), indicating TAK1 is activated downstream of PAR-1.

To further delineate the mechanism of TAK1 activation, we addressed the possible role of SOCE in

activating TAK1 in ECs. Here we used STIM1 knockdown approach in HLMVECs and EC-restricted

STIM1 knockout (Stim1∆EC) mice (79,111), since STIM1 is required for activating SOCE (74-76,83,114)

(see methods). We observed that STIM1 expression as well as thrombin-induced Ca2+ entry was

substantially reduced in STIM1-siRNA transfected cells compared with scrambled siRNA (Sc-siRNA) or control cells (see Fig. 6A). Further, I investigated the effect of STIM1 knockdown on thrombin-induced phosphorylation of TAK1. Interestingly, I observed in STIM1-siRNA treated ECs, thrombin-induced phosphorylation of TAK1 was markedly reduced (see Fig. 22B). Next, Lung ECs (LECs) isolated from

Stim1fl/fl and Stim1∆EC mice were used to determine STIM1 protein expression by immunoblot showed

absence of STIM1 expression in LECs of Stim1∆EC mice (see Fig. 8A). Thrombin-induced SOCE was

absent in LECs of Stim1∆EC mice (see Fig. 8B). As expected, TAK1 phosphorylation at Thr-187 in response to thrombin was prevented in LECs of Stim1∆EC mice (see Fig. 22C).

Characterization of inducible EC-specific TAK1 knockout mice

Since global as well EC-restricted TAK1 knockout mice are embryonically lethal (90,127); I generated

tamoxifen-inducible EC-restricted TAK1 knockout (TAK1i∆EC) mice by crossing loxP-flanked Map3k7

(Map3k7fl/fl) mice (86) with inducible Cre-ER(T) driven by 5’endothelial enhancer of the stem cell leukemia

(Fig. 23A) (128,129). The genotypes of resultant mice (see methods) were confirmed using PCR (Fig.

23B). At 8 weeks, littermates of TAK1fl/fl and TAK1fl/fl.Cre+ mice were injected with tamoxifen (1 mg/mouse)

for 2 days to generate EC-restricted TAK1 knockout (TAK1i∆EC) mice (Fig. 23C). To verify EC-TAK1

deletion, LECs were isolated from TAK1fl/fl and TAK1i∆EC mice (at 8th day) using anti-PECAM-1 mAb and

TAK1 protein expression was measured (Fig. 23D). As expected, complete loss of TAK1 expression was

observed in LECs from TAK1i∆EC vs TAK1fl/fl mice (Fig. 23D). Ablation of TAK1 in TAK1i∆EC mice was

further confirmed by immunofluorescence analysis, wherein TAK1 and VE-cadherin double staining was

performed in TAK1fl/fl and TAK1i∆EC mice (Fig. 23E). In addition, H&E staining was also performed in lung

tissues from TAK1fl/fl and TAK1iΔEC mice which showed increased hemorrage as a result of loss of EC-

TAK1 (Fig. 23F). This suggests that TAK1 is crucial for regulation of the endothelium.

A C IB Vehicle OZ (TAK1 inhibitor) IB: Thrombin (min) 0 10 15 30 60 0 10 15 30 60 Thr (min) 0 15 0 15 T187 Phos -TAK1 PhosT187-TAK1

Total TAK1 Total-TAK1 β-actin

3.00 * B 2.50 Control TAK1 IB Control sc-siRNA STIM1-siRNA - 2.00 sc - siRNA Thr (min) 0 15 30 0 15 30 0 15 30 1.50 PhosT187-TAK1 1.00 TAK1/Total

Total-TAK1 units)(arbitrary - 0.50

p STIM1-siRNA 0.00 Thr (min) 0 15 30

Figure 22. PAR-1-STIM1-SOCE axis activates TAK1 in ECs. A, Thrombin induced TAK1- phosphorylation was measured. HLMVECs pretreated with DMSO or TAK1 inhibitor OZ (1µM, 30 min) were exposed to thrombin (50 nM) for different time intervals. Cell-lysates were used for immunoblot (IB)

with anti -phospho-T187-TAK1 and anti-TAK1 antibodies. B, Control HLMVECs, HLMVECs transfected

with sc-siRNA or STIM1-siRNA were used to measure thrombin-induced phosphorylation of TAK1 as in A. Representative blot is shown in top panel and quantified results of 3 experiments are shown in bottom panel. **p<0.001. C, LECs from Stim1fl/fl and Stim1∆EC mice were used to measure thrombin-induced phosphorylation of TAK1 by IB.

TAK1fl/fl A B TAK1fl/fl End-Cre 1 2 F 0 ♀ x ♂ Functional loxP loxP gene fl/WT F1 TAK1 End.Cre + Cre 2 Non- functional

loxP gene TAK1fl/fl x TAK1fl/WT End.Cre ♀ ♂

end-SCL-Cre-ERT fl/fl + F2 TAK1 .Cre

SCL endothelial enhancer SV Cre-ERT/pA

+ Tamoxifen

SCL endothelial enhancer SV Cre-ERT/pA Fl 550 bp WT 436 bp

End. Cre 500 bp Xi et al. PNAS 2006 C D TAK1fl/fl.Cre+ Days 1 2 3 4 5 6 7 LECs kDa Tamoxifen IB: T AK 1 (1 mg/mouse) 80

β-Actin 38 TAK1iΔEC Experiment E TAK1 VE-cad Merge F AW AW TAK1fl/fl TAK1i∆EC

BV BV TAK1fl/fl

AW AW

TAK1i∆EC BV BV

Figure 23. Generation of TAK1iΔEC mice. A, Mice carrying a Map3k7 (TAK1) gene in which exon 1 flanked by 2 loxP sites were bred with End-SCL-Cre-ER(T) mice containing tamoxifen-inducible Cre- ER(T) driven by 5'-endothelial enhancer of the stem cell leukemia (SCL) locus (which drives Cre expression specifically in endothelium). B, Shows breeding scheme for generating TAK1fl/fl-Cre+ mice. Standard PCR of tail DNA from indicated mice. C, TAK1fl/fl and TAK1fl/fl-Cre+ mice were injected with tamoxifen (1 mg/mouse, i.p. daily) for two days. Five days after tamoxifen treatment, mice were used for experiments. D, Lung endothelial cells (LECs) isolated from TAK1fl/fl and TAK1iΔEC mice were used for IB revealed the loss of TAK1 expression in LECs from TAK1iΔEC mice but not in wild type (TAK1fl/fl) mice. E, Lung sections from TAK1fl/fl and TAK1iΔEC mice were stained with anti-TAK1 (red) Ab, anti-VE-cadherin (EC marker) (green) Ab, and DAPI (blue) shows absence of TAK1 in blood vessel (BV) of TAK1iΔEC mice. AW, air way. F. H&E staining for Lung tissues from TAK1fl/fl and TAK1iΔEC mice. Arrows indicate blood vessels with hemorrage.

Loss of Endothelial-TAK1 augments PAR1-induced lung vascular leak and polymicrobial sepsis induced death

To determine whether EC-expressed TAK1 regulates lung vascular barrier integrity, we measured basal

as well as PAR-1 agonist peptide (TFLLRNPNDK)-induced lung vascular leak in vivo assessed by EBA

uptake. We observed that PAR-1-induced EBA uptake was highly augmented in Tmice compared with

WT (TAK1fl/fl) littermates, at 30 min after injection (Fig. 24A). This suggests that EC-expressed TAK1 signaling plays a critical role in restoring endothelial barrier integrity. Since, thrombin is produced in large quantities in septic conditions I used cecal ligation and puncture (CLP) as a severe model of sepsis to induce polymicrobial sepsis in age-, sex-, and weight-matched TAK1fl/fl and TAK1i∆EC mice. In this study,

we noted 85% mortality in TAK1i∆EC mice within 72 hrs of CLP, whereas only 28% of TAK1fl/fl mice died in the same period (Fig. 24B).

TAK1 deficiency augments PAR-1-induced Ca2+ entry in ECs

Next, we deleted TAK1 gene in TAK1fl/fl LECs by infecting LECs with Adeno-Cre virus. Here, LECs from

TAK1fl/fl mice were infected with recombinant Adenovirus expressing Cre (Adeno-Cre) for 48 days in culture. Adeno-Cre infection depleted TAK1 in LECs of TAK1fl/fl mice (Fig. 25A). Consistent with TAK1

inhibitor study, we observed in TAK1 deficient LECs, thrombin-induced SOCE was markedly enhanced as compared with control LECs (Fig. 25B), suggesting that EC-expressed TAK1 regulates endothelial

barrier integrity through modulation of Ca2+ influx.

A B

35 ** 100 TAK1fl/fl 30 80 TAK1fl/fl 25 TAK1iΔEC i∆EC 60 TAK1 20

15 40

10 Survival (%) 20

5 * EBA EBA uptake (µg/g Lung Tissue) 0 0 PAR-1 (min) 0 30 0 1 2 3 4 5 Time after CLP (days)

Figure 24. Persistent PAR-1-induced lung vascular leak and increased mortality in TAK1iΔEC mice. A, Basal and PAR-1-induced lung vascular leak was measured by Evans blue dye conjugated albumin (EBA) uptake. N=4, in each group, *p<0.05; **p<0.001. B, Polymicrobial sepsis was induced in TAK1fl/fl and TAK1iΔEC mice using cecal ligation puncture (CLP) method. Mild-CLP model was generated by punching five holes in cecum using a 21-gauge needle. Mice were observed for upto 5 days after CLP. n=10, in each group, *p<0.05

A TAK1fl/fl -LECs Adeno Cre (pfu/cell) 0 25 50

TAK1

β-Actin

B 2 mM Ca2+

4 0 mM Ca2+ 2 mM Ca2+ 3 3 2.5

fl/fl + fl/fl TAK1 Adeno Cre 2 TAK1 2 + Adeno Cre Thr fl/fl 1.5 TAK1 TAK1fl/fl

340/380nm ratio 1 Thr 340/380nm ratio 1

0 0.5 0 200 400 600 0 200 400 600 Time (sec) Time (sec)

Figure 25. TAK1 deficiency augments PAR-1-induced Ca2+ entry in ECs. A, Lung endothelial cells (LECs) from TAK1fl/fl mice were infected with adeno-Cre. Successful deletion of TAK1 (after 48 h) was assessed by IB. B, Control (TAK1fl/fl) LECs or TAK1fl/fl-LECs treated with adeno-Cre were used to measure thrombin-induced SOCE. Results shown are representative of 3 experiments.

TAK1 kinase activity is essential for terminating SOCE

To determine whether augmented Ca2+ entry is indeed a result of loss of TAK1 function, we performed

calcium measurement experiments using TAK1 expression constructs. We transfected human dermal

microvascular endothelial cells (HMECs) with HA-TAK1 (WT) or HA-TAK1 K63W (kinase-dead) plasmids

(see methods) and confirmed expression of constructs at 48 h by immunoblotting for HA expression in the cell lysates (Fig. 26A). Further, the control, HA-TAK1 or HA-TAK1 K63W plasmids transfected cells were used to measure thrombin induced SOCE. We observed highly augmented SOCE in HA-TAK1

K63W transfected cells as compared to the control or HA-TAK1 transfected cells (Fig. 26B), which suggests the TAK1 kinase functions is essential for inhibition of SOCE activity.

PAR-1-SOCE-CaMKKβ axis activates TAK1 in ECs

First, we showed that thrombin-induced phosphorylation of TAK1 at Thr-187 in HLMVECs (Fig. 22A).

Since, CaMKKβ is a Ca2+/calmodulin-dependent protein kinase kinase activated downstream of PAR-1

(130,131), we studied the effect of CaMKKβ inhibition on thrombin-induced phosphorylation of TAK1 in

ECs. CaMKKβ inhibitor STO-609 blocked thrombin-phosphorylation of TAK1 in ECs (Fig. 27A). This

experiment suggested that CaMKKβ could be acting upstream of TAK1 activation in thrombin response.

Therefore, we studied SOCE activity in response to CaMKKβ inhibititon. We observed that similar to

TAK1 inhibition, CaMKKβ inhibitor (STO-609) augmented thrombin-induced SOCE (Fig. 27B).

HMECs

IB HA

β actin

HMECs 0 mM Ca2+ 2 mM Ca2+ 1.2

HA-TAK1 K63W 1

0.8

Thr Control 0.6 ratio340/380nm 0.4 HA-T AK 1

0.2 0 1 2 3 4 5 6 7 T ime (min)

Figure 26. TAK1 kinase activity is essential for terminating SOCE. A, HMECs were transfected with either HA-TAK1 (WT-TAK1 construct) or HA-TAK1 K63W (Kinase-defective TAK1 mutant construct). Figure shows immunoblot analysis of HA-tag expression in HMECs cell lysates (harvested 48 hrs after transfection) B, Control and transfected cells (similar to A) were used to measure thrombin (50nM) induced store Ca2+-release and Ca2+-entry. Data are representative of three experiments.

B 0 mM Ca2+ 2 mM Ca2+ 1.2

A STO-609 1 DMSO STO-609 0.8 Thr (min) 0 15 30 0 15 30 p-TAK1 (Thr 187) 0.6 Total-TAK1 Control 340/380nm ratio340/380nm 0.4

0.2 0 2 4 6 8 10 T ime (min)

Figure 27. CaMKKβ signaling downstream of SOCE required for PAR-1 induced TAK1 activation and SOCE inhibition in ECs: A, HLMVECs pretreated with DMSO or CaMKKβ inhibitor STO-609 (1µM, 30 min) were exposed to thrombin (50 nM) for different time intervals. Cell-lysates were used for immunoblot (IB) with anti-phospho-T187-TAK1 and anti-TAK1 antibodies. B, HLMVECs as in A were used to measure thrombin (50 nM) induced ER-stored Ca2+ release and Ca2+ release-induced Ca2+ entry (SOCE). Data are representative of three experiments.

Intermediate role of TAK1 in mediating STIM1 phosphorylation in ECs

To dissect the role of TAK1 in mediating SOCE inhibition downstream of PAR-1-SOCE-CaMKKβ axis, we used TAK1 knockdown approaches in HLMVECs. Firstly, we transfected HLMVECs with either Sc- siRNA or TAK1 siRNA, and after 48 hours used the cell lysates to confirm reduced TAK1 expression by immunoblotting for TAK1 antibody (Fig. 28A) Then we also used these cells to study the effect of loss of

TAK1 on thrombin-induced phosphorylation of AMPKα1 (Fig. 28B). In conjunction with our previous

studies where we have shown that loss of CaMKKβ prevents AMPKα1 activation downstream of SOCE

(83), we found that silencing of TAK1 also prevented AMPKα1 activation on ECs.

Further we checked for thrombin-induced phosphorylation of human p38 isoforms (α, β, γ), where we

found only p38α and p38β to be phosphorylated in HLMVECs in response to thrombin (Fig. 29A). Next,

we challenged cells with thrombin for various time points and immunoprecipitated the cell lysates with

TAK1 antibody. These samples were then subjected to immunoblot analysis using antibody against p38α

and p38β. Interestingly, we found p38β to be predominantly interacting with TAK1 in response to thrombin

(Fig. 29B). We also found that silencing of TAK1 in ECs prevents thrombin-induced phosphorylation of

p38β in ECs (Fig. 29C).

Thus, on the basis of these studies, we examined the role of EC-TAK1 in SOCE induced phosphorylation

of STIM1. HLMVECs transfected with Sc-siRNA or TAK1-siRNA were exposed to thrombin and the cell

lysates were used to measure STIM1 phosphorylation by immunblot analysis. In support of our notion,

silencing of TAK1 in ECs abrogated SOCE-induced phosphorylation of STIM1 (Fig. 30). Therefore, these

studies suggest that STIM1-mediated Ca2+ entry in lung ECs activates CaMKKβ→TAK1→AMPKα→p38β pathway (Fig. 31), resulting in phosphorylation of STIM1, and inactivates Ca2+ entry and thereby the

endothelial permeability response.

2.5 A α 2 Total-TAK1 AMPK β-actin - 1.5 /T B α 1 Control sc-siRNA TAK1-siRNA AMPK - 0 15 30 0 15 30 0 15 30 units)(arbitrary 0.5 * T172 T172

Phos -AMPKα p Total-AMPKα1 0 Thr (min) 0 15 30 Control sc-siRNA TAK1 siRNA

Figure 28. TAK1 activation downstream of SOCE is required for AMPKα activation in ECs. A, HLMVECs transfected with either Sc-siRNA or TAK1 siRNA (100 nM). B, were used to measure thrombin-induced phosphorylation of AMPKα1 at Thr-172. Data are representative of three experiments. *p<0.05

A IP: phos-p38 B IP: TAK1 Thr (min) 0 15 Thr (min) 0 15 30 IB: p38α IB: p38α p38β p38 β p38γ TAK1

4 C 3.5 IP: phos-p38 3 sc-siRNA TAK1-siRNA β 2.5 Thr (min) 0 15 30 0 15 30 p38 - 2

SE: IB: p38β /T * β 1.5

IB: p38β p38 LE: - 1 p (arbitrary units)(arbitrary Total lysate 0.5 IB: p38β 0 Thr (min) 0 15 30 sc-siRNA TAK1 siRNA

Figure 29. SOCE-induced TAK1 activation triggers p38β MAPK signaling in ECs. A, HLMVECs challenged with or without Thrombin (50 nM) were harvested and immunoprecipitated using phospho- p38 antibody. The lysates were then probed with antibodies for p38 isoforms (α, β and γ). B, HLMVECs challenged with Thrombin (50 nM) for different time points, were harvested and immunoprecipitated using TAK1 antibody. The lysates were then probed with antibodies for p38 isoforms (α, β). C, HLMVECs transfected with either Sc-siRNA or TAK1 siRNA (100 nM) lysates were immunoprecipitated using phospho-p38 antibody. The lysates were then probed with antibodies for p38 β. (SE: short exposure; LE: long exposure). Data are representative of three experiments. *p<0.05

1.2

1.0

STIM1 0.8 sc-siRNA TAK1-siRNA - Thr (min) 0 15 30 0 15 30 0.6 IP: phos-Ser STIM1/T IB: STIM1 - 0.4 (arbitrary units)(arbitrary Total lysate * pSer 0.2 IB: STIM1 0.0 Thr (min) 0 15 30 sc-siRNA TAK1 siRNA

Figure 30. TAK1 activation downstream of SOCE is required for phosphorylation of STIM1. HLMVECs transfected with either Sc-siRNA or TAK1 siRNA (100 nM). After 48 hours, cells were challenged with thrombin and cell lysates were immunoprecipitated using phospho-Serine antibody. The lysates were then probed with antibody for STIM1. Data are representative of three experiments.

Thrombin

PAR-1

STIM1/SOCs

2+ Ca Vascular leak

CaMKKβ

TAK1

AMPK

P38β

Phos-STIM1

Figure 31. Schematic representation for SOCE-mediated TAK1 activation responsible for STIM1 phosphorylation and thus inhibition of SOCE. Thrombin activation of PAR-1 causes STIM1 oligomerisation and interaction with SOCs. This interaction is known to cause Ca2+ influx in ECs, which in turn promotes vascular leak. Here I show that TAK1 is the key molecule which acts downstream of SOCE-induced CaMKKβ to activate AMPK-p38β signaling pathway that is known to phosphorylate STIM1 and thus cause termination of SOCE. Thus, TAK1 is a crucial molecule to restrict PAR-1-induced vascular leak.

6.2 Hypothesis 2: TAK1 activation downstream of SOCE inactivates GSK-3β via p38

MAPK, which in turn enhances β-catenin expression at endothelial AJs and thus

restores vascular barrier function

Decreased expression of β-catenin and VE-cadherin induced by TAK1 deficiency in ECs

To investigate the role of TAK1 regulating the assembly of AJs, we measured β-catenin expression in

LECs of TAK1fl/fl and TAK1iΔEC mice. Expression of β-catenin was markedly reduced in LECs of TAK1iΔEC

mice as compared with WT (Fig. 32). Because an important function of β-catenin at AJs is to stabilize

VE-cadherin, we also measured VE-cadherin expression. VE-cadherin expression was markedly

reduced in TAK1iΔEC mice (Fig. 32). This study suggested that EC-expressed TAK1 controls endothelial barrier integrity through regulating the assembly of VE-cadherin junctions. In addition, since

GSK-3β regulates β-catenin expression

We next addressed the question of upstream regulation of β-catenin. We probed the possibility that GSK-

3β was involved since it is Ser/Thr kinase known to negatively regulate β-catenin (105,132-134). GSK-

3β-induced phosphorylation of β-catenin promoted ubiquitination and subsequent degradation of β-

catenin via proteasomal pathway (105,106,108,132-136). GSK-3β also has an important site that can be

inactivated by phosphorylation of its Ser-9 residue (106,108,135,136). Hence, we investigated the

possible role of GSK-3β in stabilizing β-catenin and determined whether it participated in stabilizing AJs.

Interestingly, PAR-1-induced SOCE is required for GSK-3β inactivation (phosphorylation at Ser-9) (Fig.

33). Here I showed STIM1 knockdown HLMVECs, GSK-3β inactivation was prevented in response

thrombin (Fig. 33). More importantly, TAK1 is upstream of GSK-3β inactivation pathway because loss of

TAK1 function in ECs blocked PAR-1-induced GSK-3β phosphorylation at Ser-9 (Fig. 34)

Further, we also studied role of TAK1 in thrombin-induced ubiquitination of β-catenin. Here, I silenced

TAK1 in HLMVECs (Fig. 34C) and after 72 hrs treated the cells with MG-132 (which is lysosomal inhibitor;

to prevent K-48 tagged protein degradation via lysosomal pathway). The cells were then challenged with

thrombin for specific time interval and then the cell lysates were used for immunoprecipitation with β-

catenin antibody. Immunoprecipitated samples were subjected to immunoblot analysis for Ubiquitin

antibody. We observed that loss of EC-TAK1 augmented thrombin-induced ubiqutination of β-catenin

which also suggests increased degradation of β-catenin. These results collectively support our

hypothesis SOCE-CaMKKβ-TAK1-p38 MAPK pathway can reassemble AJs through inhibiting GSK-3β

activity, and thus increasing the expression of β-catenin.

TAK1fl/fl TAK1iΔEC Tamoxifen (days) 0 0 2 2 2 2

β-catenin VE-Cadherin

β-actin

Figure 32. EC-restricted TAK1 deletion in mice downregulates AJs protein expression. Lung tissues harvested from TAK1fl/fl and TAK1i∆EC mice were homogenized and the lysates were subjected to IB analysis for β-catenin and VE-cadherin antibody. Data are representative of three experiments.

STIM1 β-actin HLMVECs

Control sc-siRNA STIM1-siRNA B Thr (min) 0 15 30 0 15 30 0 15 30 p-GSK-3β (Ser 9) Total-GSK-3β

β 3.00 3 - 2.50 Control GSK - 2.00 sc-siRNA 1.50 /Total β

3 1.00 -

(arbitrary units)(arbitrary 0.50 GSK

- STIM1-siRNA p 0.00 Thr (min) 0 15 30

Figure 33. STIM1 deletion blocks thrombin induced GSK-3β inactivation. A, HLMVECs transfected with either Sc-siRNA or STIM1 siRNA (100 nM). Immunoblot shows STIM1 expression after 48 h. B, After transfection, cells exposed to thrombin for different time intervals were used for immunoblot analysis to determine GSK-3β phosphorylation at Ser-9. Data are representative of three experiments.

A HLMVECs Total-TAK1 β-actin

Control sc-siRNA TAK1-siRNA B Thr (min) 0 15 30 0 15 30 0 15 30 p-GSK-3β (Ser 9)

Total-GSK-3β

β 2.50 2.00

GSK3 Control - 1.50 sc-siRNA

/Total

β 1.00

(arbitrary units)(arbitrary 0.50 GSK3 - TAK1-siRNA p 0.00 Thr (min) 0 15 30

Control sc-siRNA TAK1-siRNA C Thr (min) 0 30 60 0 30 60 0 30 60

IP: β-catenin IB: Ub

Total cell lysate IB: β-catenin

Figure 34 TAK1 deletion blocks thrombin-induced GSK-3β inactivation and augments β-catenin ubiquitination. A, HLMVECs transfected with either Sc-siRNA or TAK1 siRNA (100 nM). Immunoblot shows TAK1 expression after 48 h. B, After transfection, cells exposed to thrombin for different time intervals were used for immunoblot analysis to determine GSK-3β phosphorylation at Ser-9. C, Further, transfected cells were pretreated with MG-132 and then exposed to thrombin for indicated time intervals and the cells lysates were immunoprecipitated with β-catenin antibody. Following immunoprecipitation, the samples were immunoblotted against ubiquitin antibody. Data are representative of three experiments.

Inducible EC-restricted GSK-3β deletion promotes restoration of lung vascular integrity

As GSK-3β deletion in mice results in embryonic lethality (137,138), I generated EC-restricted tamoxifen- inducible GSK-3β knockout (GSK-3βi∆EC) in adult mice. Mice carrying a GSK-3β gene in which exon 2

flanked by 2 loxP sites (137) were bred with End-SCL-Cre-ER(T) mice containing tamoxifen-inducible

Cre-ER(T) driven by 5’ endothelial enhancer of the stem cell leukemia locus (128,129). At 8 wk,

littermates of GSK-3βfl/fl (WT) and GSK-3βfl/fl,Cre+ mice were treated with tamoxifen (1 mg/mouse; i.p.) daily for 2 days or 5 days to generate “wild type” and inducible EC-restricted GSK-3β knockout (GSK-3βi∆EC)

mice. After a gap of 5 days, lung tissues were harvested and GSK-3β expression was measured. GSK-

3β expression was remarkably suppressed in lung tissues of GSK-3βi∆EC vs. GSK-3βfl/fl mice (Fig. 35A).

In addition, I also observed that β-catenin expression was augmented in lung tissues of mice lacking

GSK-3β (Fig. 35A). To confirm deletion of GSK-3β in endothelial cells, LECs from GSK-3βi∆EC vs. GSK-

3βfl/fl mice were isolated at 7th day (two days tamoxifen, i.p.). As shown in (Fig. 35B, left panel), GSK-3β

expression was disrupted in LECs of GSK-3βi∆EC mice (residual GSK-3β in blots in Fig. 35B, due to contamination from non-ECs present in mouse lung EC preparations). Interestingly, I observed augmented expression of β-catenin in LECs of GSK-3βi∆EC mice compared to WT littermates (Fig. 35B, right panel), indicating a key role for GSK-3β in regulating β-catenin expression in ECs. Next, I determined whether augmented β-catenin expression enhanced the endothelial barrier. Here, I observed that PAR-

1- induced lung vascular leak was substantially reduced in lungs of GSK-3βi∆EC mice as compared with

WT (Fig. 35C).

A LT GSK3βfl/fl GSK3βiΔEC Tamoxifen (days) 2 2 5 5 2 2 5 5 kDa

IB: GSK3β 46

β-Cat 92

β-actin 38

LECs B

GSK-3β β-catenin β-actin β-actin

C 45 GSK-3βfl/fl 40 * GSK-3βiΔEC GSK-3βfl/fl GSK-3βiΔEC 35

30 25 20

15 10 5

EBA EBA uptake (µg/g Lung Tissue) 0 Basal PAR-1

Figure 35. Generation of tamoxifen-inducible EC-restricted GSK-3β knockout (GSK-3βi∆EC) mice. A, GSK-3βfl/fl and GSK-3βfl/fl were administered Tamoxifen (1 mg/mouse; i.p.) for the indicated number of days. After a gap of 5 days for each time point, lung tissues were harvested and were then subjected to IB analysis to measure expression of GSK-3β and β-catenin. B, As in A, GSK-3βfl/fl and GSK-3βfl/fl.Cre+ mice tamoxifen injected for 2 days and at 7th day LECs isolated were used for IB to determine GSK-3β (left panel) and β-catenin (right panel) expression. In each group, 3 mice were used for LECs isolation. C, PAR-1-induced EBA uptake in lungs of GSK-3βfl/fl and GSK-3βi∆EC mice was measured to assess lung vascular leak. One h after PAR-1 peptide injection, lungs harvested from mice were used for EBA uptake measurement. Left panel, shows image of lungs harvested from GSK-3βfl/fl and GSK-3βi∆EC mice after PAR-1 peptide injection. Right panel, shows quantified results. N= 4, in each group; **p<0.01.

7 DISCUSSION II

Ca2+ is crucial for maintenance of vascular endothelial barrier through regulation of endothelial cell- cell and matrix tethering and actin-myosin contractile machinery (54,55). Previous studies from our laboratory have shown that thrombin, an inflammatory pro-coagulant mediates lung vascular leak by activating PAR-1 on the EC surface (55). Further, studies from our laboratory showed that PAR-1-induced

Ca2+ entry in lung vascular ECs mediates lung vascular leak (50). In recent studies from our laboratory, it was shown that STIM1 is crucial for activation of SOCE in endothelial cells (76,121,139). However, the intricate mechanisms involved in termination of SOCE are still unclear.

In 2009, research conducted in Putney’s lab (81) suggested SOC function may be inhibited by

phosphorylation of STIM1 on its C-terminus residues. Based upon the deduced STIM1 sequence which

contains 10 putative phosphorylation sites for p38 MAPK; research from our has laboratory established

that SOCE activates CaMKKβ-AMPK-p38 MAPK signaling pathway (131) which in turn mediates phosphorylation of STIM1, and thus inhibition of SOCE. In the first part of my thesis, I showed how STIM1- mediated SOCE signaling disturbs the endothelial barrier. However, in the second part I focused on PAR-

1-induced restoration of vascular barrier integrity. Recent published studies in our laboratory, have established that SOCE induced by thrombin also activates CaMKKβ-AMPK-p38 MAPK signaling pathway in ECs. This study showed AMPKα1-p38β MAPK axis signaling induced phosphorylation of STIM1 to

“turn off” SOCE in ECs. In addition, we showed that ablation of AMPK and p38β MAPK enhances thrombin-induced SOCE and thereby augments permeability in ECs.

Although there is little evidence showing Ca2+-dependent activation of TAK1 (86,140), role of EC-

TAK1 downstream of PAR-1 still remains unknown. TAK1 interaction with AMPK and p38 MAPK is well known downstream of various signaling pathways. Therefore, I performed studies to understand the role of EC-TAK1 in mediating thrombin response. First, I showed that TAK1 function is crucial to reverse PAR-

1-induced lung vascular leak.

TAK1, a member of MAPK family, is well established as a crucial kinase in innate immune signaling pathways activated by various stimuli, including cytokines, B-cell, T-cell, and Toll-like receptor ligands.

TAK1 functions as a nodal kinase by phosphorylating IκB kinase, p38 MAPK, JNK, and ERK; thereby causing activation of NF-κB and MAPK signaling pathways. Global as well as endothelial cell-restricted

TAK1 knockout in mice showed embryonic lethality, indicating that TAK1 expression in ECs is essential for vascular development and endothelial barrier function. However, there is no study till date to address whether TAK1 signaling in endothelial cells regulate endothelial barrier integrity.

Because TAK1 is ubiquitously expressed in all tissues and its function differs in a cell-dependent manner (91), therefore I studied function of TAK1 downstream of PAR-1 in ECs. First, I observed PAR-

1-dependent phosphorylation of EC-TAK1 at Thr-187 residue, which has been shown to be essential for auto-phosphorylation and thus activation of TAK1(141). The phosphorylation of TAK1 at Thr-187 was blocked when the cells were treated with an irreversible inhibitor of TAK1. Thus, suggesting that TAK1 activation downstream of PAR-1 occurs in a time-dependent manner through auto-phosphorylation at

Thr-187 residue. To determine the role of PAR-1-induced SOCE in activation of TAK1 activity, I transfected HLMVECs with siRNA specific to STIM1 and measured TAK1 phosphorylation at Thr-187.

As shown in part I of the thesis, that STIM1 is essential for PAR-1-induced SOCE in endothelial cells. I observed that loss of endothelial STIM1 blocked PAR-1-induced phosphorylation of TAK1. This study was also performed using STIM1∆EC LECs, and similar effect was observed. Thereby, suggesting that

PAR-1-induced Ca2+ influx is essential for activation of TAK1 in ECs.

Lie at al. (142) have demonstrated that activated AMPK interacts with the scaffold protein TAB1

(which is an adaptor for TAK1), and the resulting complex associates with the p38 MAPK, thus promoting

auto-phosphorylation of p38 MAPK in ischemic hearts. Previous sudies from our laboratory have shown

that inhibition of endothelial-AMPK prevented thrombin-induced p38 MAPK, thus establishing that AMPK

lies upstream of p38 MAPK (131). In addition, p38 MAPK inhibition in endothelial cells suppressed

thrombin-induced STIM1 phosphorylation and enhanced thrombin-induced SOCE. Since, TAK1 is a

known upstream activator for both AMPK and p38 MAPK in multiple signaling pathways. Therefore, I studied the role of TAK1 in regulating SOCE. Similar to inhibition of p38 MAPK in ECs (83), I observed

TAK1 inhibition augmented SOCE in HLMVECs. Further, I measured the effect of TAK1 inhibition on endothelial permeability using transendothelial electrical resistance in HLMVECs. In support of our previous findings that augmented Ca2+-influx leads to hyperpermeability, I observed that TAK1 inhibition

in ECs augmented thrombin-induced decrease in TER, which did not recover to basal even after 12 h. In

addition, I performed staining of VE-cadherin at junctions in control and TAK1-inhibitor treated cells after

thrombin treatment, which revealed that TAK1 is essential for reannealing of VE-cadherin at AJs after

thrombin treatment.

Second, we showed that EC-TAK1 kinase function is essential for termination of SOCE activity, and

its loss also leads to increase in lung vascular permeability through destabilization of the endothelial AJs.

Second, I generated an adult mouse model of tamoxifen-inducible endothelial cell-restricted TAK1

knockout (TAK1i∆EC) mice to study role of TAK1 in regulating lung vascular barrier function. Third, I

showed TAK1 is essential for reversing PAR-1-induced lung vascular leak and preventing sepsis induced

mortality, this was found to be in conjunction with markedly reduced expression of VE-cadherin and β- catenin in TAK1i∆EC mice. Fourth, I showed that STIM1-mediated SOCE-CaMKKβ signaling is essential

for PAR-1-induced TAK1 activation in ECs. Fifth, I show that TAK1 is an upstream of AMPKα1-p38β

MAPK axis which induces phosphorylation of STIM1 to “turn off” SOCE in ECs. Sixth, I showed STIM1- mediated TAK1 activation is essential for thrombin-induced inactivation of GSK-3β. Finally, to identify

i∆EC role GSK-3β in regulating SOCE-induced disassembly of adherens junctions, I generated GSK-3β

i∆EC mice. In my studies using GSK-3β mice, I observed that loss endothelial GSK-3β augments β-catenin

expression and thus dampens PAR-1 induced vascular permeability response. Finally, these results

together show that TAK1-mediated inhibition of SOCE and inactivation of GSK-3β is essential to reverse

PAR-1-induced lung vascular leak (Fig. 36).

Figure 36. Proposed model for role of STIM1-mediated SOCE signaling in TAK1 activation which terminates SOCE and resolves lung vascular hyperpermeability. As shown in part-I of my thesis, thrombin, a pro-inflammatory mediator binds to PAR-1 on the EC surface which results in SOC-induced Ca2+ signaling to increase endothelial permeability via disassembly of the endothelial AJs components (VE-cadherin/β-catenin complex). Here I show supporting data to prove the hypothesis that TAK1 activation secondary to STIM1-mediated SOCE signaling induces STIM1 phosphorylation which in turn inhibits SOCE. This also results in reversal of signaling responsible for increased vascular permeability. Moreover, GSK-3β is known to mediate β-catenin phosphorylation, which promotes β-catenin degradation via proteasomal pathway to destabilize endothelial AJs. I show supporting data to prove the second hypothesis, that TAK1 activation secondary to STIM1-mediated Ca2+ entry inactivates GSK-3β via phosphorylation at Ser-9. Inactivation of GSK-3β thus promotes increased β-catenin expression at AJs which restores endothelial barrier integrity and thereby resolves pulmonary edema.

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9 VITAE

NAME: Dheeraj Soni

EDUCATION: • 2017- Ph.D., Cellular and Molecular Pharmacology, University of Illinois at Chicago, Chicago, Illinois • 2009- BS, Biotechnology, North Dakota State University, Fargo, North Dakota • 2008- Diploma, Biotechnology, Ansal Institute of Technology, Gurgaon, India

HONORS & AWARDS: • GSC Travel Award for attending IVBM (2016) meeting in Boston provided by Graduate Student Council at UIC • Student Presenter Award for presenting at IVBM (2016) meeting in Boston awarded by UIC- Graduate College • ASBMB Graduate Travel Award for Experimental Biology conference by American Society for Biochemistry and Molecular Biology (2016) • Chancellor’s student service and leadership award UIC (2016) • Special Recognition UIC Dept. of Pharmacology for developing and moderating Trainee-database website ($500 cash award) (2015) • Graduate Student Poster Prize Experimental Biology meeting in Boston awarded by American Physiological Society-Respiration Section (2015) • GSC Travel Award for attending Experimental Biology meeting in Boston awarded by Graduate Student Council at UIC (2015) • Chancellor’s student service and leadership award UIC (2015) • Pre-doctoral Education for Clinical and Translational Scientists (PECTS) Scholarship UIC-Center for Clinical and Translational Science 1-year scholarship ($27000) (2014) • Honorary Member, Phi Kappa Phi Academic Honor Society. Top 5% in class, inducted by NDSU Chapter (2009) • Honorary Member, Golden Key International Honor Society Top 15% in class, inducted by NDSU Chapter (2009) • Magna Cum Laude, BS Biotechnology, NDSU • Dean’s list (2 semesters) NDSU (2008-09) • Academic & Cultural Sharing scholarship, NDSU • Director's Scholarship, Each semester at AIT, India

• Dean's list (6 semesters) AIT, India (2006-2008)

PROFESSIONAL MEMBERSHIPs: • American Association for the Advancement of Science (AAAS) • American Society for Pharmacology and Experimental Therapeutics (ASPET) • American Society for Biochemistry and Molecular Biology (ASBMB) • American Physiological Society (APS) • American Heart Association (AHA) • North American Vascular Biology Organization (NAVBO)

ABSTRACTS:

• Soni, D., Billings, S., Vonnahme, K.A., Bilski, J.J., Caton, J.S., Redmer, D.A., Reynolds, L.P., and Grazul-Bilska, A.T. (2009) Effects of nutrient restriction and dietary selenium on expression of gap junctional protein connexin (Cx) 43 in fetal ovaries obtained from sheep in late pregnancy; implications for developmental programming. Proceedings of North Dakota Academy of Sciences 63, 22 • Soni, D., DebRoy, A., Wang, D. M., Vogel, S.M., Tiruppathi, C. (2015) TAK1 Signaling Downstream of PAR-1 in Endothelial Cells Restores Lung Vascular Barrier Integrity. FASEB J 29, 661.2 • Soni, D., DebRoy, A., Wang, D. M., Vogel, S.M., Tiruppathi, C. (2016) Ca2+-dependent Pyk2 Activation Destabilizes Endothelial Adherens Junctions by Disrupting Interaction between VE- PTP and VE-Cadherin. FASEB J 30,1138.1 • Soni, D., Tiruppathi, C. STIM1-dependent Pyk2 activation phosphorylates VE-PTP triggering Src induced VE-cadherin phosphorylation to increase the vascular permeability response to thrombin. NAVBO-IVBM 2016

PUBLICATIONS: • Grazul-Bilska, A. T., Vonnahme, K. A., Bilski, J. J., Borowczyk, E., Soni, D., Mikkelson, B., Johnson, M. L., Reynolds, L. P., Redmer, D. A., and Caton, J. S. (2011) Expression of gap junctional connexin proteins in ovine fetal ovaries: effects of maternal diet. Domest Anim Endocrinol 41, 185-194 • Tiruppathi, C., Soni, D., Wang, D. M., Xue, J., Singh, V., Thippegowda, P. B., Cheppudira, B. P., Mishra, R. K., DebRoy, A., Qian, Z., Bachmaier, K., Zhao, Y. Y., Christman, J. W., Vogel, S. M.,

Ma, A., and Malik, A. B. (2014) The transcription factor DREAM represses the deubiquitinase A20 and mediates inflammation. Nat Immunol 15, 239-247 • DebRoy, A., Vogel, S. M., Soni, D., Sundivakkam, P. C., Malik, A. B., and Tiruppathi, C. (2014) Cooperative signaling via transcription factors NF-kappaB and AP1/c-Fos mediates endothelial cell STIM1 expression and hyperpermeability in response to endotoxin. J Biol Chem 289, 24188- 24201 • Mittal M, Tiruppathi C, Nepal S, Zhao YY, Grzych D, Soni D, Prockop DJ, Malik AB. TNFα- stimulated gene-6 (TSG6) activates macrophage phenotype transition to prevent inflammatory lung injury. Proc. Natl. Acad. Sci. U.S.A 113 (50), E8151-E8158 (2016) • Ayee MAA, LeMaster E, Shentu TP, Singh DK, Barbera N, Soni D, Tiruppathi C, Subbaiah PV, Berdyshev, Bronova I, Cho M, Akpa BS, Levitan I. Molecular-scale biophysical modulation of an endothelial membrane by oxidized phospholipids. Biophysical Journal 112 (2), 325-338 (2017) • Soni, D., Regmi, S.C., Wang, D. M., DebRoy, A., Zhao, YY., Vogel, S.M., Malik, A.B., Tiruppathi, C. Pyk2 Phosphorylation of VE-PTP Downstream of STIM1 induced Ca2+ entry Regulates Disassembly of Adherens Junctions. AJP-Lung (2017)

MANUSCRIPTS UNDER REVIEW OR IN PREPARATION: • Mittal M, Nepal S, Hecquet CM, Soni D, Rehman J, Tiruppathi C, Malik AB. Neutrophil Activation of Endothelial Cell-Expressed TRPM2 mediates inflammatory cell migration and vascular injury. (under review) • Soni, D., Wang, D. M., DebRoy, A., Vogel, S.M., Tiruppathi, C. (2016) TAK1 signaling downstream of PAR-1 in endothelial cells restores lung vascular barrier integrity (in preparation)

10 APPENDIX

1 Permission letter for Figure 4