UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Lipopolysaccharide signaling in endothelial cells Dauphinee, Shauna Marie 2010

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Item Metadata

Download

Media
24-ubc_2010_spring_dauphinee_shauna.pdf [ 4.37MB ]
Metadata
JSON: 24-1.0069549.json
JSON-LD: 24-1.0069549-ld.json
RDF/XML (Pretty): 24-1.0069549-rdf.xml
RDF/JSON: 24-1.0069549-rdf.json
Turtle: 24-1.0069549-turtle.txt
N-Triples: 24-1.0069549-rdf-ntriples.txt
Original Record: 24-1.0069549-source.json
Full Text
24-1.0069549-fulltext.txt
Citation
24-1.0069549.ris

Full Text

    LIPOPOLYSACCHARIDE SIGNALING IN ENDOTHELIAL CELLS  by  Shauna Marie Dauphinee  B.Sc., Dalhousie University, 2001  M.Sc., Dalhousie University, 2004   A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF  THE REQUIREMENTS FOR THE DEGREE OF   DOCTOR OF PHILOSOPHY   in   The Faculty of Graduate Studies  (Experimental Medicine)    THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)   March 2010    © Shauna Marie Dauphinee, 2010   ii  ABSTRACT The endothelium plays a critical role in coordinating the innate immune response through the regulation of vascular tone, leukocyte recruitment and transmigration, and hemostasis. These functions are mediated, in part, by the signaling cascades initiated upon recognition of bacterial and viral products by a family of transmembrane receptors known as Toll-like receptors (TLRs). In endothelial cells, exposure to lipopolysaccharide (LPS), a major cell wall constituent of Gram-negative bacteria, results in endothelial activation through TLR4. Recruitment of the adapter protein, MyD88, to the receptor facilitates association of serine-threonine kinases of the IL-1 receptor associated kinase (IRAK) family. The IRAKs initiate a phosphorylation cascade through TNFR-associated factor 6 (TRAF6) culminating in activation of proinflammatory signaling pathways including NF-κB and c-Jun NH2-terminal kinase (JNK) pathways. This thesis investigates signaling molecules and pathways downstream of TLR4 in endothelial cells. Specifically, contained herein is a description of the role of heterotrimeric G proteins in endothelial TLR signaling. This thesis identifies for the first time the function of these proteins in multiple TLR signaling pathways. In addition, the work presented here describes the identification and characterization of a novel TLR4 signaling molecule, SAM and SH3 domain containing protein 1 (SASH1). SASH1 promotes LPS-induced NF-κB and JNK, by functioning as a scaffold molecule to bind TRAF6, transforming growth factor−β−activated kinase (TAK1) and IκB-kinase (IKK), thereby increasing proinflammatory cytokine production. The distinct functions of the endothelium in innate immunity highlight the need for an understanding of the signaling cascades initiated by LPS in endothelial cells and will be crucial to our understanding of the pathophysiology of sepsis in the clinic. iii  TABLE OF CONTENTS ABSTRACT .............................................................................................................................. ii  TABLE OF CONTENTS ......................................................................................................... iii  LIST OF TABLES ................................................................................................................. viii  LIST OF FIGURES ................................................................................................................. ix  LIST OF ABBREVIATIONS .................................................................................................. xi  ACKNOWLEDGEMENTS .................................................................................................. xvii   CHAPTER 1: INTRODUCTION ..............................................................................................1  1.1       THE IMMUNE SYSTEM .................................................................................2  1.2       THE INNATE IMMUNE SYSTEM .................................................................2  1.3       SEPSIS ...............................................................................................................4  1.4       ENDOTHELIAL CELLS ..................................................................................4  1.5       ENDOTHELIAL DYSFUNCTION IN SEPSIS ...............................................5  1.5.1       Vasoregulation ....................................................................................5  1.5.2       Vascular permeability .........................................................................5  1.5.3       Leukocyte recruitment and adhesion ..................................................8  1.5.4       Hemostasis ..........................................................................................9  1.6       LIPOPOLYSACCHARIDE .............................................................................10  1.7       TOLL-LIKE RECEPTORS .............................................................................12  1.8       MYD88-DEPENDENT TLR4 SIGNALING ..................................................18  1.8.1       Mitogen activated protein kinases ....................................................21  1.8.2       Phosphoinositide 3-kinases ...............................................................23  1.9       HETEROTRIMERIC G PROTEINS IN TLR4 SIGNALING ........................24  iv  1.10     MYD88-INDEPENDENT TLR4 SIGNALING ..............................................27  1.11     NEGATIVE REGULATION OF TLR4 SIGNALING ...................................28  1.11.1    Fas-associated death domain protein (FADD) ...................................29  1.11.2    A20 .....................................................................................................29  1.11.3    β-arrestin ............................................................................................31  1.11.4    Sterile α and Armadillo motif-containing molecule ..........................31  1.11.5    Additional endogenous regulators of TLR4 signaling .......................31  1.12     SCAFFOLD PROTEINS IN IMMUNE SIGNALING ...................................33  1.12.1    Pellino family .....................................................................................34  1.13     SAM AND SH3 DOMAIN CONTAINING PROTEIN 1 (SASH1) ...............34  1.14     AIM OF THE PRESENT STUDY ..................................................................37   CHAPTER 2: MATERIALS AND METHODS .....................................................................40  2.1        MATERIALS ..................................................................................................41  2.2        CELL CULTURE ...........................................................................................41  2.3        SASH1 ANTIBODY PURIFICATION ..........................................................42  2.4        RECOMBINANT PLASMIDS AND GENE TRANSFER ............................42  2.5        cAMP ASSAY ................................................................................................43  2.6        RNA ISOLATION ..........................................................................................43  2.7 REVERSE TRANSCRIPTION AND POLYMERASE CHAIN REACTION (RT-PCR) ...................................................................................44  2.8        PROTEIN ASSAY .........................................................................................46  2.9        IMMUNOBLOT ANALYSIS ........................................................................46  2.10      CO-IMMUNOPRECIPITATION ...................................................................47  v  2.11      TRAF6 UBIQUITINATION ..........................................................................47  2.12      LUCIFERASE REPORTER ASSAY .............................................................47  2.13      RNA INTERFERENCE .................................................................................48  2.14      LIPID RAFT ISOLATION .............................................................................50  2.15      MASS SPECTROMETRY ANALYSIS ........................................................50  2.16       IN SILICO ANALYSIS .................................................................................51  2.17       ELISA ............................................................................................................51  2.18       GENERATION OF SASH1 GENE-TARGETED MICE .............................52  2.19       β-GALACTOSIDASE STAINING ...............................................................52  2.20       ISOLATION OF MOUSE ENDOTHELIAL CELLS ...................................53  2.21       STATISTICAL ANALYSIS .........................................................................53   CHAPTER 3: HETEROTRIMERIC Gαi/o PROTEINS MODULATE ENDOTHELIAL TLR SIGNALING ............................................................................................................................54  3.1       INTRODUCTION ...........................................................................................55  3.2       RESULTS ........................................................................................................57  3.2.1     TLR ligands activate heteotrimeric Gαi/o proteins in endothelial  cells ....................................................................................................57  3.2.2  Gαi/o proteins contribute to LPS-induced activation of ERK1/2 and                  JNK in endothelial cells .....................................................................59  3.2.3     Pertussis toxin does not activate the MAPKs in endothelial cells ......59  3.2.4     Gαi/o proteins play a role in LPS-induced activation of Akt in                                       endothelial cells .................................................................................61  3.2.5 Gαi/o proteins do not contribute to LPS-induced activation of NF-κB                           in endothelial cells .............................................................................63  3.2.6 Gαi/o proteins do not function along a TRAF6-dependent pathway in                                       endothelial cells .................................................................................66 vi   3.2.7 Gαi/o proteins contribute to TLR2 and TLR3 signaling ....................68  3.2.8 PI3K is upstream of Akt and JNK in TLR4 signaling ........................72  3.2.9 TLR4 interacts with, and functions though, Gαi2 ..............................74  3.3 DISCUSSION ..................................................................................................76   CHAPTER 4: THE IDENTIFICATION OF SASH1 AS A NOVEL TLR4 SIGNALING MOLECULE ............................................................................................................................80  4.1 INTRODUCTION ...........................................................................................81  4.2       RESULTS ........................................................................................................83  4.2.1     Proteomic analysis identifies SASH1 as a putative player in the               TLR4 signaling pathway ....................................................................83  4.2.2     SASH1 positively regulates LPS signal transduction .........................85  4.2.3 SASH1 is not important for LPS-induced IFN signaling ...................88  4.2.4 SASH1 plays a role in TLR3, but not TLR2, signaling ......................88  4.2.5 SASH1 interacts with the C-terminal domain of TRAF6 ...................91  4.2.6 SASH1 regulates TRAF6 ubiquitination ............................................94  4.2.7 SASH1 does not interact with other TRAF molecules .......................97  4.2.8 SASH1 acts as a scaffold molecule through binding TAK1,  IKKα and IKKβ .................................................................................99  4.2.9 Generation of Sash1 gene-targeted mice ..........................................102  4.2.10 Sash1 expression in vivo ...................................................................102  4.3      DISCUSSION .................................................................................................108   CHAPTER 5: SUMMARY AND FUTURE PERSPECTIVES ............................................112  5.1      THE STUDY OF TLR SIGNALING .............................................................113  vii  5.2 HETEROTRIMERIC G PROTEINS IN ENDOTHELIAL TLR                        SIGNALING ...................................................................................................114  5.2.1 TLR4 as a single transmembrane spanning GPCR ...........................114  5.2.2 Is the LPS-induced Gαi/o-mediated signal dependent on                           CD14/TLR4?....................................................................................115  5.2.3 Cooperation between TLR4 and a GPCR .........................................116  5.2.4 Autocrine stimulation of a GPCR by an LPS-inducible product ......117  5.3 SASH1 AS A NOVEL TLR4 SIGNALING MOLECULE ..........................117  5.3.1 Scaffold molecules in innate immune signaling ...............................118  5.3.2 SASH1 in other TRAF6 signaling cascades .....................................119  5.3.3 SASH1 as a negative regulator of TLR4 signaling ...........................119  5.3.4 Future directions studying SASH1 in vivo........................................120  5.4       GENERAL SUMMARY AND FUTURE PERSPECTIVES ........................121   REFERENCES ......................................................................................................................125   APPENDIX I .........................................................................................................................142   APPENDIX II ........................................................................................................................169               viii   LIST OF TABLES Table 1.1    TLR ligands in human and mouse ........................................................................12 Table 1.2     Heterotrimeric Gα subunit effectors ....................................................................26 Table 2.1     List of primers ......................................................................................................45 Table 3.1     Densitometry analysis of PTx-sensitive Gαi/o endothelial TLR4 signaling .......63 Table 3.2     Densitometry analysis of PTx-sensitive Gαi/o endothelial TLR2 signaling .......71 Table 3.3     Densitometry analysis of PTx-sensitive Gαi/o endothelial TLR3 signaling .......72                                ix   LIST OF FIGURES Figure 1.1     Structure of LPS .................................................................................................11 Figure 1.2     Cellular localization of TLRs .............................................................................14 Figure 1.3     Structure of TLR4...............................................................................................16 Figure 1.4     TLR4 signaling ...................................................................................................17 Figure 1.5     TRAF family protein domain structure ..............................................................20 Figure 1.6     Gαi/o signaling ...................................................................................................25 Figure 1.7     Endogenous regulators of TLR4 signaling .........................................................30 Figure 1.8     SASH1 structure .................................................................................................36 Figure 3.1     cAMP production is decreased downstream of TLR2, TLR3 and TLR4                      in endothelial cells ..............................................................................................58  Figure 3.2     PTx inhibits LPS-induced ERK1/2 and JNK activation in endothelial cells .....60 Figure 3.3     LPS-induced Akt activation is inhibited by PTx in endothelial cells .................62 Figure 3.4     Heterotrimeric Gαi/o proteins are not important for LPS-induced NF-κB activation in endothelial cells ..............................................................65  Figure 3.5     Gαi/o proteins do not function along the TRAF6 signaling cascade .................67 Figure 3.6     PTx-sensitive Gαi/o are important for endothelial TLR2 and TLR3                      signaling ..............................................................................................................70  Figure 3.7     PI3K contributes to LPS-induced JNK activation ..............................................73 Figure 3.8     TLR4 binds to and signals through Gαi2 ...........................................................75 Figure 3.9     A model for Gαi/o in TLR4 signaling ................................................................79 Figure 4.1     Identification of SASH1 in MEF FADD KO cells by mass spectrometry                      analysis ................................................................................................................84  Figure 4.2     SASH1 promotes endothelial TLR4 signaling ...................................................86 Figure 4.3     SASH1 knockdown decreases LPS signaling ....................................................87 x  Figure 4.4       SASH1 does not play a role in signaling to the IFN pathway ..........................89 Figure 4.5       SASH1 promotes endothelial TLR3, but not TLR2, signaling ........................90 Figure 4.6       SASH1 interacts with TRAF6 through a conserved TRAF6 binding motif ....92 Figure 4.7       SASH1 interacts with the C-terminus of TRAF6 .............................................93 Figure 4.8       SASH1 regulates TRAF6 ubiquitination ..........................................................95 Figure 4.9       SASH1 does not interact with Ubc13 ...............................................................96 Figure 4.10     SASH1 does not interact with TRAF2 or TRAF3 ............................................98 Figure 4.11     SASH1 is a scaffold protein that binds to the TAK1-IKK complex ..............100 Figure 4.12     The TRAF6 binding mutant of SASH1 binds to TAK1 and IKKβ ................101 Figure 4.13     Generation of gene-targeted SASH1 mice .....................................................103 Figure 4.14     SASH1 mRNA is expressed in mouse tissues ................................................104  Figure 4.15     Sash1 is expressed predominantly in the endothelium of spleen, thymus                          and lung .........................................................................................................105  Figure 4.16     SASH1 is expressed in the parenchyma and microvasculature of murine                          liver, kidney and brain ..................................................................................107  Figure 4.17     A model for the role of SASH1 in endothelial TLR4 signaling .....................111              xi  LIST OF ABBREVIATIONS AC, adenylyl cyclase  AP1, activator protein 1  ASK1, apoptosis signal regulating kinase  ATP, adenosine triphosphate  Bcl10, B-cell CLL/lymphoma 10  BCR, B-cell receptor  Btk, Bruton’s tyrosine kinase  CAM, cellular adhesion molecule  cAMP, cyclic adenosine monophosphate  CARD11, caspase recruitment domain-containing protein 11  CARMA1, CARD-MAGUK protein 1  CBP, CREB binding protein  CXCR, chemokine receptor  DD, death domain  DMEM, Dulbecco modified Eagle medium  EGF, epidermal growth factor  eNOS, endothelial nitric oxide synthase  ELISA, enzyme-linked immunosorbent assay  ELM, eukaryotic linear motif  ERK, extracellular signal regulated kinase  FADD, Fas-associated death domain  FBS, fetal bovine serum  FCS, fetal calf serum xii   FHA, fork-head associated  FRET, fluorescence resonance energy transfer  Gαi/o, G inhibitory class  GAPDH, glyceraldehyde 3-phosphate dehydrogenase  GDF5, growth differentiation factor 5  GDP, guanosine diphosphate  GTP, guanosine triphosphate  GPCR, G-protein-coupled receptor  GPI, glycosyl-phosphatidylinositol  HA, hemagluttinin  HACS1, hematopoietic adaptor containing Src homology 3 and sterile α motif domains 1  HMEC, human microvessel endothelial cell  Hsp, heat shock protein  HUVEC, human umbilical vein endothelial cell  IB, immunoblotted  ICAM-1, intracellular adhesion molecule  IFNβ, interferon β  Ig, immunoglobulin  IL-1β, interleukin-1 β  IL-1R, interleukin-1 receptor  IL-4, interleukin 4  IL-6, interleukin 6  IL-8, interleukin 8 xiii   IL-10 interleukin 10  IL-13, interelukin 13  IP, immunoprecipitated  IP-10, interferon gamma inducible protein  IκB, inhibitor of nuclear transcription factor-κB  IKK, inhibitor of nuclear transcription factor-κB kinase  iNOS, inducible nitric oxide synthase  IRAK, interleukin-1 receptor associated kinase  IRF, interferon regulatory factor  ISRE, interferon stimulated response element  JIP3, c-jun NH2-terminal kinase interacting protein  JNK, c-jun NH2-terminal kinase  K48, lysine 48  K63, lysine 63  LBP, lipopolysaccharide binding protein  LOH, loss of heterozygosity  LPS, lipopolysaccharide  LRR, leucine-rich repeat  Mal, myeloid differentiation factor 88 adaptor-like protein  MALT1, mucosa-associated-lymphoid-tissue-lymphoma-translocation gene 1  MAPK, mitogen activated protein kinase  MEF, mouse embryonic fibroblast  MEKK3, mitogen-activated kinase kinase kinase 3 xiv   MIP2, macrophage inflammatory protein 2  MKK, mitogen-activated protein kinase kinase  MyD88, myeloid differentiation factor 88  NAK, nuclear transcription factor-κB activating kinase  NAP1, nuclear transcription factor-κB activating kinase associated protein  NEMO, nuclear transcription factor-κB essential modulator  NF-κB, nuclear transcription factor-κB  NLR, nucleotide-binding oligomerization (NOD)-like receptors  NO, nitric oxide  PAMP, pathogen associated molecular pattern  PAR, protease-activated receptor  PBS, phosphate buffered saline  PFA, paraformaldehyde  PI3K, phosphatidylinositol 3-kinases  PIP2, phosphatidylinositol 4,5-bisphosphate  PKB, protein kinase B  PKR, double-stranded RNA-activated serine/threonine kinase  PLCβ, phospholipase C β  PLCγ2, phospholipase C γ2  PMK1, p38 mitogen-activated protein kinas homolog  PRR, pattern recognition receptor  PtdIns, phosphatidylinositides  PTx, pertussis toxin xv   Pyk2, proline rich tyrosine kinase 2  RANK, receptor activator of nuclear transcription factor-κB  RGS, regulators of G protein signaling  RING, really interesting new gene  RIP1, receptor interacting protein 1  RNA, ribonucleic acid  ROS, reactive oxygen species  RLR, retinoid acid-inducible gene I (RIG-1)-like receptors  RT-PCR, reverse transcriptase polymerase chain reaction  S1P, sphingosine 1-phosphate  SAM, sterile α motif  SARM, sterile α and armadillo motif containing protein  SASH1, sterile α motif and Src homology 3 domain containing protein 1  SDS, sodium dodecyl sulfate  SH3, Src homology 3  SHIP, Src Homology 2 Domain-Containing Inositol-5-Phosphate  SIGIRR, Single immunoglobulin IL-1 receptor-related molecule  SLY, Src homology 3 domain expressed in lymphocytes  Src, V-Src avian sarcoma viral oncogene  Syk, spleen tyrosine kinase  TAB, transforming growth factor-β-activated kinase 1-binding protein  TAK1, transforming growth factor-β-activated kinase 1  TANK, tumor necrosis factor receptor associated factor family member-associated nuclear xvi  transcription factor-κB activator  TBK1, tumor necrosis factor receptor associated factor family member-associated nuclear transcription factor-κB activator binding kinase 1  TCR, T-cell receptor  TGF-β, transforming growth factor-β  TICAM1, TIR domain-containing adaptor molecule 1  TICAM2, TIR domain-containing adaptor molecule 2  TIR, Toll/interleukin-1 receptor  TIRAP, TIR domain-containing adaptor protein  TLR, Toll-like receptor  TNFα, tumor necrosis factor α  Tpl2, tumor progression locus 2  TRAF, tumor necrosis factor receptor associated factor  TRAM, Toll/interleukin-1 receptor-containing adaptor inducing interferon β-related adaptor molecule  TRIF, Toll/interleukin-1 receptor-containing adaptor inducing interferon β  TxB2, thromboxane B2  Uev1A, ubiquitin-conjugating enzyme E2 variant  Ubc13, ubiquitin-conjugating enzyme 13, S. cerevisiae homolog  VCAM-1, vascular cell adhesion molecule          xvii  ACKNOWLEGEMENTS  I would like to thank my wonderful parents, Wayne and Allana Dauphinee, for their support, encouragement and understanding throughout the years. I could not have succeeded without the unconditional support that you have provided throughout my life. I would also like to thank my supervisor Dr. Aly Karsan for his guidance and support. I am enormously grateful for the many opportunities that you have provided me with over the years. I have enjoyed learning with you. I would like to acknowledge the members of my laboratory for their help and support over the years. A special thank you to Iva Kulic, whose conversation and insight has made graduate school a much more enjoyable experience. Thank you to Megan Abbott for her tireless help with animal studies. Thank you to Zinaida Tebaykina, Eugene Park and Cindy Yang for their technical assistance over the years. Your work has contributed greatly to this project. I would also like to thank Dr. Josip Blonder at the National Cancer Institute for collaboration on the proteomics project. I am grateful to my graduate committee members, Drs. Robert Nabi, Yossi Av-Gay and Neil Reiner for their thoughtful comments and dedication to my project. I would like to express my gratitude to my undergraduate supervisor, Dr. Peter Dolphin, for providing the inspiration that started me down this road. This thesis would not even exist if you had not encouraged me at the start of my career. Finally, I owe my deepest gratitude to my husband, David Walsh, for sharing his passion for science and life with me. Your optimism, intellect and confidence are an inspiration. I could not have done this without you.  1          CHAPTER 1: INTRODUCTION                  2   INTRODUCTION 1.1 THE IMMUNE SYSTEM The immune system is a complex network of cells and processes that has evolved to distinguish self from non-self and thereby protect the host from invading pathogenic microorganisms. The immune system can be divided into two broad classes of functional responses - the innate response and the adaptive response. The innate immune response is a non-specific response that leads to an immediate defense and rapid clearance of the microorganism, but does not confer long-term protective immunity to the host. In contrast, the delayed antigen-specific responses mediated by the adaptive immune system create immunological memory, so that a subsequent exposure to the same antigen will induce a heightened response (1). Increasing evidence has suggested that the innate immune system is indispensable for initiation of the adaptive immune response in the host, and is important for adaptive immune functions such as clonal selection of lymphocytes and maturation of dendritic cells (2, 3). 1.2 THE INNATE IMMUNE SYSTEM The innate immune system is the host’s first line of defense against invading pathogens, present in individuals from birth, and highly evolutionarily conserved. The innate immune system provides four main barriers to infection: anatomic, physiologic, cellular and inflammatory (4). Physical barriers that prevent invasion by a microorganism include the skin and mucus membranes. In addition to providing a mechanical barrier, the skin hosts an acidic environment that inhibits most bacterial growth (5). Furthermore, the epidermal and dermal layers of the skin produce antimicrobial peptides such as cathelicidins and defensins (5, 6). In 3  addition to the skin, the mucus membranes, which line the passages of the body that are exposed to the external environment, also protect the host from pathogens (7). The membrane epithelium secretes a viscous fluid that traps microorganisms and tail-like projections, called cilia, act to sweep away invading pathogens (7). Additionally, normal flora residing in the mucus membranes will typically out-compete the invading pathogens for nutrients and attachment sites (8). Physiological barriers important in innate immunity include pH, temperature, and the production of various antimicrobial substances, such as complement. For example, pathogens are typically unable to survive the acidic environment of the gut and, for some microbes, host body temperature is not conducive to growth. In addition, complement, a group of serum proteins activated by bacterial products, plays a central role in pathogen destruction or clearance through activation of an enzymatic cascade (9). Cellular barriers to infection are a third mechanism to prevent pathogen invasion. The major function of these cellular barriers is internalization of pathogenic material through a process called phagocytosis (10). Pathogenic microbes are recognized by cell surface receptors that initiate phagocytic signaling cascades to ultimately control actin dynamics and bacterial uptake, resulting in delivery of the pathogen to the lysosome for degradation (10). The ultimate barrier to infection is inflammation, which is caused by the influx of blood to the affected tissue and is characterized by rubor (redness), tumor (swelling), calor (heat) and dolor (pain) (11). During inflammation, immune cells are recruited to the site of infection through the production of proinflammatory mediators, called cytokines. Proinflammatory cytokines activate the genes necessary for leukocyte recruitment and 4  passage from the circulation into tissue, as well as neutrophil degranulation and subsequent tissue damage (12). If these barriers are not sufficient to prevent invasion by a microorganism, the specific, adaptive immune response will be initiated. Together, these two branches of the immune response work together synergistically to protect the host and provide a strong, effective response to pathogenic insult. 1.3 SEPSIS  Sepsis occurs as the result of a systemic inflammatory response to a severe bacterial infection (13). When bacterial products are sensed by receptors on immune and endothelial cells, intracellular signaling cascades are initiated that activate families of transcription factors, such as nuclear transcription factor-κB (NF-κB) and interferon regulatory factor (IRF), to upregulate the production of proinflammatory mediators (14). Under normal conditions, a controlled cellular response to bacterial products will protect the host from infection. However, in sepsis, an initial hyperactivation of the systemic response leads to excessive production of these chemical mediators resulting in cellular injury, tissue damage and death (15). Subsequently, in an attempt to restore immune homeostasis, the production of anti-inflammatory mediators creates a state of immunosuppression, which results in the inability to combat infections post-sepsis (16). 1.4 ENDOTHELIAL CELLS  The vasculature is an extensive network of blood vessels that distributes oxygen and nutrient-rich blood throughout the body. Endothelial cells are the cells that line the interior surface of blood vessels and form a selectively permeable membrane between circulating blood and the surrounding tissues (17). The endothelium has many normal functions that help 5  to maintain organ homeostasis, including vasoregulation, vascular permeability and the ability to provide an anticoagulant surface (18). During infection, the regulated balance of these physiological functions is disturbed, leading to endothelial dysfunction and the clinical manifestations of sepsis, including edema, hypotension and organ failure (19). However, it remains unclear whether the changes in vascular function are a consequence of the direct interaction of bacterial products, such as lipopolysaccharide (LPS), with the endothelium or are secondary to the production of inflammatory mediators by immune cells. 1.5 ENDOTHELIAL DYSFUNCTION IN SEPSIS 1.5.1 Vasoregulation  Septic patients often become hypotensive, which is the result, in part, of the altered production of vasoactive mediators secreted by the endothelium. The endothelium secretes many chemically diverse compounds that maintain a balance in vascular tone, including vasodilating compounds, such as nitric oxide (NO) and prostacyclin (PGI2), as well as vasoconstrictors, such as angiotensin (20). In endothelial cells, NO production is mediated by an isoform of the nitric oxide synthase (NOS) family, called eNOS (21). Recent evidence suggests that in response to pathogens, direct activation of eNOS by LPS increases production of NO. In turn, this results in an increase in the expression of inducible NOS (iNOS) in the vasculature, contributing to impaired vasoregulation and endotoxemia in mice (22, 23). Indeed, mice that are deficient for eNOS are resistant to LPS-induced hypotension and endotoxic shock (22, 23). 1.5.2 Vascular permeability Endothelial cells act as gatekeepers to regulate the flux of material at the interface between the vasculature and the surrounding tissue (18). The endothelium uses both paracellular and transcellular pathways to regulate vascular permeability. The paracellular pathway involves 6  passage between adjacent endothelial cells, while transcellular pathways involve passage through the cell. The endothelial monolayer forms intercellular junctions that connect adjacent endothelial cells to restrict the passage of macromolecules into the underlying tissue. These intercellular junctions include the adherens junctions, the tight junctions and the gap junctions. In addition to mediating cell-cell contact, these junctions play an important role in relaying intracellular signals that control endothelial cell proliferation and function. The adherens junctions (also called zona adherens) are organized by homotypic interactions between transmembrane glycoproteins called vascular endothelial (VE) cadherin on adjacent cells. The intracellular domains of VE-cadherin interact with β-catenin to link adherens junctions to the cytoskeleton. In turn, the juxtamembrane domain of VE-cadherin binds to p120-catenin, and it is this interaction that provides stability to the adherens junction. The complex between VE-cadherin and the intracellular catenins is dynamic and dependent on the functional state of the cells (24). When the cells are migrating in vivo or loosely confluent in vitro, the interactions are weak and VE-cadherin, in its tyrosine phosphorylated form, is found primarily linked to p120 and β-catenin. In contrast, when the cells are stabilized or tightly confluent, VE-cadherin is dephosphorylated and complexed with actin and the catenin family member, plakoglobin. The interaction between junction proteins and the actin cytoskeleton is critical for maintaining junctional integrity in the endothelium. An increase in vascular permeability is mediated by contraction of the endothelial layer and phosphorylation of the light chain of nonmuscle myosin (25). This results in conformational changes that promote the interaction between actin and myosin, supporting the contractile state (25). This contraction essentially pulls VE-cadherin inward, thereby resulting in dissociation of adjacent cells and opening of intercellular channels. The modulation of cadherin-catenin 7  interactions by phosphorylation events is often catalysed by factors that are known to increase vascular permeability such as LPS, histamine and VEGF (26). LPS has been shown to activate Src family members, c-Src, Fyn and Yes, resulting in VE-cadherin phosphorylation and a loss of barrier integrity (27). Moreover, LPS can also contribute to a loss in barrier function through caspase-mediated cleavage of adherens junction proteins (28).  In contrast to the relatively abundant adherens junctions, tight junctions (also called zona occludens) are less abundant. They are composed of occludins, claudins and junctional adhesion molecule (JAM) proteins, which form homotypic interactions with adjacent cells to regulate the passage of ions and solutes into the underlying tissue. There is increasing evidence that JAMs play an important role in vascular inflammation (reviewed in (29)), which is supported by the observation that JAM family members are upregulated during inflammation and redistributed to the apical surface of the endothelial cell, thereby promoting leukocyte recruitment and extravasation (30, 31). The third class of intercellular junction is the gap junction, which is comprised of a collection of four different connexin (Cx) proteins, Cx37, Cx40, Cx43 and Cx45 (32). In addition to facilitating the exchange of ions and other small metabolites, gap junctions are important for communication between adjacent endothelial cells and the smooth muscle cell layer (33), mediating signals important for regulation of vasomotor tone (34). In addition to paracellular transport, molecules can also pass into the tissue through transcellular mechanisms. Depending on the vascular bed, the endothelium can be either continuous, fenestrated or discontinuous (35). Fenestrae are small cellular pores that extend through the cell and enable the rapid passage of molecules across the endothelial cell. 8  Typically, fenestrated endothelium is found in organs where a high rate of exchange of molecules occurs between the intravascular space and the surrounding tissue. For example, fenestrated endothelium is found in capillaries of the endocrine glands, gastrointestinal mucosa and the glomeruli of the kidney. These fenestrations are typically smaller (~60-70 nm in diameter) and permit the passage of water and small solutes, but not macromolecules (36, 37).  In contrast, continuous non-fenestrated endothelium is found in organs with limited filtration capacity such as the brain, heart, lung and skin. The discontinuous endothelium, also referred to as sinusoidal endothelium, is found in organs such as the liver and spleen, and owing to the increased diameter of the fenestrae (~100-200 nm in diameter), may be important for the exchange of lipid particles and cellular debris (38, 39). 1.5.3 Leukocyte recruitment and adhesion At the site of infection, recruitment of leukocytes occurs through association between selectin molecules, which are found on the surface of the circulating leukocytes (L-selectin) and the endothelium (E- and P-selectin), and oligosaccharides on the contacting endothelium (40). The integrin family, comprised of heterodimeric molecules expressed on the surface of leukocytes, also mediates the rolling and adhesion of the leukocytes along the endothelium. Cell adhesion molecules (CAMs), found on the surface of the endothelial cells (ICAM-1 and VCAM-1), are important for association with molecules of the integrin family (41). LPS has been shown to directly increase the expression of E-selectin and integrin counter receptors through an NF-κB-dependent mechanism (42-45). Furthermore, mice that are deficient for expression of endothelial E--and P-selectin are resistant to lethality in an animal model of sepsis (46).  9  1.5.4 Hemostasis  The endothelium is normally maintained in an anticoagulatory state through the action of thrombin on the endothelial cell surface (47). Thrombin binds to thrombomodulin on the surface of the endothelium and initiates the cleavage of protein C into its active form, thereby inhibiting the coagulation cascade (48). However, during infection, the action of thrombin shifts toward the cleavage of soluble fibrinogen in plasma into insoluble strands of fibrin. This is due, in part, to an LPS-induced reduction in the expression of thrombomodulin on the endothelium (49, 50). This shift toward a procoagulant state occurs in an attempt to limit the spread of invading pathogens. However, when uncontrolled, the result is disseminated intravascular coagulation, which is a common finding in septic patients (51). 1.6 LIPOPOLYSACCHARIDE The host recognition of small microbial molecules, called pathogen associated molecular patterns (PAMPs), initiates the inflammatory response characteristic of innate immunity (14). PAMPs include bacterial LPS, peptidoglycans, flagellin and lipotechoic acid, as well as viral nucleic acids (14). The most extensively studied PAMP is LPS (also called endotoxin). LPS is an essential component of the outer membrane of various Gram-negative bacteria, with the exception of the genus Sphingomonas, whose outer membrane contains glycosphingolipids in place of LPS (52, 53). LPS is an amphiphilic molecule composed of three structural elements: a core oligosaccharide, a covalently linked O-specific chain made up of repeating sequences of hydrophilic oligosaccharides and a membrane anchoring lipid A region, which is responsible for the proinflammatory properties of LPS (Figure 1.1) (52). LPS provides structural integrity to the bacteria and creates a negative charge that stabilizes the cell membrane and protects against entry of toxic compounds from the environment (54). 10  In addition, the diversity of the LPS molecular structures found on different bacterial strains provides the specific antigenic properties that activate host innate immunity. 1.7 TOLL-LIKE RECEPTORS  Cellular signaling cascades are initiated when PAMPs are recognized by a class of germ-line receptors, called Pattern Recognition Receptors (PRRs). PRRs are expressed on the surface of cells of the innate immune system and are comprised of several classes, including Toll-like receptors (TLRs) (55), retinoid acid-inducible gene I (RIG-I)-like receptors (RLRs) (56) and nucleotide-binding oligomerization domain (NOD)-like receptors (NLRs) (57). TLRs are a family of receptors classified on the basis of homology to the cytoplasmic domain of the interleukin-1 receptor (IL-1R) family, known as the Toll/IL-1R (TIR) domain. Mammalian TLRs were originally identified based on their homology to the Drosophila melanogaster Toll protein, which shows homology to the IL-1R TIR domain and plays a role in the antifungal response in adult flies (58, 59). To date, there have been 13 TLRs identified in mice (60, 61), and 11 TLRs found in humans (55). The family of TLRs is responsible for recognizing a diversity of ligands from bacteria and viruses (Table 1.1). Although the exact cellular location of human TLRs remains under study, the family can be segregated into those receptors that are found on the plasma membrane, TLR1, TLR2, TLR4, TLR5, TLR6, and TLR10, and those that are present in the endosomal compartment, TLR3, TLR7, TLR8 and TLR9 (Figure 1.2). The localization of TLR11, TLR12 and TLR13 has not been described. The intracellular location of TLRs that recognize bacterial and viral DNA/RNA may be an important determinant in the ability to distinguish between viral and self nucleic acids (62).  11                     Figure 1.1 Structure of LPS LPS is an amphipathic molecule composed of three structural elements: a covalently linked O-specific side chain consisting of N=4-40 repeating sequences of hydrophilic oligosaccharides, a core oligosaccharide, and a lipid A region that confers the proinflammatory properties of LPS. Kdo, 3-deoxy-D-manno-octulosonic; GlcN, glucosamine.  Phosphate Heptose N=4-40 NH3+ - - - - - - - Kdo GlcN  O-SPECIFIC CHAIN CORE REGION LIPID A - 12    Table 1.1 TLR ligands in human and mouse Ligands of TLR1-13 found in human (h) or mouse (m). Additional species and their respective TLRs are not shown as they are not relevant to this thesis.   Receptor Ligand Ligand location Species (h or m) TLR1 triacyl lipopeptides Bacteria h/m TLR2 glycolipids Bacteria h/m lipopeptides Bacteria lipoproteins Bacteria lipotechoic acid Bacteria HSP70 Host zymosan Fungi TLR3 double-stranded RNA Viruses h/m TLR4 lipopolysaccharide Bacteria h/m heat shock proteins Bacteria fibrinogen Host heparan sulfate Host hyaluronic acid Host TLR5 flagellin Bacteria h/m/ TLR6 diacyl lipopeptides Mycoplasma h/m TLR7 single-stranded RNA Viruses h/m TLR8 single-stranded RNA Viruses h/m TLR9 unmethylated CpG DNA Bacteria h/m TLR10 unknown unknown h TLR11 profilin Protozoa m TLR12 unknown unknown m TLR13 unknown unknown m 13  Toll-like receptor 4 (TLR4) was identified as the signaling receptor for LPS based on genetic evidence from the LPS-insensitive mouse strain, C3H/HeJ, which has a single point mutation in the TIR domain of TLR4. In addition, the C57BL/10ScCr mouse strain, which has a null mutation in the TLR4 gene, is also resistant to LPS (63, 64). The first host protein identified as a component of the LPS-sensing machinery was LPS-binding protein (LBP) (65). LBP belongs to a class of proteins, called acute-phase proteins, whose plasma concentrations rise in response to inflammation (66). It is synthesized primarily by hepatocytes as a 50-kDa protein and then released into the bloodstream as a 58-kDa protein following glycosylation (65). The primary role of LBP is to bring LPS to the cell surface through binding with high affinity to the amphipathic lipid A region of LPS and forming a ternary complex with the LPS receptor molecule, CD14 (65). In endothelial cells, LBP can also function to enhance LPS uptake (67). Formation of the complex between CD14 and LPS facilitates the transfer of LPS to the receptor complex of TLR4 and MD-2 (68). CD14 is a glycoprotein that can be found in two forms: a membrane-bound glycosyl- phosphatidylinositol (GPI)-anchored protein on the cell surface (mCD14), or a soluble proteolytic fragment found in the serum (sCD14) (69). Endothelial cells lack mCD14 and thus require sCD14 that has been cleaved from mCD14 bearing cells, such as macrophages and monocytes (70). While most endothelial cell lines do not express mCD14, it has been reported that early passage human umbilical vein endothelial cells (HUVECs) do express functional mCD14 and that experimental removal of mCD14 from the cell surface prevents endothelial detection of LPS and subsequent leukocyte recruitment (71, 72). Although most endothelial cells rely on sCD14 for LPS recognition, it has been shown that supplementation of cell culture medium with sCD14 can only partially rescue the ability of an endothelial cell 14                     Figure 1.2 Cellular localization of TLRs There are 11 TLRs characterized in humans, which are responsible for recognizing a diversity of pathogen-associated ligands. Mammalian TLRs can be classified into two groups, those that are found on the plasma membrane (TLR1/2, TLR4, TLR5, TLR2/6 and TLR10) and those that are found in the endosome (TLR3, TLR7, TLR8 and TLR9). The localization of TLR11, TLR12 and TLR13 is unknown.  TLR1 TLR2 TLR4 TLR5 TLR6 TLR2 TLR3 TLR7 TLR8 TLR9 Lipoproteins (Pam3CSK4) LPS Flagellin Lipoproteins (Zymosan) dsRNA (Poly(I:C)) ssRNA CpG DNA Plasma membrane Endosome TLR10 15  to recognize LPS (73, 74). Since CD14 lacks an intracellular domain, it does not have the capacity to initiate intracellular signaling (69), thus the complex of TLR4/MD-2 functions as the signaling entity for LPS. MD-2 is a secreted glycoprotein that binds to the ectodomain of TLR4 and is indispensable for LPS recognition (75). Signaling initiated by LPS requires two important domains in the TLR gene family: the extracellular domain containing leucine-rich repeats (LRRs) and the intracellular TIR domain (Figure 1.3). LRRs are a structural motif that form horseshoe-like 3D structures of repeating molecular sequences, typically rich in the hydrophobic amino acid, leucine (LXXLXLXXN, where X is any amino acid, L is leucine and N is asparagine) (76). The extracellular domain of TLR4 contains 23 LRRs (76) and it is this LRR structure that plays an essential role in binding ligand and the co-receptor molecules, CD14 and MD-2 (77).  The sequence of the TIR domain is highly similar between species and within the TLR family, which is reflected by the important role of docking intracellular adaptor molecules that will define the specificity of downstream signaling (78). The intracellular TIR domain is also important for dimerization of TLR4 and it is the binding of MD-2 to the extracellular domain that facilitates this homotypic interaction (79). Consequently, a dimerization-induced conformational change results in rearrangement of the TIR domain interface, which facilitates the binding of intracellular TIR-domain containing adaptor proteins (80). Ligation of TLR4 leads to activation of two intracellular signaling pathways, the specificity of which is determined by the particular adaptor molecules that are recruited to the intracellular domain of the receptor (81). These adaptors include myeloid differentiation factor 88 (MyD88), TIR-domain containing adaptor protein (TIRAP; also called MyD88 adaptor-like protein (Mal)), TIR-containing adaptor inducing IFNβ (TRIF; also called TICAM-1), and 16                      Figure 1.3 Structure of TLR4 TLR4 is composed of an extracellular domain containing LRRs responsible for ligand recognition and binding, and an intracellular TIR domain that mediates association with downstream adaptor molecules. LRR, leucine rich repeat; TIR, Toll/IL-1R domain.  LRR Transmembrane TIR 17                 Figure 1.4 TLR4 signaling LPS binds to the TLR4 receptor complex, resulting in recruitment of adaptor molecules. In MyD88-dependent signaling, IRAKs are recruited to the receptor via interaction with MyD88. IRAK1 recruits TRAF6, leading to activation of NF-κB and MAPKs, and proinflammatory cytokine production. In MyD88-independent signaling, TRIF associates with TRAF3 or TRAF6 leading to downstream activation of IRF3 or NF-κB, respectively. LPS, lipopolysaccharide; TLR4, Toll-like receptor 4; TIRAP, toll-interacting protein; MyD88, myeloid differentiation factor 88; IRAK, interleukin-1 receptor-associated kinase; TRAF6, tumor necrosis factor receptor associated factor-6; TAB, transforming growth factor-β-activated kinase-1 binding protein; TAK1, transforming growth factor-β-activated kinase-1; NF-κB, nuclear transcription factor-κB; IKK, IκB kinase; MKK, mitogen activated protein kinase kinase; JNK, c-jun NH2-terminal kinase; TRAM, Toll/interleukin-1 receptor- containing adaptor inducing interferon β-related adaptor molecule; TRIF, Toll/interleukin-1 receptor-containing adaptor inducing interferon β; TBK1, TANK-binding kinase; IFN, interferon; IRF3, interferon regulatory factor 3; Ub, ubiquitin.   TLR4 MyD88 IRAK1 IRAK4 TRAF6 TIRAP LPS MD2sCD14 Ub IKKα/β/γ NF-κB TAK1 TRAM TRIF MyD88-dependent signaling  MyD88-independent signaling  Proinflammatory cytokines IFNβ IRF3 TRAF3 TBK1 IKK-ε TAB2/3 MKK JNK/p38 IFN-inducible genes Ub Ub 18   TRIF-related adaptor molecule (TRAM; also called TICAM-2) (Figure 1.4). More recently, the sterile α and Armadillo motif containing protein (SARM) has been identified as a fifth member of the TLR-adaptor family (82). 1.8 MYD88-DEPENDENT TLR4 SIGNALING  MyD88 was originally characterized as an IL-6 inducible myeloid differentiation primary response gene during the differentiation of M1 myeloleukemic cells into macrophages (83). Subsequently, MyD88 was cloned and identified as an adaptor molecule belonging to the IL-1R signaling complex and shown to have a C-terminal TIR domain, which interacts with the intracellular tail of TLRs, and an N-terminal death domain (DD) that interacts with the serine/threonine kinase, IL-1 receptor associated kinase (IRAK) (84-86). Following receptor ligation, recruitment of MyD88 to the cytoplasmic domain of TLR4 is facilitated by TIRAP. TIRAP contains a phosphatidylinositol 4,5-bisphosphate (PIP2) binding domain that mediates recruitment to the plasma membrane and functions to shuttle MyD88 to the activated receptor complex (87). Furthermore, ligation of both TLR2 and TLR4 activates Bruton’s tyrosine kinase (Btk), resulting in phosphorylation of TIRAP and ultimately, NF-κB activation (88, 89). Both MyD88 and TIRAP are essential for MyD88- dependent signaling to activate NF-κB and proinflammatory cytokine production, as demonstrated by the fact that MyD88-/- mice are resistant to LPS-induced death (90) and TIRAP-deficient mice show delayed kinetics with respect to TLR2- and TLR4-induced NF- κB and MAPK activation (91, 92).  The assembly of the adaptor complex at the receptor permits interaction between MyD88 and IRAK family members. IRAK was originally identified based on homology to Pelle, a kinase in the D. Melanogaster Toll pathway (93). To date, there are four IRAK 19  family members, IRAK1, IRAK2, IRAK4 and IRAKM (94). IRAK4 is the critical kinase necessary for activation of the classical NF-κB pathway (95, 96), whereas IRAK1 selectively activates IRF5/7 and NF-κB downstream of TLRs (97, 98). Although IRAK1 and IRAK4 possess intrinsic serine/threonine kinase activity, IRAK2 and IRAKM are catalytically inactive. While IRAK2 plays a positive role in TLR signaling (99), IRAKM acts as a negative regulator of the signaling cascade (100).  The binding of IRAK4 to the receptor complex facilitates the transphosphorylation of IRAK1, inducing IRAK1 kinase activity (101). The autophosphorylation of IRAK1 results in its dissociation from MyD88 (85) and the ability to bind tumor necrosis factor (TNF) receptor associated factor-6 (TRAF6) (102), through three conserved TRAF6 binding domains (103).  TRAF6 belongs to a family of seven TRAF members that mediate signaling downstream of TNF receptor (TNFR) superfamily members (104, 105) (Figure 1.5). With the exception of TRAF1 and TRAF7, all TRAF proteins possess an N-terminal RING finger, followed by several zinc fingers, and a C-terminal TRAF domain, which consists of a coiled-coil domain and a highly conserved TRAF-C domain (106). The N-terminus functions as an E3 ubiquitin ligase that is critical for downstream activation (107) and the C-terminal TRAF domain is involved in self-association and heterologous protein interactions (106). In contrast to the other TRAF family members, TRAF7 does not contain a canonical TRAF domain, but rather has seven WD40 repeats at the C-terminus that facilitate interactions with its protein partners (108).  The complex of IRAK1 and TRAF6 dissociates from the receptor to form a complex with transforming growth factor-β (TGF-β)-activated kinase 1 (TAK1) and the adaptor molecules, TAK1-binding protein 1, 2 and 3 (TAB1, TAB2 and TAB3). TAB1 functions as an activator of TAK1 (109), while TAB2 and TAB3 bind polyubiquitin chains to link TAK1  20                    Figure 1.5 TRAF family protein domain structure The domain structure of TRAF family members, TRAF1-6. Aside from TRAF1, all other TRAF molecules have a RING domain that imparts E3 ligase activity. All TRAF molecules have atleast one zinc finger for ligase activity. The CC domain is important for homo- oligomerization of TRAF members and the TRAF domain is key for binding to protein partners. The WD40 domain is important for TRAF7 binding to protein partners.   Zn finger TRAF6 TRAF5 TRAF4 TRAF3 TRAF2 TRAF1 TRAF7 TRAF RING CC WD40 21  to TRAF6 and facilitate TRAF6 ubiquitination (110-112). The interaction between TAK1 and TAB2/3 is essential for activation of downstream signaling cascades (113). Although TAK1 phosphorylation occurs at the plasma membrane and is dependent on formation of the TRAF6-TAK1-TAB1/2/3 complex, it does not become active until the complex translocates to the cytoplasm (114). Once in the cytosol, it forms a complex with the E2 ubiquitin conjugating enzyme complex of Ubc13/Uev1A, to catalyze the formation of a polyubiquitin chain linked through lysine 63 (K63) of ubiquitin on TRAF6 (115). In contrast to the K48- linked polyubiquitin chains that target a protein substrate for proteasomal degradation, K63- linked polyubiquitin chains are important for proteasome-independent events, such as signal transduction (116). When TAK1 is activated, IRAK1 is released from the complex and eventually degraded by the ubiquitin-proteasome system (117). The formation of K63-linked polyubiquitin chains on TRAF6 results in TAK1-mediated phosphorylation of the inhibitor of NF-κB (IκB) kinase (IKK) complex and the MAPK kinases (MKKs) (118). The IKK complex is composed of two catalytic subunits, IKKα and IKKβ, and a regulatory subunit IKKγ (also known as NF-κB essential modulator, NEMO) (119). Activation of the IKKs leads to downstream phosphorylation of members of the IκB family, resulting in ubiquitin-mediated proteolysis of the IκB members, thus permitting the release and nuclear translocation of the transcription factor, NF-κB, activating proinflammatory genes (120). 1.8.1 Mitogen activated protein kinases In addition to the activation of NF-κB, activation of TAK1 also leads to the activation of a family of MAPKs consisting of p38, extracellular signal-regulated kinase (ERK) and c- jun NH2-terminal kinase (JNK) (118). Downstream of TLR4, MAPK are activated by both MyD88-dependent and independent mechanisms (91, 121). Relatively little is known about 22  the activation of the MAPKs downstream of the TLRs, however, tyrosine phosphorylation within the TLR4 signaling pathway has emerged as a critical event in the activation of downstream signaling. Phosphorylation of two tyrosine residues within TLR4 by the Lyn kinase facilitates the interaction between TLR4-MyD88 and the association with activated IRAK1, which is critical for the activation of NF-κB, p38 and JNK (122). The tyrosine kinase, Csk, is involved in activation of p38, ERK and NF-κB, but not JNK, possibly through the inhibition of Src family tyrosine kinases (123). In endothelial cells, LPS stimulation activates the proline-rich tyrosine kinase, Pyk2, leading to activation of NF-κB and p38, but not ERK, and subsequent upregulation of the chemokines, IL-8 and MCP1 (124, 125). A functional phospholipase C gamma 2 (PLCγ2) is important for phosphorylation of IκB, p38 and Akt, but not ERK, in bone marrow derived macrophages (126). In MEFs, MAPK kinase kinase 3 (MEKK3) is indispensable for the activation of p38, but not ERK, through interaction with TRAF6 (127). In contrast, the activation of p38 and ERK in endothelial cells does not require TRAF6, since a dominant negative mutant of TRAF6 does not inhibit their activation (128). In immune cells, LPS-induced reactive oxygen species (ROS) control the formation of a complex between TRAF6 and apoptosis signal-regulating kinase 1 (ASK1) to allow activation of p38, but not JNK (129). The activation of ERK involves the MAP3K, Tpl2 (also known as Cot), in bone marrow derived macrophages and B cells (130). It has been shown that ERK activation is not dependent on TRAF6 in endothelial cells and macrophages (128, 131). In contrast, we and others have seen that activation of JNK lies downstream of TRAF6 in both endothelial cells and macrophages (128, 131). In macrophages, the activation of JNK is dependent on interaction between the scaffold protein JNK-interacting protein 3 (JIP3) and TLR4 (132). 23  Genetic deletion of the double-stranded RNA (dsRNA)-activated serine/threonine kinase R (PKR) reduces JNK and NF-κB, but not p38 or ERK, activation in primary alveolar macrophages (133). In neutrophils, JNK activation downstream of TLR4 involves the tyrosine kinase, Syk, and PI3K (134). 1.8.2 Phosphoinositide 3-kinases (PI3Ks)  PI3Ks are a family of enzymes that catalyze the phosphorylation of the hydroxyl group at the 3-position of the inositol ring of phosphoinositides (PtdIns) (135). The resulting products, PtdIns 3,4-bisphosphate and PtdIns 3,4,5-triphosphate, act as second messengers to activate downstream effectors, such as Akt (also known as protein kinase B, PKB) to control cell survival, remodeling of the cytoskeleton and intracellular trafficking events (136, 137). The PI3K family consists of four classes of kinases, IA, IB, II and III. Class IA enzymes are comprised of a regulatory subunit (p85α, p55α, p50α, p85β, or p55γ) and a catalytic subunit (p110α, p110β or p110δ), which are activated downstream of multiple TLRs and the IL-1R (138-142). B cells isolated from mice deficient in the regulatory subunits of PI3K fail to proliferate in response to LPS, indicating that PI3K plays a positive role in TLR4 signaling (143). Furthermore, in endothelial cells, activation of NF-κB by LPS is mediated by PI3K- Akt, as suggested by inhibition of NF-κB activation by a p85 dominant-negative mutant. (144). PI3K has been shown to positively regulate cytokine expression through formation of a complex between the p85 regulatory subunit, TLR4 and MyD88 in murine macrophages (145). However, more recently, sustained interaction of this complex at the intracellular domain of TLR4 has been shown to inhibit NF-κB luciferase activity (146). Indeed, PI3K has been reported to both positively and negatively regulate TLR signal transduction. In endothelial cells, monocytes and dendritic cells, inhibition of PI3K-Akt enhances 24  LPS-induced cytokine production (147-149). Moreover, macrophages isolated from p85-/- mice show reduced activation of MAPK pathways and cytokine production in response to LPS (150). Thus, PI3K may act as a molecular switch to control TLR signaling dependent on cellular needs. 1.9 HETEROTRIMERIC G-PROTEINS IN TLR4 SIGNALING  Heterotrimeric guanine nucleotide binding proteins (G-proteins) are the intracellular partners to seven transmembrane domain G-protein-coupled receptors (GPCRs). Under basal conditions, G-proteins are composed of a GDP-bound α subunit tethered to a βγ heterodimer. Binding of a ligand to the GPCR results in GDP-GTP exchange on the α subunit and subsequent dissociation of Gα from Gβγ, allowing the components to bind and regulate downstream effector molecules, such as ion channels and the enzymes, adenylyl cyclase (AC), PI3K and PLCβ (Figure 1.6) (151). Based on their sequence identity and the ability to differentially regulate downstream effectors, Gα subunits have been divided into four classes: Gαs, Gαi/o, Gαq and Gα12/13 (Table 1.2). GPCRs couple selectively to Gα subunits to determine the specificity of downstream signaling. Regulation of GPCR signaling is maintained by the exchange of GTP for GDP by a class of “regulators of G protein signaling” (RGS), resulting in reassociation of the Gαβγ heterotrimer, and termination of the signal (152). The family of arrestin proteins also mediates this cycle of activation/termination through translocation of β-arrestin from the cytoplasm to the cell membrane to interact with the activated receptor complex. This results in uncoupling of the receptor and the G-protein, internalization of the receptor, and signal attenuation (153). More recently, β-arrestins have also been implicated as scaffold molecules that positively coordinate signaling downstream of GPCRs (154). 25                    Figure 1.6 Gαi/o signaling Upon receptor-ligand interaction, GDP-GTP exchange occurs on the α subunit, resulting in dissociation of Gα from Gβγ. This leads to regulation of various downstream effector molecules, such as ion channels (K+ and Ca2+) and the enzymes, AC, PI3K and PLCβ. AC, adenylyl cyclase; ATP, adenosine triphosphate; cAMP, cyclic adenosine monophosphate; GTP, guanosine triphosphate; GDP, guanosine diphosphate; RGS, regulators of G-protein signaling; PLCβ, phospholipase C β; PI3K, phosphoinositide 3-kinase;   αATP cAMP α AC PLCβ  PI3K K+ Ca2+ β γ β γ β γ β γ GTP GDP RGS γ β 26         Table 1.2 Heterotrimeric Gα subunit effectors   Effector molecules activated, or inhibited as indicated, downstream of heterotrimeric G proteins of various classes. AC, adenylyl cyclase; HCK, hematopoietic cell kinase; PLCβ, phospholipase C; Btk, bruton’s tyrosine kinase; PI3Kβ, phosphoinositide 3-kinase; MEK5, mitogen activated kinase kinase 5; ASK1, apoptosis signaling regulated kinase 1.  Subunit Gene Distribution Effectors Gαs class AC Gαs GNAS Ubiquitous HCK, c-Src Gαsxl GNASXL Neuroendocrine Ca2+ channel Gαsolf GNAL Olfactory epithelium, brain Gαi/o class AC (inhibition) Gαi1 GNAI1 Widely distributed RhoGEFs Gαi2 GNAI2 Ubiquitous c-Src Gαi3 GNAI3 Widely distributed Rap1GAP Gαo GNAO Neuronal, neuroendocrine Gαz GNAZ Neuronal, platelets Gαgust GNAT3 Taste cells, brush cells Gαt-r GNAT1 Retinal rods, taste cells Gαt-c GNAT2 Retinal cones Gαq/11 class PLCβ Gαq GNAQ Ubiquitous Btk Gα11 GNA11 Almost ubiquitous Pyk2 Gα14 GNA14 Kidney, lung, spleen PI3Kβ Gα15/16 GNA16 (Gna15) Hematopoietic cells RhoGEFs Csk Gα12/13 class Src Gα12 GNA12 Ubiquitous Btk Gα13 GNA13 Ubiquitous MEK5 Inhibition of RasGAP Pyk2 ASK1 Cdc42 p115-RhoGEF 27  G-proteins of the G inhibitory class (Gαi/o) have been shown to be important for LPS-induced MAPK activation through association with CD14 (155). Studies that use pertussis toxin (PTx), an inhibitor of receptor-Gαi/o protein coupling, have shown decreased activation of MAPK, but not NF-κB, in monocytes and macrophages (156, 157). Furthermore, genetic deletion of Gαi2 or Gαi1/3 results in a decrease in LPS-induced production of proinflammatory mediators, such as IL-10, TNF-α and thromboxane (TxB2), in macrophages (158). In contrast, the production of these mediators is increased in splenocytes isolated from Gαi2-/- and Gαi1/3-/- mice (158). This study highlights the need for an understanding of cell-specific effects of these, and other, novel LPS signaling molecules. 1.10 MYD88-INDEPENDENT SIGNALING  MyD88-independent signaling was identified in MyD88-deficient mice, which retain the ability to activate both NF-κB and MAPK pathways, albeit with delayed kinetics (90). While the MyD88-dependent pathway is primarily responsible for promoting the expression of proinflammatory cytokines, such as interleukin (IL)-6 and tumor necrosis factor α (TNFα), the MyD88-independent pathway induces expression of interferon (IFN)-inducible genes, such as IFNγ inducible protein 10 (IP-10), through activation of the transcription factor, IFN regulatory factor 3 (IRF3) (159). IRF3 is a member of a family of transcription factors that are involved in the induction of type I IFNs that translocate to the nucleus upon phosphorylation and dimerization (160). Once inside the nucleus, IRF3 controls IFNβ transcription by recruiting co-activators, p300 and CBP (160). MyD88-independent signaling begins with recruitment of the adaptor molecule, TRAM, to the cytoplasmic domain of TLR4 (161). TRAM acts as a bridging adaptor to bring a second adaptor molecule, TRIF, to the receptor complex (161). Myristoylation of TRAM 28  results in localization to the plasma membrane, followed by translocation of the TLR4/adaptor complex to endosomes (162). Endocytosis of the receptor is essential for TRIF-dependent signaling leading to IFNβ activation (163). Although it has been suggested that endothelial cells do not express TRAM, and are thus restricted to signaling through MyD88, we and others have seen that TRAM is expressed in various endothelial cell types (Dauphinee et al. unpublished, (74, 164)). While TRAM is restricted to the TLR4 pathway, TRIF has been shown to play an essential role in IRF3 activation downstream of both TLR3 and TLR4 (165). The recruitment of these adaptor molecules to the receptor complex results in binding of either TRAF6 or TRAF3 leading to late-phase activation of NF-κB or IFNβ, respectively (166). TRAF3 interacts with TRIF and shuttles the protein kinase, TRAF family member-associated NF-κB activator (TANK) binding kinase 1 (TBK1, also called NAK) to the TLR signaling complex within endosomes (166). This results in recruitment of the non- canonical IκB kinase homolog, IKKε, and the NF-κB-activating kinase-associated protein 1 (NAP1) to the signaling complex and subsequent phosphorylation and activation of IRF3, followed by IFNβ production (167, 168). The partial localization of TLR4 in the endosomal compartment is in agreement with the fact that other endosomal TLRs, such as TLR3/7/8/9, are able to induce IFNs, whereas TLRs found on the plasma membrane are not able to induce this response (169). In a parallel MyD88-independent pathway, TRIF recruits TRAF6 and receptor interacting protein (RIP1), also leading to activation of NF-κB (170). 1.11 NEGATIVE REGULATION OF TLR4 SIGNALING  Signaling through TLRs can induce very potent inflammatory responses, thus modulation of the signaling cascade is imperative to protect the host from the inflammation-associated damage induced by bacterial and viral products. There are 29  numerous molecules that have been described as regulators of the TLR4 signaling pathway (Figure 1.7). Due to their relevance to this thesis, some of these regulators will be described in more detail here. 1.11.1 Fas-associated death domain (FADD)  FADD is a proapoptotic adaptor molecule that couples the cytoplasmic domain of death receptor molecules, such as Fas, to effector caspases (171). FADD has been shown to play an inhibitory role in LPS-induced NF-κB and JNK activation (172, 173). Our lab has shown that the mechanism by which FADD exerts its regulation is through interaction with both IRAK1 and MyD88, thereby preventing the obligatory association of these two molecules (173). Furthermore, we have shown that the negative regulation imparted by FADD results in a decrease in LPS-induced proinflammatory cytokine production and endothelial cell sprouting (173). 1.11.2 A20 A20 is a zinc-finger protein that was originally identified as an antiapoptotic gene and has since been shown to have both ubiquitinating and deubiquitinating activities (174-176). TLR4 regulation by A20 is dependent on its ability to interact with TRAF6 (177) and catalyze the removal of ubiquitin chains from TRAF6 (178), thereby inhibiting TLR4-mediated NF-κB activation and cytokine production (178, 179). In endothelial cells, the expression of A20 is induced by LPS in an NF-κB-dependent manner, suggesting that A20 may function in a feedback loop to regulate TLR4 signaling in these cells (180). In vivo, A20-deficient mice show decreased responsiveness to LPS and develop severe inflammation (181). Furthermore, mice reconstituted with A20-deficient stem cells exhibit higher concentrations of serum TNF, compared to controls, following LPS stimulation. (178, 181). 30                   Figure 1.7 Endogenous regulators of TLR4 signaling The TLR4 signaling regulators relevant to this thesis are shown. LPS, lipopolysaccharide; TLR4, Toll-like receptor 4; TIRAP, toll-interacting protein; MyD88, myeloid differentiation factor 88; IRAK, interleukin-1 receptor-associated kinase; TRAF6, tumor necrosis factor 6; NF-κB, nuclear transcription factor-κB; JNK, c-jun NH2-terminal kinase; Tollip, Toll- interacting protein; SHIP, Src homology 3 domain expressed in lymphocytes; TRAM, Toll/interleukin-1 receptor-containing adaptor inducing interferon β-related adaptor molecule; TRIF, Toll/interleukin-1 receptor-containing adaptor inducing interferon β; IFN, interferon; SARM, sterile α and armadillo motif containing protein; Ub, ubiquitin.            TLR4 MyD88  IRAK1 IRAK4 TRAF6 TIRAP LPS MD2sCD14 Ub   NF-κB TRAM TRIF          Proinflammatory               cytokines  IFNβ JNK/p38 IFN-inducible genes Ub Ub ST2 SIGIRR FADD A20 β-arrestin Tollip IRAK-M MyD88s SHIP SARM 31  1.11.3 β-arrestin  The arrestin family is comprised of four subtypes: the ubiquitously expressed β-arrestin 1 and β-arrestin 2, and two visual arrestins that are expressed exclusively in photoreceptors of the retina (182). The β-arrestins are most well known for their role in regulating cell signaling pathways through desensitization and/or endocytosis of receptor molecules, but they have more recently been implicated as scaffold molecules that regulate cell signaling cascades (183). β-arrestin 1 and β-arrestin 2 inhibit NF-κB and AP-1 activation following stimulation with IL-1β by interacting with TRAF6 to prevent its oligomerization and autoubiquitination (184). Furthermore, mouse embryonic fibroblasts isolated from Arrb1-/-Arrb2-/- mice have elevated levels of transcript for the proinflammatory cytokine, IL-6 (184) and mice deficient for β-arrestin 2 are more susceptible to endotoxic shock (184). 1.11.4 Sterile α and Armadillo repeat motif (SARM)  SARM is the fifth TLR adaptor molecule identified and has been studied in mammals, D. melanogaster and C. elegans protein. In mammalian systems, SARM has been shown to be a negative regulator of TRIF-dependent NF-κB and IFN activation by its physical association with TRIF (82), which may prevent the interaction of TRIF with signaling molecules downstream of TLR3 and TLR4. The orthologous C. elegans protein, called TIR-1 (185, 186), is required for activation of the p38 homolog, PMK-1, in worm immunity (187) and RNA interference of TIR-1 leads to decreased worm survival following fungal infection (188). 1.11.5 Additional endogenous regulators of TLR4 signaling Tollip was originally identified as a protein partner of the IL-1R complex (189). It forms a complex with IRAK to suppress IRAK kinase activity and thus the downstream 32  activation of NF-κB (189, 190). Indeed, ectopic expression of Tollip in human dermal microvessel endothelial cells (HMEC) inhibits LPS-induced NF-κB activation (191). In contrast, bone marrow derived macrophages isolated from Tollip-/- mice do not exhibit defects in signaling to NF-κB or MAPK, but do show a decrease in production of the proinflammatory cytokines, such as IL-6 and TNFα, suggesting that Tollip is important for regulating the magnitude of proinflammatory responses to LPS (192).  Molecules such as ST2 and single immunoglobulin IL-1 receptor-related molecule (SIGIRR), which mimic the intracellular domain of TLR4, have also been shown to inhibit LPS signaling (193-195). ST2 (also known as T1) is a TIR-domain-containing orphan receptor that, when overexpressed in HEK293 cells, results in a decrease in LPS-induced NF- κB activation (194). The inhibition imparted by ST2 is the result of interaction between its TIR domain and the TIR domains of MyD88 and IRAK, thus sequestering these adaptors during initiation of the LPS signal (193). ST2 has been shown to be expressed in endothelial cells and thus may play a role in regulation of TLR4 signaling in these cells (196). SIGIRR is another TIR-domain-containing receptor that has been shown to interact with TLR4 to attenuate interaction with downstream adaptor molecules (195). Since expression of SIGIRR is relatively low in endothelial cells, it may play a minor role in regulation of endothelial TLR4 signaling (197).  Splice variants of the adaptor molecules MyD88 and IRAK have also been shown to play a negative regulatory role in TLR4 signaling (198). MyD88s is a splice variant of MyD88 that lacks the intermediate domain required for recruitment of IRAK4, thereby acting as a dominant-negative molecule to inhibit NF-κB activation (198, 199). The expression of MyD88s in endothelial cells has not been determined. IRAK-M is a member of the 33  IRAK/Pelle family and is expressed exclusively in macrophages (200). IRAK-M plays a negative role in TLR4 signaling by preventing the dissociation of IRAK4 and MyD88, thus inhibiting the formation of the IRAK-TRAF6 complex (100).  Src Homology 2 domain-containing Inositol-5-Phosphate (SHIP) is a cellular phosphatase that catalyses the removal of a phosphate group from PtdIns 3,4,5-triphosphate, a product of PI3K activity (201). However, the role of SHIP as a negative regulator of LPS-induced signaling is independent of PI3K. Rather, the regulation imparted by SHIP is proposed to be through inhibiting formation of the complex between TLR4 and MyD88 (202). 1.12 SCAFFOLD PROTEINS IN IMMUNE SIGNALING The complexity of signaling cascades dictates a need for the proper localization and coordinated regulation of the many proteins involved in transduction of a signal from a cellular receptor to downstream effector molecules. Scaffold proteins are ideal candidates for mediating these functions. They can be defined as molecules that bind and assemble multiple signaling proteins to regulate signal transduction either by altering subcellular localization of the complex, coordinating positive and negative regulators or protecting the complex from inactivation (203). The function of scaffold molecules has been investigated using mathematical modeling to suggest that scaffold proteins can function to amplify or inhibit a pathway depending on variables such as: relative expression of various signaling molecules (including the scaffold molecule itself), localization of signaling components, stability of the interactions and basal catalytic activity of the cell (204, 205). Engineered scaffold proteins have also been useful to demonstrate that scaffold molecules can create signaling thresholds that mediate whether a signal is analogue, digital, sustained, transient or oscillatory (203). 34  The ability to modulate the amplitude and duration of a signal, as suggested by the oscillatory model, might represent the ability of a scaffold protein to facilitate negative feedback loops through binding/activation of a negative regulator of a signaling pathway. Although much emphasis has been placed on studying TLR signaling cascades, very little is known about the role of scaffold proteins in regulating these signaling pathways. 1.12.1 Pellino family The Pellino proteins are a family of E3 ubiquitin ligases that were originally identified in D. melanogaster as an interacting partner of the IRAK1 homologue, Pelle (206). Highly similar in their domain structure, all three mammalian Pellino proteins (Pellino 1, 2 and 3) contain an N-terminal FHA domain to mediate interaction with phosphorylated IRAK (207) and a C-terminal RING domain that has E3 ubiquitin ligase activity to catalyze K63-linked polyubiquitination of IRAK1 (208). K63-linked polyubiquitination of IRAK1 has been shown to be important for binding of IKKγ and subsequent activation of NF-κB (209, 210). Pellino proteins have also been shown to bind to TRAF6 and TAK1 downstream of IL- 1R signaling (211). This has led to the model that Pellino proteins function as scaffold molecules to facilitate the association between ubiquitinated IRAK1-IKKγ and the other Pellino binding partners, TRAF6 and TAK1 (208). However, the function of Pellino proteins remains unclear. 1.13 SAM AND SH3 DOMAIN CONTAINING PROTEIN 1 (SASH1)  SASH1 was originally cloned from a human brain cDNA library and found to be ubiquitously expressed in most human tissues (212). SASH1 is found on chromosome 6q24.3 and is comprised of 20 exons that encode two transcripts of 4.4 and 7.5 kb, differing in the location of their polyadenylation signals (213). SASH1 is a large protein of 1247 amino acids 35  with a predicted molecular weight of ~140 kDa that contains two sterile-α-motif (SAM) domains and a Src homology 3 (SH3) domain. SAM domains are protein-protein interaction domains that can form homo- or heterooligomers with SAM domain-containing proteins and can even bind to non-SAM domain-containing proteins (214). SH3 domains also mediate protein-protein interactions through binding to proline-rich sequences with a consensus motif of PXXP (where P is proline and X is any amino acid) (215). In addition to the protein- interaction domains, SASH1 has a predicted coiled-coil domain and a proline-rich region, which both lie at the N-terminus of the SASH1 protein sequence (Figure 1.8). The presence of multiple interaction domains within SASH1 suggests that it may play a role as an adaptor or scaffold molecule in cell signaling.  SASH1 mRNA is ubiquitously expressed in human tissues, with the highest levels of expression found in the lung, placenta, spleen and thymus (213). In silico profiling and expression analysis in tumor tissues from breast cancer patients originally implicated SASH1 as a tumor suppressor in breast cancer and other solid tumors, such as lung and thyroid (213). Loss of heterozygosity (LOH) at 6q24.3 is associated with increased tumor size and poor prognosis (213). LOH at this region has also been associated with the invasiveness of primary colorectal cancers (216). Indeed, SASH1 mRNA expression is reduced in 42% of colon cancer patients with an even greater reduction in mRNA expression seen in post- operative metastases in the liver (217). Although this correlative data suggests that SASH1 may function as a tumor suppressor, no functional data has been published to corroborate these findings.   36            Figure 1.8 SASH1 structure A. SASH1 genomic structure. The SASH1 gene is composed of 20 exons. B. SASH1 protein domain structure. The genomic location of the SH3 and SAM domains is denoted. SH3, Src homology 3 domain; SAM sterile α motif; CC, coiled coil; ATG, start codon; TAG, stop codon.   37  SASH1 belongs to a family of SAM and SH3 adaptor proteins, consisting of SH3 domain expressed in lymphocytes (SLY1), hematopoietic adaptor containing SH3 and SAM domains 1 (HACS1) (also called SLY2) and SASH1 (213, 218-221). SLY1, so named for its expression in lymphoid tissues, such as spleen, thymus and bone marrow, is serine phosphorylated upon B or T cell receptor stimulation (221). SLY1 mutant mice, expressing a protein that lacks the serine phosphorylation site and part of the nuclear localization sequence (SLY1d/d), exhibit impaired lymphoid organ development, defects in antigen-receptor mediated lymphocyte activation, weakened humoral immune responses and prolonged allograft survival, suggesting that SLY1 is critical for mediating adaptive immune responses in mice (218). HACS1 is expressed in normal and malignant hematopoietic tissues and is upregulated in B cells stimulated with IL-4, IL-13, anti-IgM, anti-CD40 and LPS, suggesting a role in B cell activation and differentiation cascades (219, 222). Collectively, these functions indicate that the SLY family plays a role in immune regulation. 1.14 AIM OF THE PRESENT STUDY The signaling pathways activated downstream of TLR4 have been extensively studied in immune cells. However, signaling through TLRs in endothelial cells remains comparatively unexplored. During bacterial infection, the normal physiological functions of the endothelium are disturbed, leading to the clinical manifestations of sepsis, including edema, hypotension and organ failure (19). Indeed, recent studies have highlighted the importance of endothelial-specific signaling in response to pathogenic ligands (223). Therefore, furthering our knowledge of the molecular signaling pathways activated by LPS in endothelial cells will contribute to our understanding of the endothelial dysfunction and 38  vascular collapse characteristic of sepsis.  The overarching aim of this study was to characterize the function of novel signaling molecules in the endothelial response to LPS.  Heterotrimeric G-proteins have been implicated in the LPS-induced activation of MAPKs in macrophages and monocytes (156). However, a role for these proteins has not been clearly established in endothelial cells. Furthermore, it remains unclear if heterotrimeric G-proteins participate in signaling downstream of other TLRs. The first part of this thesis investigates the role of pertussis toxin sensitive Gαi/o proteins in endothelial TLR signaling. We posit that heterotrimeric G proteins, of the Gαi/o class, are important for signaling downstream of TLRs in endothelial cells. Indeed, the data presented in this thesis demonstrates that Gαi/o proteins are important for signaling mediated by multiple pathogen- associated ligand-receptor interactions. The work presented in the second part of this thesis arose from work previously published from our laboratory, which shows that FADD is a negative regulator of TLR4 signaling in endothelial cells and FADD null cells demonstrate hyperactivation of JNK in response to LPS (160). We hypothesized that proteins important for promoting activation by LPS would be differentially expressed in FADD wild-type versus FADD null cells. A mass spectrometric (MS) analysis of LPS stimulated FADD null cells identified SAM and SH3 domain-containing protein (SASH1) as a putative candidate for positive regulation of the signaling events initiated by LPS. The work in the second part of this thesis characterizes the role of SASH1 in endothelial TLR4 signaling. In summary, the data presented herein demonstrates that SASH1 acts as a scaffold protein to positively regulate TLR4 signaling in endothelial cells. 39  Together, the experiments presented in this thesis further our understanding of the molecular events that are critical for coordinating the response to pathogen-associated ligands in endothelial cells.                       40            CHAPTER 2: MATERIALS AND METHODS                  41  2.1 MATERIALS LPS, pertussis toxin (PTx), 3-isobutyl-1-methylxanthine (IBMX) and forskolin (FSK) were all purchased from Sigma-Aldrich (St. Louis, MO). The synthetic double-stranded RNA molecule, Poly(I:C), and the synthetic tripalmitoylated lipopeptide, Pam3CSK4, were purchased from InvivoGen (San Diego, CA). LY294002 was purchased from Cell Signaling Technology (Beverly, MA). TrueBlot anti-rabbit immunoglobulin immunoprecipitation beads and rabbit immunoglobulin G (IgG) were obtained from eBioscience (San Diego, CA). 2.2 CELL CULTURE The human microvascular endothelial cell line (HMEC) was provided by the Centers for Disease Control and Prevention (Atlanta, GA). HMEC lines were cultured in MCDB 131 medium (Invitrogen, Carlsbad, CA) supplemented with 10% heat-inactivated fetal calf serum (FCS) (HyClone, Logan UT), 2 mM glutamine (Sigma-Aldrich, St. Louis, MO), 10 ng/mL epidermal growth factor (EGF) (Sigma-Aldrich, St. Louis, MO) and 100 U each of penicillin and streptomycin (Sigma-Aldrich, St. Louis, MO). Human umbilical vein endothelial cells (HUVEC) were isolated as previously described (224) and cultured in MCDB 131 (Invitrogen, Carlsbad, CA) supplemented with 10% heat-inactivated FCS, 10% heat- inactivated fetal bovine serum (FBS) (HyClone, Logan, UT), 2 mM glutamine, 20 ng/mL endothelial cell growth supplement (BD Bioscience, San Jose, CA), 16 U/mL Heparin (Sigma-Aldrich, St. Louis, MO) and 100 U each of penicillin and streptomycin. HEK293- TLR4-MD2-CD14, HEK293T, mouse embryonic fibroblasts (MEFs) and the retroviral producer cell line AmphoPhoenix (gift of G. Nolan, Stanford University, Stanford, CA) were cultured in Dulbecco modified Eagle medium (Invitrogen, Carlsbad, CA) supplemented with 10% FBS, 2 mM glutamine and 100 U each of penicillin and streptomycin. 42  HEK293-TLR4-MD2-CD14 were maintained in medium containing 10 μg/mL blasticidin and 50 μg/mL of hygromycin. All cells were maintained at 37°C in 5% CO2. 2.3 SASH1 ANTIBODY PURIFICATION A rabbit polyclonal antibody to human SASH1 was raised for our laboratory against a synthetic peptide conjugated to keyhole limpet haemocyanine (KLH) to recognize amino acid 8-20 at the N-terminus of the deduced SASH1 protein sequence (University of British Columbia). The SASH1 antibody was then purified by protein-A sepharose chromatography. Briefly, SASH1 peptide was coupled to a CNBr-activated sepharose column and washed with Tris-buffered saline (TBS) solution (50 mM Tris, 0.15 M NaCl, 0.02 % NaN3, pH 7.4). Serum was added to the column and rotated overnight at 4°C. Unbound material was washed off with TBS, the antibody was eluted with 5 M MgCl2 and concentrated using a Centriprep® filter column (Millipore, Billerica, MA) according to manufacturer’s directions. 2.4 RECOMBINANT PLASMIDS AND GENE TRANSFER Human SASH1 constructs were amplified by PCR and cloned into pcDNA3 by restriction digest to generate N-terminally tagged (HA and Flag) constructs (see Table 2.1 for primers). The expression plasmid encoding Flag-TAK1 was generated by PCR and cloned into pcDNA3 (see Table 2.1 for primers). The expression plasmid encoding Gαi2-GFP was generated by PCR and cloned into pEGFPN2 by restriction digest (see Table 2.1 for primers). Myc-SASH1ΔT6BD was obtained by inverse PCR to generate a deletion of amino acids 852- 860 (see Table 2.1 for primers). Flag-TRAF6ΔN, Flag-TRAF6ΔC and Flag-TRAF6ΔCC were cloned from Flag-TRAF6 into pcDNA3 or the retroviral vector LNCX (gift of A. D. Miller, Fred Hutchinson Cancer Research Center, Seattle, WA) (128) by restriction digest. Other plasmids were obtained as follows: Flag-TRAF6 (Tularik, San Francisco, CA), Flag- 43  IKKβ, Flag-IKKα and Flag-IKKγ (gift of T. D. Gilmore, Boston University, Boston, MA), Flag-TLR4 (Addgene, Cambridge, MA), ISRE-Luc (Stratagene, La Jolla, CA) and NF-κB- Luc (gift of F. R. Jirik, University of Calgary, Calgary, AB). Transient transfections were performed using TransIT-siQUEST® transfection reagent according to the manufacturer’s instructions (Mirus Bio Corporation, Madison, WI). For retroviral transduction, transient transfections of the retroviral packaging cell line AmphoPhoenix were carried out using TransIT-siQUEST® transfection reagent according to the manufacturer’s instructions (Mirus Bio Corporation, Madison, WI). Viral supernatants were used to transduce HMEC 48 h post- transfection, for a total of four rounds of infection approximately every 12 h. Cell lines were selected using G418 (300 μg/mL). Expression of the proteins was verified by immunoblotting. 2.5 cAMP ASSAY HMEC were pretreated with the phosphodiesterase inhibitor, IBMX (1 mM, 15 min) and then treated with forskolin (FSK; 10 µM, 30 min) and either LPS (100 ng/mL), Pam3CSK4 (100 ng/mL) or Poly(I:C) (2 μg/mL) for 30 min. In some instances, cells were pretreated with PTx for 16 h prior to the treatment with IBMX, FSK and LPS. Cells were lysed in a solution of 0.1 N HCl for 20 min at room temperature, cell lysis was confirmed by visualization and lysates were centrifuged at 600 × g for 5 min. Intracellular cAMP levels were assayed using the cAMP Direct Immunoassay Kit (Calbiochem, San Diego, CA), according to manufacturer’s instructions. 2.6 RNA ISOLATION Total RNA was isolated from cultured cells in vitro or tissues in vivo using the TRIZOL® method (Invitrogen, Carlsbad, CA) according to manufacturer’s directions. 44  2.7 REVERSE TRANSCRIPTION AND POLYMERASE CHAIN REACTION (RT- PCR) Total RNA was extracted for reverse transcription and polymerase chain reaction and cDNA synthesis was performed using 2.5 μg total RNA. DNA contamination was removed by DNase treatment (Invitrogen, Carlsbad, CA). First strand cDNA synthesis was performed using random priming (Invitrogen, Carlsbad, CA) and Superscript® II reverse transcriptase (Invitrogen, Carlsbad, CA). Samples were stored at -20°C until further analysis or used for PCR to amplify cDNA to determine gene expression levels in cells or tissues. The PCR reaction was performed using 3 μL of cDNA and 1.25 U Taq DNA polymerase (New England Biolabs, Ipswich, MA). PCR was performed on a PTC-200 PCR cycler (BioRad Laboratories, Hercules, CA) and samples were stored at 4°C until analysis by agarose gel electrophoresis in a gel containing 1.0% agarose (w/v) in Tris-acetate EDTA buffer (TAE; 0.04 M Tris-acetate and 1 mM EDTA, pH 8.0). PCR for cloning was performed using Platinum® Taq DNA polymerase High Fidelity (Invitrogen, Carlsbad, CA). The gene specific primers used are listed in Table 2.1. For RT-PCR, relative amounts of mRNA were normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) levels in each sample. In negative controls, using samples without reverse transcription, no PCR product was obtained, demonstrating the absence of genomic DNA.      45  Table 2.1 List of primers  Human RT-PCR primers Human GAPDH Forward GGACCTGACCTGCCGTCTAGAA Reverse GGTGTCGCTGTTGAAGTCAGAG Human SASH1 Forward AAGCTGGTAAACTCCACTCGCAGA Reverse TGCAGAGAGCGATTCGTCAAGTCA Human GNAI2 Forward GGCCGAGCGCTCTAAGATGATCGA Reverse GGACAGGTCATCAGGGAGCACGCC Mouse RT-PCR primers Mouse GAPDH Forward AATGTGTCCGTCGTGGATCT Reverse CCCTGTTGCTGTAGCCGTAT Mouse SASH1 Forward TGTGGAAAGCCTTCGGAGTTCTCT Reverse ACTTTGTTGTTCAGCAGGCCCATC Mouse CD31 Forward CCAACTTCACCATCCAGAAGG Reverse GGGAATGGCAATTATCCGC Mouse vWF Forward GTGGGCATGATGGAGAGGTTA Reverse GCATCGCTCCTGAGACATTTC Mouse VE-cadherin Forward CCTGGTATAACCTGACTGTGGAG Reverse TGCCCATACTTGACCGTGATG Mouse E-cadherin Forward TTAGGTTAGAGGGTTATCGCGT Reverse TAATTTTAGGTTAGAGGGTTATTGT Mouse SLY1 Forward TGAGCTGAACATCATGGACCCACA Reverse ATGTCCACCTTGGGTTCTGACACT Mouse SLY2 Forward TGCCTCAGACAAGGAGAAACACCA Reverse TCCTCCACTGCTTGAAGTCTTGCT Cloning primers Flag-SASH1 Forward GGAATTCCCACCATGGATTACAAGGATGACGACG ATAAGGAGGACGCGGGAGCAGCTGGCCCG Reverse CCTCGAGCTACATGGCCTCAGGGCCTGG HA-SASH1 Forward GGAGATCTGCCACCATGGACTACCCATACGATG TTCCAGATTACGCTGAGGACGCGGGAGCAG Reverse CCTCGAGCTACATGGCCTCAGGGCCTGG Flag-TAK1 Forward GGAATTCCCACCATGGATTACAAGGATGACGAC GATAAGTCTACAGCCTCTGCCGCCTCCTCCTCC Reverse CCCTCGAGTCATGAAGTGCCTTGTCGTTTCTGC Gαi2-GFP Forward GGCTCGAGGCCACCATGGGCTGCACGTTGAGC GCCGAAGAC Reverse CCGCGGATCCCGAAGAGGCCGCAGTCCTTC Inverse PCR primers Myc-SASH1 Forward GGTACCGGAGATCTGCCACCATGGCAGAACAAA AACTTATTTCTGAAGAAGATCTGGAGGACGCGGGA GCAG DelT6BD Reverse CACTTCAGGTACAATCTGAGGGGGTTGCTCAG CACCAGGCTC DelT6BD Forward GAGCCTGGTGCTGAGCAACCCCCTCAGATTGT ACCTGAAGTG SASH1-clone rev Reverse CCTCGAGCTACATGGCCTCAGGGCCTGG Primer Sequence (5' to 3') 46  Table 2.1 (continued) List of primers  2.8 PROTEIN ASSAY  Cell lysates were assayed for total protein concentration using the BioRad DC™ Protein Assay Kit (BioRad Laboratories, Hercules, CA) based on the Lowry method (225). Protein samples were diluted 10-fold with distilled water and 5 μL added to a dye solution containing 25 μL alkaline copper tartrate solution, 0.5 μL surfactant solution and 200 μL Folin reagent. After incubation at room temperature for 15 min, absorbance was measured at 560 nm and protein concentrations determined against a standard curve of bovine serum albumin (BSA) at concentrations ranging from 0.125 to 2.0 mg/mL. 2.9 IMMUNOBLOT ANALYSIS Immunoblotting was performed using samples containing 50-100 μg protein in SDS- PAGE sample buffer. The samples were incubated at 95°C for 5 min to disrupt non-covalent interactions and loaded onto 10% acrylamide gels. Molecular weight markers were electrophoresed adjacent to the samples. Immediately after SDS-PAGE was performed, the proteins on the gel were electro-transferred to nitrocellulose membrane (BioRad Laboratories, Hercules, CA). After the transfer was complete, the blots were submerged in a blocking solution of 5% skim milk powder (w/v) in Tris-buffered saline containing Tween- Genotyping primers Intron 14-15_1 Forward GTAGGCTTGTTCATCTTG Intron 14-15_2 Forward GTGTACAGAGATGTTTATGTG Intron 14-15_3 Forward CCAATCTTATCCTATCTCAG Intron 14-15_4 Forward CATGTGCACATGTGCACA Intron 14-15_5 Forward GATCTTTGTTCCCTGGGC bgeo Reverse AGTATCGGCCTCAGGAAGATCG Geno_WT Forward TCGAGAACTTCCATGCCCATCCTT Reverse ACTTTGTTGTTCAGCAGGCCCATC Geno_KO Forward TCGAGAACTTCCATGCCCATCCTT Reverse AATCAACTTTGGAGACATGCGGGC Primer Sequence (5' to 3') 47  20 (TTBS; M Tris-HCl, pH, NaCl, and 0.05% Tween-20) for 1 h at room temperature. The blots were incubated with the desired primary antibody overnight at 4°C in 5% skim milk TTBS, or 5% BSA (w/v) for phospho-specific antibodies. The primary antibody concentrations were as follows: anti-phosphoJNK (1:1000), anti-phospho-p38 (1:1000), anti- phospho-ERK1/2 (1:2000), anti-ERK1/2 (1:2000), anti-phospho-Akt (1:500) and anti-Akt (1:1000) from Cell Signaling Technologies (Danvers, MA); anti-JNK (1:1000), anti-TRAF6 (1:1000) and anti-Ubiquitin (1:1000) from Santa Cruz Biotechnology (Santa Cruz, CA); anti- p38 (1: 1000, Stressgen, Ann Arbor, MI); anti-FlagM2 (1:2500, Sigma-Aldrich, St Louis, MO); anti-HA (1:1000, Covance, CA); anti-GFP (Roche, Mississauga, ON) and anti-SASH1 (1:1000). Following incubation with the primary antibodies, the blots were washed three times with TTBS for 10 min each and then incubated in the appropriate secondary antibody (1:5000, HRP-conjugated IgG, Sigma-Aldrich, St. Louis, MO) in 5% skim milk-TTBS. The blots were then washed again three times for 10 min each in TTBS. This was followed by 1 min incubation in a 1:1 mixture of enhanced chemiluminescence (ECL) reagents (PerkinElmer Life Sciences, Boston, MA). The membranes were blotted with Whatman paper to remove any excess ECL and wrapped in plastic wrap for autoradiography. 2.10 CO-IMMUNOPRECIPITATION HEK293T or HEK293-TLR4-MD2-CD14 cells were co-transfected with 5 μg of each expression plasmid using TransIT-siQUEST® transfection reagent (Mirus Bio Corporation, Madison, WI) according to manufacturer’s instructions. Forty-eight hours later, cell lysates were collected using modified RIPA buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% NP-40, 0.25% sodium deoxycholate plus protease inhibitors) and protein concentrations were determined. Subsequently, 1 mg of protein was precleared by incubation 48  with agarose beads for 1 h followed by incubation overnight with either anti-FlagM2-agarose beads or control IgG-agarose beads (Sigma-Aldrich, St. Louis, MO). Immunoprecipitation was also performed with anti-HA (1 μg) or anti-TRAF6 (1 μg) overnight followed by incubation with protein G agarose beads (Cell Signaling Technologies, Danvers, MA) for an additional 24 h. Following immunoprecipitation, the beads were washed three times in lysis buffer and boiled in SDS-PAGE sample buffer for immunoblot analysis. 2.11 TRAF6 UBIQUITINATION To examine polycovalent ubiquitination of TRAF6, HEK293-TLR4-MD2-CD14 cells were transfected with 5 μg of expression plasmids using TransIT-siQUEST® transfection reagent (Mirus Bio Corporation, Madison, WI) according to manufacturer’s directions. Cells were serum starved 24 h post-transfection and stimulated with LPS (100 ng/mL, 30 min) the next day. Cells were lysed (50 mM Tris, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Triton X- 100, 0.1% SDS, 20 mM N-ethylmaleimide and protease inhibitors) and protein concentrations were determined. Following this, 3 mg of protein lysate was used for immunoprecipitation with anti-TRAF6 (1 μg) for 3 h followed by overnight incubation with TrueBlot anti-rabbit immunoglobulin immunoprecipitation beads (eBioscience, San Diego, CA). After three washes, beads were boiled in SDS-PAGE sample buffer for immunoblot analysis. 2.12 LUCIFERASE REPORTER ASSAY HMEC were seeded in 24-well plates to a density of 100,000 cells per well and transient transfections were performed 24 h later using Superfect transfection reagent (Qiagen, Mississauga, ON) according to manufacturer’s directions. HMEC were co-transfected with 300 ng of the expression plasmids, or stably transduced with lentivirus, 49  as indicated, and 7.5 ng of a plasmid encoding an NF-κB luciferase reporter (with five tandem NF-κB binding sites, pNF-κB-Luc) or an interferon stimulated response element luciferase reporter (pISRE-Luc, Stratagene, La Jolla, CA). As an internal control, 7.5 ng of the Renilla luciferase plasmid, pRL-CMV (Promega, Madison, WI, for pNF-κB-Luc) or pRL-TK (Promega, Madison, WI, for pISRE-Luc), were transfected simultaneously. Cells were serum starved overnight 24 h post-transfection and stimulated with LPS (100 ng/mL), Pam3CSK4 (100 ng/mL) or Poly(I:C) (2 μg/mL) for 8 h, followed by passive lysis. Luciferase activity was measured by dual luciferase assay (Promega, Madison, WI) on a luminometer. 2.13 RNA INTERFERENCE siRNAs targeting human SASH1 mRNA (NM_015278); shSASH1 573- GCTAATGATGGTCAAAGATTCAAGAGA-593, human Gαi2 mRNA (NM_002070) shGαi2 266-GCAACCTGCAGATCGACTTTG-286, and shRandom GTTGCTTGCCACGTCCTAGAT were cloned into the HpaI and XhoI sites of the pLentilox3.7 vector. Constructs were sequence verified and screened for efficient knockdown. Lentiviral particles were produced from HEK293T cells by co-transfection of 6 μg pLentilox shRNA vector, 3 μg pVSVG, 3 μg pMDL g/p RRE, and 3 μg RSV-REV for a 100 mm dish using TransIT-siQUEST® transfection reagent according to the manufacturer’s instructions (Mirus Bio Corporation, Madison, WI). Viral supernatants were used to transduce target cells and green fluorescent protein-positive cells were selected by flow sorting (FACS 440, Becton Dickson, NJ).   50  2.14 DETERGENT RESISTANT MEMBRANE (DRM) ISOLATION  MEF FADD WT or FADD KO were treated with LPS (100 ng/mL, 5 min) and harvested for sucrose density gradient separation. Briefly, cells were washed three times with ice-cold phosphate buffered saline and lysed in 400 μL of ice-cold lysis buffer (50 mM HEPES pH 7.4, 150 mM NaCl, 1% Brij-35 and protease inhibitors). Cell lysates were sheared by five successive passages through 26 gauge needles and then centrifuged at 800 × g for 5 min at 4°C to pellet nuclei. The supernatant was mixed 1:1 with 80% sucrose (w/v) and transferred to 11 × 34 mm ultracentrifuge tubes (Beckman Instruments, Inc., CA). The samples were overlaid with 1.3 mL of 30% sucrose and 500 μL 5% sucrose and centrifuged using a TLS-55 swinging bucket rotor on a TL-100 tabletop ultracentrifuge (Beckman Instruments, Inc., CA) at 100,000 × g for 18 h at 4°C. Following centrifugation, 11 fractions of 200 μL each were collected starting at the top of the gradient. Fractions 1-6, representing the interface between 30% and 5% gradients, were pooled and prepared for mass spectrometry. Briefly, an equal volume of 20% trichloroacetic acid (TCA) was added to pooled protein fractions and incubated on ice for 30 min. Samples were centrifuged at 12,000 × g for 15 min at 4°C and the pellet was washed with 10% TCA and centrifuged at 12,000 × g for 15 min at 4°C. The sample was washed twice in 300 μL cold acetone, the pellet dried, and stored at -80°C until further analysis. 2.15 MASS SPECTROMETRY ANALYSIS DRM protein pellets (~30 μg each) were solubilized in 0.2 mL of CH3OH (60% v/v) buffered with NH4HCO3, pH 7.9. Samples were digested overnight using a 1:20 (w/w) trypsin/protein ratio. Trypsin-catalysed 16O (FADD WT)/18O (FADD KO) exchange/labeling was carried out as previously described (226). Peptide pools were combined and lyophilized 51  prior to strong cation exchange (SCX) fractionation. SCX fractionation was performed using a HP 1090 LC System (Agilent Technologies, CA) as previously described (226). Sixteen fractions were collected, lyophilized and reconstituted in 0.1% TFA prior to nanoflow (nano) reversed-phase (RP) liquid chromatography (LC)-Fourier-transform ion cyclotron resonance (FT-ICR) mass spectrometry (MS) (LTQ-FT, ThermoElectron, CA), as previously described (214). Briefly, the MS spectra were analysed using SEQUEST algorithm against the UniProt mouse proteomic database from the European Bioinformatics Institute (http://www.ebi.ac.uk/). For a peptide to be acceptably identified it had to achieve charge state and proteolytic cleavage-dependent cross correlation (Xcorr) scores of 2.0 for [M + H]1+, 2.3 for [M + 2H]2+, 3.3 for [M + 3H]3+, and a minimum delta correlation score (ΔCn) of ≥0.08. Relative abundances for differentially labeled protein-specific peptides (i.e. peptide matched to only one protein) were calculated using PepQuan™ software (ThermoElectron, CA) and reported as heavy-to-light 18O/16O ratios for a particular protein identified by ≥2 peptides. 2.16 IN SILICO ANALYSIS Prediction of TRAF6 as a SASH1 interacting partner was found using the Eukaryotic Linear Motif resource (ELM) (http://elm.eu.org/links.html) (227). 2.17 ELISA HMEC were seeded in 24-well plates to a density of 100,000 cells per well and transient transfection of expression plasmids was performed 24 h later using Superfect transfection reagent (Qiagen, Mississauga, ON) according to manufacturer’s directions. Cells were serum starved 24 h post-transfection and then stimulated with LPS (100 ng/mL, 6 h) the 52  following day. Cell culture supernatants were assayed for IL-6 (eBioscience, San Diego, CA) or IP-10 (R&D Systems, Minneapolis, MN) by ELISA following manufacturer’s directions. 2.18 GENERATION OF SASH1 GENE-TRAP MICE The ES cell line CC0006 from the Sanger Institute Gene Trap Resource was used to generate Sash1 gene-trap mice. Briefly, ES cells were injected into blastocysts from 129Ola mice, the resulting chimeras were mated with C57BL/6 mice and agouti F1 progeny that were heterozygous for the Sash1 allele (+/-) were backcrossed to C57BL/6 mice and interbred to produce F2 progeny. A comparison of the EST from the Sanger database and Ensembl suggested that the insertion site for the gene-trap vector, pGT0LxfT2, is within intron 14-15 of Sash1, which creates a fusion protein consisting of β-galactosidase fused to the N- terminus of Sash1 (exon 1-14). PCR of genomic DNA, followed by DNA sequencing, was used to map the exact location of the insertion into the Sash1 gene (see Table 2.1 for primers). Genomic DNA was isolated from littermates by tail-digest with proteinase K followed by PCR using primers specific to the insertion site of the gene-trap construct to distinguish homozygous mice (see Table 2.1 for primers). 2.19 β-GALACTOSIDASE STAINING Adult mouse tissues from Sash1+/- mice and their wild-type littermate controls were dissected and immediately fixed in 2% PFA at 4°C for 4 h. Tissues were washed three times with PBS and cryopreserved in 15% sucrose at 4°C for 1 h, followed by incubation in 30% sucrose at 4°C overnight. Tissues were placed into OCT compound and frozen at –80°C until sectioning. Sections (10 μm) were fixed with 0.2% glutaraldehyde solution and washed three times with PBS, followed by incubation with β-galactosidase staining solution (100 mM sodium phosphate, pH 7.3, 2 mM MgCl2, 0.01% sodium deoxycholate, 0.02% Nonidet-P40, 53  1 mg/mL X-gal, 5 mM ferrocyanide and 5 mM ferricyanide) at 37°C overnight. Sections were stained with nuclear fast red until the desired intensity was reached and dehydrated with ethanol and xylenes according to standard methods. 2.20 ISOLATION OF MOUSE ENDOTHELIAL CELLS Adult lung tissue was harvested from C57BL/6 mice and rinsed with cold PBS. Tissue was minced and digested with collagenase for 2 h at 37 °C, followed by 15 min incubation with DNase (0.2 mg/mL) (Sigma-Aldrich, St. Louis, MO). Tissue homogenate was passed through a 100 μm cell strainer, centrifuged and red blood cell lysis was carried out. Cells were passed through a 40 μm cell strainer and then incubated with rat anti-mouse CD105 hybridoma supernatant (or non-specific hybridoma supernatant). Cells were sorted for a CD105+ population (FACS 440, Becton Dickson, NJ) and resuspended in TRIZOL® to be processed for RNA and RT-PCR. 2.21 STATISTICAL ANALYSIS Results are expressed as means ± s.d. Data were analysed using a two-tailed Student’s t-test using the GraphPad Prism statistical program. P values of less than 0.05 were considered significant. Error bars depict s.d.               54                      CHAPTER 3: HETEROTRIMERIC Gαi/o PROTEINS MODULATE ENDOTHELIAL TLR SIGNALING                        55   3.1 INTRODUCTION Previous studies have suggested a role for heterotrimeric G-proteins in LPS signaling in macrophages and monocytes (156). Genetic deletion of the Gαi isoforms (Gαi1, Gαi2 and Gαi3) suggests that Gαi proteins differentially regulate TLR-stimulated mediator production in a cell-type specific manner (158). These in vivo studies highlight the need to understand the role Gαi/o proteins play in the various cell types important in innate immunity. Indeed, the molecular mechanism of activation of Gαi/o proteins by LPS in endothelial cells is unknown. Recently, Lentschat et al. implicated heterotrimeric G-proteins in TLR4, but not TLR2, signaling in endothelial cells (228). This study was performed using the wasp venom- derived peptide, mastoparan, to disrupt heterotrimeric G-protein signaling. However, others have shown that mastoparan binds the toxic lipid A portion of the LPS molecule with high affinity, effectively reducing the activity of LPS in a manner similar to that of the LPS neutralizing agent, polymyxin B (229). Furthermore, mastoparan potently reduced TLR4 mRNA expression in a dose-dependent manner (229). These findings suggest that the results obtained from studies utilizing mastoparan as an inhibitor of TLR4-mediated G-protein signaling should be interpreted with caution since the effects of this treatment may be to specifically bind and inhibit LPS rather than inhibit G-protein activation or signaling. Thus, the role of G-proteins in TLR signaling in endothelial cells remains unclear. Gαi proteins have been shown to co-immunoprecipate with the GPI-anchored CD14 (155). Since CD14 does not contain an intracellular domain available for binding adaptor molecules, such as the Gαi subunit, it is likely that CD14 cooperates with another receptor that has a transmembrane domain and intracellular docking region that could 56  facilitate formation of the CD14-Gαi complex. Moreover, endothelial and epithelial cells lack the membrane-bound form of CD14 found on many cell types, and thus require soluble CD14 cleaved from cells found circulating in the plasma that bear mCD14 (70). Therefore, it is of interest to investigate the role of heterotrimeric G-proteins in cells lacking GPI- anchored CD14 and to determine whether another receptor with an intracellular signaling domain, such as TLR4, could be involved in this complex. In this chapter, the role of heterotrimeric G-proteins in endothelial cells in response to various TLR ligands was investigated. The findings presented in this chapter demonstrate that heterotrimeric G- proteins are important for the activation of MAPK and Akt downstream of TLR2, TLR3 and TLR4 and suggest a common mechanism of activation for multiple TLRs.              57  3.2 RESULTS 3.2.1 TLR ligands activate heteotrimeric Gαi/o proteins in endothelial cells Heterotrimeric Gαi/o proteins participate in activation of macrophages and monocytes downstream of TLR4 (156, 157). Heterotrimeric Gαi/o proteins, once activated, result in the inhibition of adenylyl cyclase, thereby reducing intracellular cAMP levels. To determine whether Gαi/o proteins are activated downstream of TLRs, intracellular cAMP was measured in response to various TLR ligands. HMEC were pretreated with PTx or vehicle control, followed by stimulation with LPS, Pam3CSK4 or Poly(I:C) and assayed for cAMP production. Treatment of HMEC with LPS decreased forskolin-induced cAMP production and cAMP levels were restored with PTx pretreatment (Figure 3.1). The same trend was also observed in Pam3CSK4 and Poly(I:C) stimulated HMEC. Thus, these results demonstrate that Gαi/o proteins are activated downstream of TLR2, TLR3 and TLR4 in endothelial cells.                       58                    Figure 3.1 cAMP production is decreased downstream of TLR2, TLR3 and TLR4 in endothelial cells  HMEC were pretreated with 1 mM IBMX for 15 min and then treated with forskolin (FSK; 10 µM, 30 min) and either LPS (100 ng/mL, 1 h), Pam3CSK4 (100 ng/mL, 1 h) or Poly(I:C) (2 μg/mL, 1 h) as indicated. In some instances, cells were pretreated with PTx for 16 h. Cell lysates were used to determine intracellular cAMP levels. *P < 0.001 relative to FSK treated cell lysates and #P ≤ 0.001 relative to ligand treated cell lysates, as determined by Student’s t- test. Error bars indicate s.d., n = 3 independent experiments. 59   3.2.2 Gαi/o proteins contribute to LPS-induced activation of ERK1/2 and JNK in endothelial cells Heterotrimeric Gαi/o proteins contribute to TLR4-induced ERK1/2, p38 and JNK activation in macrophages and monocytes (156, 157). To determine whether Gαi/o proteins play a role in endothelial LPS signaling, HUVECs were pretreated with PTx and stimulated with LPS for various times. Pretreatment of cells with PTx attenuated ERK1/2 activation and significantly reduced JNK activation in HUVECs (Figure 3.2A). However, in contrast to monocytes, PTx pretreatment did not affect phosphorylation and activation of p38 in HUVEC (Figure 3.2A). Interestingly, p38 activation appears to be slightly delayed in HMEC treated with PTx (Figure 3.2B). Pretreatment of HMEC with cholera toxin, an inhibitor of the stimulatory class of G proteins (Gαs), did not alter MAPK activation (data not shown). These results suggest that Gαi/o proteins differentially regulate TLR4 activation of MAPK pathways in endothelial cells, as compared to macrophages and monocytes (156, 157). 3.2.3 Pertussis toxin does not activate the MAPKs in endothelial cells PTx has been suggested to activate MAPK pathways in peritoneal macrophages (230). To determine whether PTx activates the MAPKs in endothelial cells, HMEC were stimulated with PTx, or LPS as a positive control, for the times indicated. Activation of JNK, p38 and ERK1/2 was apparent after 60 min LPS stimulation as detected by immunoblotting with phospho-specific antibodies to each of the MAPKs. In contrast, PTx stimulation did not result in activation of JNK, p38 or ERK1/2 (Figure 3.2C). Thus, in contrast to macrophages, PTx does not activate MAPK pathways in endothelial cells.   60           Figure 3.2 PTx inhibits LPS-induced ERK1/2 and JNK activation in endothelial cells  (A) HUVEC or (B) HMEC were pretreated with PTx (100 ng/mL, 1 h) and stimulated with LPS (100 ng/mL) for the indicated times. Cell lysates were subjected to immunoblot analysis with antibodies against the phosphorylated form of ERK1/2, JNK and p38. (C) HMEC were stimulated with LPS (100 ng/mL as a positive control) or PTx (100ng/mL) for the times indicated. Activation of the MAPKs was assessed by immunoblotting using phospho-specific antibodies for ERK1/2, JNK and p38. In all cases, total MAPK protein was used as a control for protein loading.  61  3.2.4 Gαi/o proteins play a role in LPS-induced activation of Akt in endothelial cells PI3Ks are a family of kinases that catalyze the phosphorylation of phosphoinositides to yield the phosphorylated lipid products, phosphatidylinositol (PtdIns) 3,4-bisphosphate and PtdIns 3,4,5-triphosphate (135), which can then act as second messengers to activate downstream events, including activation of Akt (136). We, and others, have shown that PI3K/Akt is activated downstream of TLR4 in endothelial cells, however the mechanism of activation of PI3K/Akt in response to LPS is not well characterized. Several studies also suggest that the PI3K isoform, PI3Kβ, is activated downstream of GPCRs (219, 220). To investigate whether Gαi/o play a role in the activation of PI3K downstream of TLR4, endothelial cells were treated with LPS in the presence or absence of PTx. As seen in Figure 3.3, pretreatment with PTx resulted in a decrease in the active, phosphorylated form of Akt compared to cells treated with vehicle alone. Similar results were obtained for both HMEC and HUVEC (Figure 3.3A and 3.3B, respectively).              62                   Figure 3.3 LPS-induced Akt activation is inhibited by PTx in endothelial cells  (A) HMEC or (B) HUVEC were pretreated with PTx (100 ng/mL, 1 h) and stimulated with LPS (100 ng/mL) for the indicated times (in minutes). Cell lysates were subjected to immunoblot analysis with a phospho-Akt antibody. Total Akt was used as a control for protein loading.    63   Table 3.1 Densitometry analysis of PTx-sensitive Gαi/o endothelial TLR4 signaling.  *Data are represented as relative quantification of phospho-protein:total-protein and expressed as the fold-change relative to untreated. Error indicates s.d., n ≥ 2 independent experiments.  3.2.5 Gαi/o proteins do not contribute to LPS-induced activation of NF-κB in endothelial cells Activation of NF-κB results in the expression of proinflammatory molecules following LPS stimulation (231). In endothelial cells, NF-κB and JNK are activated along the MyD88/IRAK axis which bifurcates downstream of TRAF6 to activate transcription (128). To examine the effect of inhibition of Gαi/o on endothelial NF-κB activation, we used an NF-κB luciferase reporter assay. LPS-induced NF-κB luciferase reporter activation was unchanged by pretreatment with PTx (Figure 3.4A). Conversely, pretreament of HMEC with mastoparan abolished NF-κB luciferase activity, which is in agreement with the earlier finding that mastoparan acts indirectly on LPS signaling by neutralizing the ligand and decreasing TLR4 mRNA expression (data not shown) (229). Activation of NF-κB is coupled -PTx +PTx -PTx +PTx -PTx +PTx -PTx +PTx 0 1 0.40 ± 0.51 1 1.23 ± 0.46 1 1.00 ± 0.13 1 0.44 ± 0.19 30 1.69 ± 0.75 0.30 ± 0.58 2.49 ± 0.57 2.20 ± 0.48 1.321 ± 0.04 1.66 ± 0.78 1.52 ± 1.18 0.46 ± 0.26 60 2.31 ± 1.66 1.07 16.79 ± 2.12 10.73 ± 0.58 3.04 ± 1.77 1.43 ± 0.78 3.19 ± 1.79 1.31 ± 1.20 90 1.76 ± 0.51 1.12 ± 0.50 9.48 ± 3.56 7.45 ± 5.88 2.75 ± 1.38 1.73 ± 0.42 2.92 ± 1.24 3.46 ± 1.87 120 1.21± 0.51 0.11 ± 0.08 3.03 ± 2.26 2.64 ± 0.03 2.72 ± 1.69 1.42 ± 0.18 2.18 ± 1.68 2.71 ± 2.82 pp38:p38LPS (100ng/mL, min) pERK1/2:ERK1/2 pJNK:JNK pAkt:Akt 64  to the degradation of IκB, which normally binds NF-κB in the cytoplasm to inhibit nuclear translocation (120), and LPS-induced NF-κB activation requires degradation of IκBα in endothelial cells (232). To confirm that Gαi/o proteins do not contribute to activation of the canonical NF-κB pathway, LPS-induced IκBα degradation was monitored in the presence or absence of PTx. After 60 min of LPS stimulation, we observed a decrease in IκBα protein levels, which was maintained for up to 2 h. This reduction in protein levels was unaffected by pretreatment with PTx (Figure 3.4B). Taken together, these results indicate that Gαi/o proteins do not regulate activation of the canonical NF-κB pathway. This suggests that LPS signaling feeds through both Gαi/o-dependent and MyD88-dependent pathways for full activation of ERK1/2, JNK and Akt, but not NF-κB or p38.               65             Figure 3.4 Heterotrimeric Gαi/o proteins are not important for LPS-induced NF-κB activation in endothelial cells  (A) HMEC were co-transfected with an NF-κB-Luc construct and a constitutively active Renilla luciferase plasmid. Cells were pretreated with PTx (100 ng/mL, 1 h) and stimulated with LPS (100 ng/mL) for the times indicated. (B) HMEC were pretreated with PTx (100 ng/mL, 1 h) and stimulated with LPS (100 ng/mL) for the times indicated. Cell lysates were subjected to SDS-PAGE and immunoblotted with an antibody directed against IκBα. Anti- tubulin was used as a control for protein loading. Error bars indicate s.d., n = 3 independent experiments.   66  3.2.6 Gαi/o proteins do not function along a TRAF6-dependent pathway in endothelial cells Since PTx does not affect NF-κB activation in endothelial cells, we hypothesized that Gαi/o proteins do not act upstream of TRAF6 as part of the MyD88-dependent signaling pathway. Upon TLR4 activation, TRAF6 forms a complex with the ubiquitin conjugating enzyme Ubc13 and the Ubc-like protein Uev1A, to catalyze the formation of a polyubiquitin chain linked through lysine 63 (K63) of ubiquitin (115). Therefore, to investigate the effect of inhibition of Gαi/o on TRAF6 ubiquitination, HMEC were pretreated with PTx, followed by stimulation with LPS, and TRAF6 was immunoprecipitated from cell lysates for immunoblotting with an antibody specific to ubiquitin. After 30 min of LPS stimulation, a ladder of polyubiquitinated TRAF6 was seen in vehicle control cells and the presence of polyubiquitinated TRAF6 was unaffected by pretreatment with PTx (Figure 3.5A). This suggests that Gαi/o proteins do not act upstream of TRAF6. Thus, Gαi/o proteins must either converge on the signaling pathway downstream of TRAF6 or function in a parallel signaling cascade. To further explore at which point in the signaling cascade G-proteins converge, we used concurrent expression of dominant negative TRAF6 and inhibition of Gαi/o proteins. To determine whether Gαi/o act downstream of TRAF6, we expressed the C-terminal fragment of TRAF6 (TRAF6ΔN), (Figure 3.5B), and assessed MAPK activation in the presence or absence of PTx. TRAF6ΔN has been shown to act as a dominant-negative molecule to inhibit signaling  67    Figure 3.5 Gαi/o proteins do not function along the TRAF6 signaling cascade  (A) HMEC were pretreated with PTx (100 ng/mL, 1 h) and stimulated with LPS (100 ng/mL) for 30 min. Whole-cell lysates were prepared and immunoprecipitated with anti-TRAF6 antibody under denaturing conditions followed by immunoblotting with anti-ubiquitin and anti-TRAF6 antibodies. IP, immunoprecipitation; IB, immunoblotting. HMEC were transduced with the dominant negative LNCX-TRAF6ΔN or vector alone. (B) Expression of LNCX-TRAF6ΔN was confirmed by immunoblotting of cell lysates. Tubulin was used as a control for protein loading. (C) Cells were pretreated with PTx (100 ng/mL, 1 h) and stimulated with LPS (100 ng/mL) for the times indicated. Activation of MAPK and Akt was measured using phospho-specific antibodies to JNK, ERK1/2, p38 and Akt. Total MAPK and Akt was used as a positive control for protein loading. Error bars indicate s.d., n ≥ 3 independent experiments. 68   downstream of TRAF6 in IL-1R and TLR signaling (102, 233). Furthermore, our laboratory has previously shown that JNK and Akt, but not ERK1/2 nor p38, are activated downstream of TRAF6 in endothelial cells (128, 234). As expected, activation of JNK and Akt is decreased by both TRAF6ΔN and PTx, but not synergistically by both modes of pathway inhibition (Figure 3.5C). In contrast, ERK1/2 activation is unaffected by the expression of TRAF6ΔN and almost completely abolished by PTx (Figure 3.5C) Together, with data from our laboratory showing that TRAF6ΔN blocks NF-κB activation in endothelial cells (115), these results support a model in which Gαi/o signaling converges downstream of TRAF6 to activate JNK and Akt, while Gαi/o mediated ERK1/2 activation remains independent of the TRAF6 pathway. 2.2.7 Gαi/o proteins contribute to TLR2 and TLR3 signaling Since Gαi/o proteins are activated downstream of TLR2 and TLR3, the ability of Gαi/o proteins to modulate MAPK activation mediated by these other TLRs was assessed. To evaluate the role of Gαi/o in TLR2 signaling, the synthetic TLR2 agonist, Pam3CSK4, was used to stimulate endothelial cells in the presence or absence of PTx. Comparable to the results seen with LPS, pretreatment with PTx decreased ERK1/2 and Akt activation in response to TLR2 agonist (Figure 3.6A). In contrast to TLR4 signaling, Gαi/o proteins mediate Pam3CSK4-induced activation of p38, but not JNK activation (Figure 3.6A, Table 3.1). These results demonstrate that Gαi/o proteins mediate MAPK signaling downstream of TLR2. MyD88 is required for signaling downstream of all TLRs with the exception of TLR3 (235). In contrast to TLR2 and TLR4, TLR3 signaling is initiated solely through receptor 69  binding of the adaptor molecule TRIF, leading to activation of NF-κB, interferon regulatory factor 3 (IRF3) and the MAPKs. The above results suggest that Gαi/o signaling occurs parallel to MyD88 activation, thus to investigate if MAPK activation downstream of TLR3, a MyD88-independent receptor, is also dependent on Gαi/o proteins, HMEC were stimulated with Poly(I:C) in the presence or absence of PTx. Stimulation with Poly(I:C) resulted in activation of ERK1/2 and Akt (Figure 3.6B, Table 3.2) and this activation was reduced in the presence of PTx (Figure 3.6B). Activation of JNK and p38 by Poly(I:C) was weak in HMEC and unable to be assessed (data not shown). These results implicate Gαi/o proteins in signaling via a MyD88-independent pathway and suggest that G-proteins may be universally involved in TLR signaling.        70      Figure 3.6 PTx-sensitive Gαi/o are important for endothelial TLR2 and TLR3 signaling  (A-B) HMEC were pretreated with PTx (100 ng/mL, 1 h) and stimulated with (A) Pam3CSK4 (100 ng/mL), (B) Poly(I:C) (2 μg/mL) for the times indicated. Activation of the MAPKs and Akt was assessed using antibodies specific to phosphorylated forms of ERK1/2, JNK, p38 and Akt. Total Akt or MAPK protein was used as a loading control.  71  Table 3.2 Densitometry analysis of PTx-sensitive Gαi/o endothelial TLR2 signaling.    *Data are represented as relative quantification of phospho-protein:total-protein and expressed as the fold-change relative to untreated . Error indicates s.d., n ≥ 2 independent experiments.                       -PTx +PTx -PTx +PTx -PTx +PTx -PTx +PTx 0 1 0.27 ± 0.14 1 0.87 ± 0.02 1 0.44 ± 0.24 1 1.27 ± 0.46 30 1.54 ± 0.23 0.58 ± 0.08 1.03 ± 0.13 1.97 ± 0.68 1.34 ± 0.61 0.47 ± 0.34 4.17 ± 3.38 1.42 ± 0.29 45 1.33 ± 0.02 0.63 ± 0.08 4.24 ± 1.70 4.91 ± 3.53 1.49 ± 0.62 0.41 ± 0.28 3.73 ± 2.59 1.54 ± 0.73 60 1.04 ± 0.14 0.72 ± 0.11 4.63 ± 2.74 5.33 ± 5.99 1.60 ± 0.60 0.52 ± 0.42 2.71 ± 1.35 1.33 ± 0.35 90 0.91± 0.22 0.79 ± 0.08 1.71 ± 0.55 3.27 ± 2.89 1.30 ± 0.27 0.62 ± 0.40 1.50 ± 0.54 1.44 ± 0.94 120 0.62 ± 0.16 0.64 ± 0.07 1.06 ± 0.01 1.96 ± 1.65 1.30 ± 0.43 1.00 ± 0.34 1.47 ± 0.18 1.66 ± 1.02 pp38:p38Pam3CSK4 (100ng/mL, min) pERK1/2:ERK1/2 pJNK:JNK pAkt:Akt 72  Table 3.3 Densitometry analysis of PTx-sensitive Gαi/o endothelial TLR3 signaling.   *Data are represented as relative quantification of phospho-protein:total-protein and expressed as the fold-change relative to untreated . Error indicates s.d., n ≥ 2 independent experiments.   3.2.8 PI3K is upstream of Akt and JNK in TLR4 signaling Several studies have suggested that PI3Kβ is activated downstream of GPCRs (219, 220) and PI3K has been shown to be upstream of JNK activation in LPS-stimulated neutrophils (134). To determine if PI3K lies upstream of JNK in endothelial cells stimulated with LPS, HMEC were pretreated with the PI3K inhibitor, LY294002, and stimulated with LPS for the indicated times. Indeed, LPS-induced JNK activation was significantly decreased by inhibition of PI3K (Figure 3.7B), suggesting that JNK activation lies downstream of PI3K in endothelial TLR4 signaling.   -PTx +PTx -PTx +PTx 0 1 0.52 ± 0.36 1 0.93 ± 0.46 30 0.99 ± 0.12 0.31 ± 0.03 1.74 ± 0.26 0.91 ± 0.11 45 0.81 ± 0.65 0.46 ± 0.21 2.41 ± 0.19 1.14 ± 0.20 60 1.46 ± 0.54 0.65 ± 0.09 2.07 ± 0.81 1.34 ± 0.23 90 1.31± 0.14 0.37 ± 0.11 1.48 ± 0.84 1.37 ± 0.46 120 1.31 ± 0.66 1.77 ± 0.26 1.46 ± 0.96 0.79 ± 0.09 Poly(I:C) (2μg/mL, min) pERK1/2:ERK1/2 pAkt:Akt 73         -          Figure 3.7 PI3K contributes to LPS-induced JNK activation  (A-B) HMEC were pretreated with LY294002 (50 μM, 1 h) and stimulated with LPS (100 ng/mL) for the times indicated. Activation of (A) Akt and (B) JNK was assessed using antibodies specific to phosphorylated forms of JNK and Akt. Total JNK or Akt protein was used as a loading control. Data is representative of three independent experiments.     74  3.2.9 TLR4 interacts with, and functions through, Gαi2 to activate JNK There is emerging evidence that single transmembrane spanning receptors couple to heterotrimeric G proteins (236). Such nonclassical interactions have been demonstrated for receptor tyrosine kinases, epidermal growth factor receptor and the insulin and insulin-like growth factor receptors (236). Insulin-dependent lipolysis and inhibition of glucose oxidation are blocked by PTx (237) and heterotrimeric G-proteins have been shown to physically associate with, and be activated, by the insulin receptor (238, 239). Co-immunoprecipitation was used to show that TLR4 binds to Gαi2 and thus may also act as a single transmembrane spanning GPCR (Figure 3.8A). Indeed, overexpression of Gαi2 in HEK293-TLR4-MD2- CD14 expressing cells resulted in an increase in LPS-induced activation of JNK (Figure 3.8B). However, a decrease in LPS-induced JNK activation using lentiviral-mediated RNA interference using short hairpin RNA (shRNA) targeting Gαi2 alone was not detected in HMEC (Figure 3.8C). Together, these findings suggest that TLR4 binds to and signals through Gαi2.          75       Figure 3.8 TLR4 binds to and signals through Gαi2  (A) HEK293T cells were co-transfected with Flag-TLR4, Gαi2-GFP or vector control and lysates immunoprecipitated (IP) and immunoblotted (IB) using anti-GFP or anti-Flag. (B) HEK293-TLR4-MD2-CD14 cells were transfected with Gαi2-GFP or vector control. Expression of Gαi2-GFP was confirmed by IB and JNK activation following LPS stimulation (100 ng/mL) was measured for the times indicated. (C) HMEC were transduced with lentiviral vectors encoding shRNA as indicated and Gαi2 was measured by RT-PCR. Activation of JNK following LPS stimulation (100 ng/mL) was measured for the times indicated. Data is representative of two independent experiments.   76  3.3 DISCUSSION  There is increasing evidence demonstrating cross-talk between TLRs and signaling pathways, such as transforming growth factor β (TGFβ) (240, 241), β2 integrins (87), and nuclear receptors (242). Furthermore, collaboration between TLR4 and heterotrimeric Gαi/o protein signaling has been described in cells of the immune system (156-158, 243). The data in this chapter demonstrates for the first time that Gαi/o proteins are activated by LPS in endothelial cells (Figure 3.1) and mediate activation of MAPKs downstream of multiple TLRs in endothelial cells. (Figures 3.2 and 3.6) This data also shows that Gαi/o proteins play a role in endothelial TLR signaling through a pathway that is distinct from the canonical MyD88-dependent pathway. The finding that Gαi/o proteins contribute to TLR4-mediated ERK1/2 and JNK activation, but not p38, is in contrast to the ubiquitous involvement of Gαi/o proteins in MAPK activation in immune cells (156, 157). The activation of PI3K/Akt downstream of TLRs has not been extensively studied. The data in this chapter shows that inhibition of Gαi/o is sufficient to reduce the level of Akt activation back to basal levels in response to ligands for TLR2, TLR3 and TLR4 (Figures 3.3 and 3.6). These results show that Gαi/o proteins are important mediators of multiple signaling events downstream of TLRs and highlights the importance of cell type specific modulation of TLR signaling. The data in Figure 3.4 shows that Gαi/o do not mediate activation of the canonical NF-κB pathway, suggesting that Gαi/o proteins do not play a role upstream of TRAF6. Furthermore, inhibition of Gαi/o in cells expressing dominant negative TRAF6ΔN does not augment the inhibitory effect seen by either PTx or TRAF6ΔN alone (Figure 3.5). Finally, pretreatment of endothelial cells with PTx does not inhibit TRAF6 ubiquitination (Figure 77  3.5). Taken together, these results reveal that Gαi/o proteins converge on the signaling pathway downstream of the MyD88/IRAK/TRAF6 signaling cascade. The family of TLRs signal through various adaptor proteins (81). TLR2 engages MyD88/TIRAP, TLR3 engages TRIF alone, whereas TLR4 is coupled to all of MyD88/TIRAP and TRIF/TRAM (81). These results suggest that Gαi/o proteins may act through a common mechanism to initiate signaling events downstream of diverse TLR signaling pathways. Fan et al. (158) have found that Gαi proteins are important for signaling events initiated by both gram-positive and gram-negative bacteria. This data supports our finding that Gαi/o proteins are important for signaling from multiple TLRs. The TLR homology domain, TIR, is common to all TLRs and the IL-1R family members, IL-1R, IL- 18R and ST2 (244). IL-1 stimulation has been shown to increase GTP binding to cellular membranes and promote GTP hydrolysis (245). We have also seen that PTx can inhibit IL- 1R-mediated signaling to Akt, but not the MAPKs (data not shown). The data presented here shows that PI3K/Akt activation is important for JNK activation downstream of TLR4. This agrees with the finding that inhibition of PI3K reduces Syk-dependent JNK activation in LPS-stimulated neutrophils (134). Gαi proteins activate PI3K by mechanisms involving the dissociation of Gβγ subunits from the G-protein heterotrimer. In contrast to JNK activation, we have seen in our laboratory that activation of ERK1/2 is not reduced by PI3K inhibitors (data not shown). Similarly, LPS-induced ERK1/2 activation is not affected by expression of a Gβγ sequestering peptide in HEK293 cells (157). As shown in Figure 3.8, TLR4 may couple to Gαi2 to induce G protein mediated JNK activation. Indeed, there is emerging evidence that single transmembrane spanning receptors, such as the insulin receptor, bind to Gαi/o to activate downstream signaling (236). 78  However, the finding that RNA interference of a single Gαi isoform is not sufficient to abrogate signaling may suggest that TLR4 is able to bind other Gαi proteins, and together, these isoforms may exhibit functional redundancy during LPS activation. Indeed, Fan et al. have seen that transfection of a dominant negative for a single Gαi subunit is not sufficient to decrease ERK1/2 activation, but rather co-transfection of dominant negative constructs for atleast two Gαi subunits is required for inhibition of the signal (157). We propose that since Gαi2 can be immunoprecipitated with TLR4 that the difference between immune and endothelial cells may be independent of a discrepancy between mCD14 and sCD14, atleast for G-protein activation. Whether TLR4 binds to and signals through other Gαi isoforms remains to be determined and will be discussed further in chapter 5. The data presented in this chapter leads to a novel model of LPS-induced activation of pertussis toxin sensitive G-proteins, and the subsequent activation of ERK, JNK and Akt, in endothelial cells. (Figure 3.9).                   79                                   Figure 3.9 A model for Gαi/o in TLR4 signaling Gαi/o, activated downstream of TLR4, contribute to ERK1/2, JNK and Akt activation, along a pathway that is parallel to the MyD88-dependent signaling cascade and eventually converges downstream of TRAF6 for activation of JNK and Akt. TLR4, Toll-like receptor 4; MyD88, myeloid differentiation factor 88; IRAK, interleukin-1 receptor-associated kinase; TRAF6, tumor necrosis factor receptor-associated factor 6; NF-κB, nuclear transcription factor-κB; JNK, c-Jun NH2-terminal kinase; ERK1/2, extracellular signaling regulated kinase 1/2.    Gαi/o TLR4 MyD88 IRAK NF-κB JNK   PI3K ERK   Akt p38 TRAF6  80           CHAPTER 4: THE IDENTIFICATION OF SASH1 AS A NOVEL TLR4 SIGNALING MOLECULE              81  4.1 INTRODUCTION Our laboratory has previously shown that FADD is a negative regulator of TLR4 signaling in endothelial cells and FADD null cells demonstrate hyperactivation of JNK in response to LPS (160). In addition, LPS signaling components have been shown to localize to lipid-rich microdomains in the plasma membrane (236) and unpublished work from our laboratory has shown that disruption of lipid raft microdomains inhibits LPS-induced JNK activation (Zhande et al., unpublished). We hypothesized that the hyperactivation of the LPS signal in FADD null cells is due, in part, to an alteration in lipid raft microdomain protein content. Thus, to identify proteins that are differentially expressed in the lipid rafts (or more specifically, the DRM fraction) of FADD wild-type (WT) versus FADD knock-out (KO) cells, mass spectrometric (MS) analysis was used. This resulted in the identification of SAM and SH3 domain-containing protein (SASH1) as a putative candidate for positive regulation of TLR4 signaling. SASH1 belongs to a family of SAM and SH3 adaptor proteins, consisting of SH3 domain expressed in lymphocytes (SLY1), hematopoietic adaptor containing SH3 and SAM domains 1 (HACS1, also called SLY2) (218-221), which have been implicated in immune regulation, suggesting that SASH1 may also function in the immune system. However, the molecular function of SASH1 has not been studied. The data in this chapter reveals that SASH1 positively regulates signaling through TLR4 to increase activation of NF-κB and the MAPKs, culminating in the increased production of proinflammatory cytokines. Further, SASH1 binds to TRAF6 and regulates its ubiquitination through a conserved TRAF6 binding domain within SASH1. SASH1 also binds to TAK1 and the IKK complex to assemble a signaling hub at TRAF6, thus permitting 82  downstream signaling. Collectively, these results demonstrate that SASH1 functions as a novel scaffold protein to regulate innate immune signaling. Although signaling complexes are often assembled on large proteins that act as scaffolds to regulate signal transduction, identification of a scaffold protein for coordination of TLR signaling complexes has remained elusive (203). In this chapter, a novel scaffold protein in TLR signaling, SASH1, is identified and characterized.                  83  4.2 RESULTS 4.2.1 Proteomic analysis identifies SASH1 as a putative player in the TLR4 signaling pathway The plasma membrane is a disordered bilayer, originally thought to consist of a random distribution of proteins and lipids (246). However, within this bilayer, there are specialized lipid-rich microdomains, referred to as lipid rafts, which serve as a platform for organizing cellular signaling (247). TLR4 has been shown to localize to the lipid rafts of both immune and endothelial cells following stimulation with LPS (248, 249). We have previously found that FADD is a negative regulator of IRAK1/MyD88-dependent responses and we have also seen that mouse embryonic fibroblast (MEF) FADD-KO cells exhibit increased activation in response to LPS (173). Thus, FADD-KO cells can be used as a model to identify novel proteins important for TLR4 signaling. To examine the protein content of lipid microdomains from FADD-WT versus FADD-KO, DRMs were isolated from these cells and a differential proteomic analysis performed. This analysis revealed two peptides that uniquely identified SASH1 as present in the DRM fraction of FADD-KO, but not FADD-WT, cells (Figure 4.1A-C). The presence of SASH1 in the DRM fraction of FADD- KO cells was validated by Western blot analysis (Figure 4.1D). Together, this finding suggests that SASH1 is a putative candidate for positive regulation of the TLR4 pathway.  84   Figure 4.1 Identification of SASH1 in MEF FADD-KO cells by mass spectrometry analysis  MEF FADD-WT and FADD-KO were stimulated with LPS (100 ng/mL, 5 min) and DRMs were isolated. (A) Protein fractions isolated from sucrose density-gradient separation were immunoblotted with caveolin-1 to validate the separation procedure. (B) SASH1 peptides identified in MEF FADD-KO DRMs. (C) MS/MS spectrum assigned to one of the SASH1 peptides predicting the amino acid sequence of FIYVDVLNEEEEK. The y and b ions are labeled. (D) Validation of SASH1 in the MEF FADD KO DRM fraction.  85  4.2.2 SASH1 positively regulates LPS signal transduction  To demonstrate that SASH1 is important for LPS-induced signal transduction, we used HEK293-TLR4-CD14-MD2 cells overexpressing SASH1. Ectopic expression of SASH1 (Figure 4.2A) resulted in an increase in the phosphorylation and activation of JNK (Figure 4.2B). Furthermore, increased expression of SASH1 in HMEC, resulted in an increase in NF-κB luciferase reporter activity (Figure 4.2C). Activation of NF-κB results in the production of proinflammatory cytokines, therefore, we wanted to determine whether the increase in activation of NF-κB would result in a concomitant increase in production of the proinflammatory cytokines, IL-6 and IP-10 (250). As seen in Figure 4.2D, overexpression of SASH1 resulted in an increase in the production of IL-6, in response to LPS, as compared to vector control. Similarly, LPS-induced IP-10 production was also increased by ectopic expression of SASH1 (Figure 4.2E). Conversely, lentiviral mediated knockdown using short hairpin RNA (shRNA) targeting SASH1 in HMEC resulted in a decrease in activation of JNK and NF-κB, and reduced IL-6 production from LPS-stimulated HMEC (Figure 4.3A-D). 86           Figure 4.2 SASH1 promotes endothelial TLR4 signaling  (A-B) HEK293 cells expressing TLR4-MD2-CD14 were transfected with Flag-SASH1 or vector control. (A) Expression of Flag-SASH1 was confirmed by immunoblotting and (B) JNK activation following LPS stimulation (100 ng/mL) was measured by immunoblotting. (C-D) HMEC were co-transfected with Flag-SASH1 or vector control and (C) NF-κB-Luc plus pRL-CMV for measurement of reporter activity following LPS stimulation (100 ng/mL, 8 h), or (D-E) stimulated with LPS  (100 ng/mL, 6 h) and (D) IL-6 or (E) IP-10 in the supernatants was measured by ELISA. Data are expressed as the fold-change relative to normal. *P < 0.05 as determined by Student’s t-test. Error bars indicate s.d., n = 3 independent experiments.   87                Figure 4.3 SASH1 knockdown decreases LPS signaling  (A-D) HMEC were transduced with lentiviral vectors encoding shRNA as indicated, and (A) SASH1 was measured by RT-PCR. (B) Activation of JNK following LPS stimulation (100 ng/mL) was measured by immunoblotting or (C) HMEC were transfected with pNF-κB-Luc plus pRL-CMV for measurement of reporter activity following LPS stimulation (100 ng/mL, 8 h) or (D) stimulated with LPS  (100 ng/mL, 6 h) and IL-6 in the supernatants was measured by ELISA. Data are expressed as the fold-change relative to normal. *P < 0.05 as determined by Student’s t-test. Error bars indicate s.d.  88  4.2.3 SASH1 is not important for LPS-induced IFN signaling TLR4 is coupled to the activation of two intracellular signaling pathways: a MyD88- dependent pathway leading to activation of NF-κB and the MAPKs, and a MyD88- independent pathway leading to the activation of late-phase NF-κB, MAPKs and IFN- inducible genes (81). The results shown above suggest that SASH1 is important for the activation of NF-κB and JNK. Therefore, to determine if SASH1 was also able to promote activation of IFN-inducible genes, an interferon-stimulated responsive element (ISRE) coupled to luciferase was used. The data in Figure 4.4 shows that SASH1 does not increase activation of ISRE-luciferase, suggesting that SASH1 is important for activation of NF-κB and JNK, but not for the activation of IFN-regulated genes downstream of TLR4. 4.2.4 SASH1 plays a role in TLR3, but not TLR2, signaling TLR4 belongs to a family of TLRs that recognize a diversity of ligands from a wide variety of pathogens (see Table 1.1) (55). Thus, to investigate if SASH1 is important for signaling downstream of other TLRs, the role of SASH1 in NF-κB activation following activation of TLR2 and TLR3 was examined. The data indicates that SASH1 does not function downstream of TLR2 (Figure 4.5A), a MyD88-dependent pathway, but does increase signaling to NF-κB from TLR3 (Figure 4.5B), a MyD88-independent pathway, as determined by stimulation with Pam3CSK4 and Poly(I:C), respectively. These results suggest that SASH1 may be important downstream of the adaptor molecule, TRIF, leading to activation of NF-κB.      89                     Figure 4.4 SASH1 does not play a role in signaling to the IFN pathway  HMEC were co-transfected with Flag-SASH1 or vector control and pISRE-Luc plus pRL-TK for measurement of reporter activity following LPS stimulation (100 ng/mL, 8 h). Data are expressed as the fold-change relative to normal. Error bars indicate s.d., n = 3 independent experiments.    90        Figure 4.5 SASH1 promotes endothelial TLR3, but not TLR2, signaling  (A-B) HMEC were transfected with Flag-SASH1 or vector control and pNF-κB-Luc plus pRL-CMV for measurement of reporter activity following stimulation with (A) Pam3CSK4 (100 ng/mL, 8 h) or (B) Poly(I:C) (2 μg/mL, 8 h). Data are expressed as the fold-change relative to normal. *P < 0.05 as determined by Student’s t-test. Error bars indicate s.d., n = 3 independent experiments.    91  4.2.5 SASH1 interacts with the C-terminal domain of TRAF6 TRAF6 binding proteins, RANK and IRAK, possess a conserved TRAF6 interaction motif (103, 251). To determine if SASH1 contains a putative TRAF6-binding site, in silico analysis was performed. This analysis revealed that the SASH1 protein sequence contains the consensus site for TRAF6 binding at amino acid 852-860 of the human SASH1 sequence. To validate this finding, co-immunoprecipitation was used to show that SASH1 interacts with TRAF6 (Figure 4.6A). To determine if this binding is physiologically relevant, interaction between endogenous TRAF6 and SASH1 was examined in LPS-stimulated HEK293-TLR4-CD14-MD2. Immunoprecipitation of TRAF6 resulted in co-precipitation of endogenous SASH1 after 5 min, and up to 60 min of LPS treatment (Figure 4.6B), suggesting that this interaction is stimulus-dependent. To determine if SASH1 binds to TRAF6 through the putative TRAF6-binding domain identified by in silico analysis, a SASH1 mutant lacking this domain was made. Deletion of this domain abolished interaction with SASH1 (Figure 4.6C), suggesting that interaction does occur through the conserved TRAF6 binding motif. Other TRAF6 binding partners that possess a conserved TRAF6 interaction motif, interact with TRAF6 at the C-terminal TRAF domain (106). Indeed, SASH1 interacts with the C-terminus of TRAF6, near the conserved TRAF domain (Figure 4.7A and B).      92            Figure 4.6 SASH1 interacts with TRAF6 through a conserved TRAF6 binding motif  (A) HEK293T cells were co-transfected with HA-SASH1, Flag-TRAF6 or vector control and lysates immunoprecipitated (IP) and immunoblotted (IB) using anti-HA or anti-Flag. (B) Endogenous SASH1 associates with TRAF6 in HEK293-TLR4-MD2-CD14 cells stimulated with LPS (100 ng/mL, times indicated in min). (C) HEK293T cells were co-transfected with Flag-TRAF6 and Myc-SASH1ΔT6BD and IP and IB with anti-TRAF6 or anti-SASH1. 93     Figure 4.7 SASH1 interacts with the C-terminus of TRAF6  (A) Schematic of TRAF6 deletion constructs. TRAF6 aa 1-522, Flag-TRAF6ΔN aa 289-522, Flag-TRAF6ΔC aa 1-289, Flag-TRAF6ΔCC aa 1-292, 351-522. (B) HEK293T cells were co- transfected with HA-SASH1 and either Flag-TRAF6ΔN, Flag-TRAF6ΔC or Flag-TRAF6ΔCC and lysates were immunoprecipitated (IP) and immunoblotted (IB) using antibodies to HA or Flag. CC, coiled coil; aa, amino acid. 94   4.2.6 SASH1 regulates TRAF6 ubiquitination K63-linked autoubiquitination of TRAF6 is critical for formation of the signaling complex comprised of TAK1, and the adaptor proteins, TAB2 and TAB3 (107, 112, 252). To determine if the interaction between SASH1 and TRAF6 regulates TRAF6 activation, TRAF6 ubiquitination was assessed in the presence of SASH1. Indeed, ectopic expression of SASH1 in HEK293-TLR4-CD14-MD2 was sufficient to induce autoubiquitination of TRAF6 in the absence of LPS stimulation (Figure 4.8A). Moreover, expression of the TRAF6 binding domain mutant did not increase ubiquitination of TRAF6 (Figure 4.8B) or the downstream activation of an NF-κB luciferase reporter (Figure 4.8C). These results imply that binding of SASH1 to TRAF6 is critical for signaling downstream of TLR4. However, mutation of the SASH1 binding domain does not inhibit LPS-induced signaling events below the level of vector control cells, suggesting that the TRAF6 binding mutant does not act as a dominant negative. TRAF6 ubiquitination requires interaction with a heterodimeric ubiquitin conjugating enzyme composed of Ubc13 and the Ubc variant, Uev1A (115). TRAF6 has been shown to bind directly to Ubc13, but not Uev1A (253). An interaction between SASH1 and Ubc13 was not detected in unstimulated cells (Figure 4.9A), although we were able to detect interaction of TRAF6 and Ubc13 (Figure 4.9B). Whether SASH1 increases the basal interaction between TRAF6 and Ubc13 or binds directly to Uev1A, is unknown. Thus, it remains unclear how SASH1 facilitates TRAF6 ubiquitination in the absence of stimulation and it is likely that a secondary signal through LPS-TLR4 engagement is required to complete the activation cascade to NF-κB and JNK.  95            Figure 4.8 SASH1 regulates TRAF6 ubiquitination  (A) HEK293-TLR4-MD2-CD14 cells were transfected with Flag-SASH1 or Flag-TRAF6, stimulated with LPS (100 ng/mL, 30 min), IP for TRAF6 and IB with anti-ubiquitin, TRAF6 or SASH1. (B) HEK293-TLR4-MD2-CD14 cells were transfected with Flag-SASH1, Myc-SASH1ΔT6BD or vector control, stimulated with LPS (100 ng/mL, 30 min) and IP for TRAF6 and IB with anti-ubiquitin, anti-TRAF6 or anti-SASH1. (C) HMEC were co- transfected with Flag-SASH1, Myc-SASH1ΔT6BD or vector control and pNF-κB-Luc plus pRL-CMV for measurement of reporter activity following stimulation with LPS (100 ng/mL, 8 h). *P < 0.001 as determined by Student’s t-test. Data are presented as fold-change relative to the normal. Error bars indicate s.d., n = 3 indpendent experiments.   96          Figure 4.9 SASH1 does not interact with Ubc13  HEK293T cells were co-transfected with (A) HA-SASH1 and Flag-Ubc13 or (B) HA-TRAF6 and Flag-Ubc13 and lysates were immunoprecipitated (IP) and immunoblotted (IB) with antibodies to HA and Flag.   97  4.2.7 SASH1 does not interact with other TRAF molecules TRAF6 belongs to a family of TRAF molecules that mediate signaling downstream of TNFR superfamily members (104). In silico analysis also identified four putative TRAF2 binding sites within the human SASH1 protein sequence. TRAF2 is critical for the activation of NF-κB and JNK downstream of numerous receptors, including TNFR1 and CD40 (254). The presence of multiple putative TRAF2 binding sites within SASH1 could be indicative of a broader role for SASH1 in regulating TRAF family members. However, co- immunoprecipitation analysis revealed that SASH1 does not interact with TRAF2 (Figure 4.10A). TRAF3 functions downstream of TLR4 in MyD88-independent pathways, primarily contributing to the activation of IFN-dependent genes (166). To determine if SASH1 interacts with TRAF3, co-immunoprecipitation was used. However, no interaction was apparent between SASH1 and TRAF3 (Figure 4.10B), The absence of an interaction between SASH1 and TRAF3 agrees with the finding that SASH1 does not induce ISRE-luciferase, supporting a model in which SASH1 promotes activation of NF-κB and JNK, but not IFN- dependent pathways. Taken together, these results suggest that SASH1 may interact specifically with TRAF6, and not with other TRAF molecules.       98             Figure 4.10 SASH1 does not interact with TRAF2 or TRAF3  (A-B) HEK293T cells were co-transfected with HA-SASH1 and either  (A) Flag-TRAF2or (B) Flag-TRAF3 and lysates were immunoprecipitated (IP) and immunoblotted (IB) using antibodies to HA or Flag.    99  4.2.8 SASH1 acts as a scaffold molecule by binding TAK1, IKKα and IKKβ The coordination of TLR signaling cascades by scaffold proteins has not been extensively studied. Scaffold proteins have been defined as proteins that bind to two or more molecules within a signaling cascade to regulate signaling, either by altering subcellular localization of the complex, coordinating positive and negative regulators or protecting the complex from inactivation (203). Given that SASH1 is a large protein with multiple protein-interaction domains, suggestive of a scaffolding function, co-immunoprecipitation was used to determine whether SASH1 provides the framework for constructing a molecular complex around TRAF6. TAK1 binds to TRAF6 through the adaptor molecules, TAB1/2/3, thereby facilitating activation of TRAF6 (100). Co-immunoprecipitation was used to show that SASH1 interacts with TAK1 (Figure 4.11A). TAK1 phosphorylates and activates the downstream target IKKβ, leading to NF-κB activation (118). Therefore, the interaction between SASH1 and IKKβ was examined by co-immunoprecipitation. Indeed, SASH1 also binds to IKKβ (Figure 4.11B). IKKβ is part of a complex consisting of an additional catalytic subunit, IKKα, and a regulatory subunit, IKKγ (119). The data in Figure 4.11 shows that SASH1 also interacts with IKKα, but not the regulatory subunit IKKγ (Figure 4.11C-D). Whether SASH1 interacts with IKKγ following LPS stimulation remains to be determined. To determine if the interaction between SASH1 and TAK1/IKK is dependent on the interaction with TRAF6, the TRAF6 binding domain mutant was used. Interestingly, interaction of SASH1 with either TAK1 or IKKβ  was not abolished by deletion of the TRAF6 binding domain (Figure 4.12E-F), suggesting that SASH1 can bind to TAK1 and IKKβ in the absence of an interaction between SASH1 and TRAF6. Collectively, these  100    Figure 4.11 SASH1 is a scaffold protein that binds to the TAK1-IKK complex (A-D) HEK293T cells were co-transfected with HA-SASH1 and either Flag-TAK1 (A) or Flag-IKKβ (B) or Flag-IKKα (C) or Flag-IKKγ (D) and lysates were immunoprecipitated (IP) and immunoblotted (IB) with antibodies to HA and Flag.      101                          Figure 4.12 The TRAF6 binding mutant of SASH1 binds to TAK1 and IKKβ  (A-B) HEK293T cells were co-transfected with Myc-SASH1DT6BD and either Flag-TAK1 (A) or Flag-IKKβ (B) and lysates were IP and IB with antibodies to Myc and Flag.         102  results suggest that SASH1 acts as a scaffold molecule to bind TRAF6/TAK1/IKK to facilitate signaling to NF-κB and JNK. 4.2.9 Generation of Sash1 gene-trap mice To examine the in vivo role of SASH1 in TLR4 signaling, we aimed to generate Sash1-/- mice using a gene-trap approach that results in a Sash1-LacZ fusion protein under control of the endogenous Sash1 promoter. The Sash1-LacZ fusion disrupts the Sash1 gene at intron 14, truncating the SH3 domain and resulting in loss of the two SAM domains and the putative TRAF6 binding domain (Figure 4.13A). Genotypic analysis of progeny from Sash1+/- crosses did not reveal any viable Sash1-/- mice from 11 litters, with a total of 29 pups, (P < 0.05, chi square test) (Figure 4.13B). Our preliminary observations suggest that Sash1-/- mice die in utero prior to embryonic day (E) 8.5, but further work is needed to delineate the exact age and cause of embryonic lethality. 4.2.10 Sash1 expression in vivo To investigate the in vivo expression pattern of Sash1, tissues were harvested from Sash1+/- mice and analysed for mRNA expression by RT-PCR. Sash1 mRNA was found to be expressed in all mouse tissues examined, with expression predominant in the brain, heart, liver, lung and kidney (Figure 4.14). Insertion of the β-geo cassette into the Sash1 locus allows investigation of gene expression at the cellular level by X-gal staining of tissues from heterozygous mice. Notably, Sash1 was predominantly expressed in the endothelium, but not immune cells, of the spleen and thymus (Figure 4.15A-B).  Sash1 was also strongly expressed in lung endothelial cells (Figure 4.15C-D). CD105 (also known as endoglin) is preferentially expressed on endothelial cells. Sash1 mRNA was  103         Figure 4.13 Generation of gene-trap Sash1 mice Gene-trap mice were generated from ES cells. (A) Genomic insertion was mapped to intron 14-15 using PCR. (B) Genotypic analysis of litters born from heterozygous crosses.     104                       Figure 4.14 Sash1 mRNA is expressed in mouse tissues  RT-PCR was performed on RNA isolated from tissues harvested from C57BL/6 mice using primers specific to mSASH1. GAPDH was used as an internal control.      105          Figure 4.15 Sash1 is expressed predominantly in the endothelium of the spleen, thymus and lung   (A-B) Sash1 expression was detected in the endothelium of the spleen (A), thymus (B) and lung (C) as determined by β-galactosidase activity. (D) Endothelial cells were purified by flow cytometry using anti-mouse CD105 hybridoma and RT-PCR was performed on RNA isolated from these cells using primers specific for Sash1, endothelial markers (CD31, VE- cadherin, vWF), epithelial marker (E-cadherin), Sly1 and Sly2. Gapdh was used as a control.   106  found predominantly in the endothelial, CD105+, fraction of adult lung tissue, while the other members of the SLY family, Sly1 and Sly2, were found predominantly in the non-endothelial, CD105-, population (Figure 4.15D). Sash1 expression was also detected in the parenchyma of the liver, kidney and brain. Furthermore, Sash1 expression was also detected microvasculature of these tissues, as noted  (Figure 4.16A-C). The presence of SASH1 in the endothelium, but not immune cells, could suggest that SASH1 functions as an endothelial-selective TLR4 scaffolding molecule, present in endothelial cells, but not immune cells. Whether SLY family members, SLY1 and SLY2, compensate for the function of SASH1 in immune cells remains to be determined.                    107           Figure 4.16 Sash1 is expressed in the parenchyma and microvasculature of murine liver, kidney and brain  Sash1 expression was detected in the parenchyma of the liver (A, i and ii), kidney (B, i and ii) and brain (C, i and ii) as determined by β-galactosidase activity. Arrows denote (A) hepatic sinusoids, indicative of endothelial staining; (B) renal glomeruli, indicative of microvascular staining and; (C) vascular staining in the brain.  108  4.3 DISCUSSION The data presented in this chapter describes the function of a previously uncharacterized protein, SASH1. SASH1, a member of the SLY family, functions as a novel TLR4 signaling molecule that is expressed in vivo in endothelial cells. These results show that SASH1 acts as a scaffolding molecule to create a molecular complex at TRAF6, leading to the activation of signaling molecules reported to be downstream of this complex in endothelial cells, namely NF-κB and JNK (234). Although SASH1 was found in the DRM fraction of MEFs following LPS stimulation (Figure 4.1), the subcellular localization of SASH1 has not been extensively investigated. Since one of the functions of a scaffold molecule is to direct subcellular localization of molecular complexes, it would be of interest to investigate if the localization of SASH1 changes with LPS stimulation and whether this corresponds to a change in localization of its binding partners, namely TRAF6. Indeed, TRAF6 is found in the membrane proximal region in the early moments following LPS stimulation and translocates into the cytosol as the signal is propagated (255). Recently, the interaction between TRAF6 and TIRAP has been shown to be critical for localization of TRAF6 at the plasma membrane (255). Whether SASH1 acts as a molecular shuttle for TRAF6 remains to be explored. The finding that SASH1 plays a role in activation of NF-κB, but not IFN, suggests that SASH1 is involved in MyD88-dependent, but not MyD88-independent, signaling (Figures 4.2-4.4). However, the data presented herein also indicates that SASH1 is important in the activation of NF-κB downstream of TLR3, a MyD88-independent receptor, but not TLR2, a MyD88-dependent receptor (Figure 4.5). Thus, a potential model for the role of SASH1 in endothelial cells is that SASH1 is important for the activation of 109  TRAF6/TAK1/IKK leading to the activation of NF-κB, but not IFN, downstream of the adaptor TRIF. At this point, we cannot exclude the possibility that SASH1 also plays a role in LPS-induced activation of NF-κB downstream of MyD88. It remains unclear why SASH1 is not important for signaling mediated by TLR2 and it would be of interest to determine if SASH1 is important for NF-κB signaling downstream of other TLRs which are strictly MyD88-dependent, such as TLR5. The conserved TRAF6 binding domain within SASH1 appears critical for interaction with TRAF6 and subsequently downstream signaling (Figure 4.6 and 4.8). Interestingly, deletion of this domain in SASH1 does not affect the binding of SASH1 to other TLR4 signaling molecules, TAK1 and IKKβ, suggesting that SASH1 binds these molecules independent of interaction with TRAF6 (Figure 4.12). Using TRAF6-null cells to investigate the interaction between SASH1/TAK1 and SASH1/IKKβ in the genetic absence of TRAF6 would strengthen this observation. In addition, it remains unknown if SASH1 is critically required for formation of the complex. This will be discussed in more detail in chapter 5. The endothelium plays a critical role in the inflammatory response, through recruitment and transmigration of leukocytes into infected tissue and the regulation of vascular tone (256). The expression of SASH1 in endothelial cells, but not immune cells, may provide an explanation for the cell-type specific responses to LPS and provide insight on the molecular events that lead to endothelial dysfunction and vascular collapse in sepsis. In endothelial cells, LPS activates NF-κB to induce expression of the endothelial- leukocyte adhesion molecule, E-selectin, contributing to neutrophil infiltration in an animal model of systemic inflammation (43). Activation of NF-κB also promotes inducible nitric oxide synthase (iNOS) expression, which, in turn, leads to increased nitric oxide (NO) 110  production, resulting in vasodilation and hypotension (205). In the endothelium, it is these unique consequences of NF-κB signaling that may be regulated at the molecular level by SASH1. Indeed, the importance of endothelial NF-κB signaling in the vascular sequelae of sepsis strengthens a model in which SASH1 functions as an endothelial scaffold molecule for assembly of TRAF6/TAK1/IKK for activation of downstream signaling (Figure 4.17).                   111                    Figure 4.17 A model for the role of SASH1 in endothelial TLR4 signaling Binding of SASH1 to TRAF6/TAK1/IKKβ promotes TRAF6 ubiquitination, resulting in activation of NF-κB and JNK, and culminating in the increased production of proinflammatory cytokines. TLR4, Toll-like receptor 4; MyD88, myeloid differentiation factor 8; TRIF, Toll/interleukin-1 receptor-containing adaptor inducing interferon β; SASH1, SAM and SH3 domain-containing protein; TRAF6, tumor necrosis factor receptor-associated factor 6; TAK1, transforming growth factor β-activated kinase 1; IKK, IκB kinase; NF-κB, nuclear transcription factor-κB; Ub, ubiquitin.  TLR4 NF-κBJNK TRAF6 SASH1TAK1 IKKβ MyD88 TRIF ? ?Ub Proinflammatory cytokines 112          CHAPTER 5: SUMMARY AND FUTURE PERSPECTIVES                  113  5.1 THE STUDY OF TLR SIGNALING  Human TLRs have been studied since the first report of the hToll receptor, later named TLR4, in 1997 (257). Over the past two decades, much work has been done to delineate the signaling mechanisms downstream of this, and other, TLRs. Additionally, the field has also benefited from an understanding of unique signaling cascades triggered downstream of other pattern recognition receptors, such as NLRs and RLRs (57, 258). We are continuously reminded of the complexity of the innate immune system with the discovery of new regulators (ie: SARM) (82), novel splice variants with distinct functions (ie: MyD88s) (199), and unique cell-type specific responses to pathogens (259, 260). These important discoveries contribute greatly to our molecular knowledge of mammalian host defense, cross-talk between TLRs and other signaling cascades, and, how phenomena, such as endotoxin tolerance, develop. A continued exploration of the TLR signaling cascades will expand our understanding of the pathophysiology of numerous inflammatory diseases, thus allowing us to develop targeted therapies with immense clinical benefit for patients suffering from inflammation-associated disease. This thesis identifies novel regulators of the TLR4 signaling cascade, specifically in endothelial cells. Although most attention in the field of innate immunity is focused on immune cells, growing interest in the role of the endothelium in coordinating early inflammatory responses has revealed that endothelial cells are capable of recognizing pathogenic ligands and, thus, may be considered sentinel cells themselves (259). The work presented herein contributes to our understanding of endothelial responses to microbial products, specifically LPS.  114  5.2 HETEROTRIMERIC G PROTEINS IN ENDOTHELIAL TLR SIGNALING The work presented in chapter 3 demonstrates that Gαi/o proteins are important for activation of signaling molecules downstream of multiple TLRs. In particular, Gαi/o mediate endothelial ERK1/2, JNK and Akt activation, but not the activation of p38 or NF-κB in response to LPS. The activation of Gαi/o and their downstream targets occurs along a pathway that is parallel to the MyD88-dependent pathway, converging downstream of TRAF6, and may be common to TLR2, TLR3 and TLR4. Although the data presented in chapter three suggests that TLR4 may bind to Gαi2, it remains unclear how Gαi/o are activated by TLRs in endothelial cells. In addition to the scenario where TLR4 acts as a single transmembrane spanning receptor, it is also possible that a GPCR may cooperate with TLR4, by also binding to LPS to induce cellular activation. In addition to CD14, TLR4 and MD2, Triantafilou et al. have proposed that other receptor molecules may be involved in CD14-independent signal initiation in macrophages (261, 262). Studies using affinity chromatography, and later confirmed by fluorescence resonance energy transfer (FRET), reveal that LPS associates with the heat shock proteins, Hsp70 and Hsp90, chemokine receptor 4 (CXCR4) and growth differentiation factor 5 (GDF) (261, 262). Alternatively, TLR4 activation may result in production of a secreted factor that could function as a GPCR agonist. 5.2.4 TLR4 as a single transmembrane spanning GPCR The data presented in chapter three suggests that TLR4 binds to, and activates, Gαi2. However, lentiviral-mediated RNA interference of Gαi2 alone is not sufficient to decrease LPS-induced JNK activation. Together, this data could suggest that the other Gαi isoforms, Gαi1 and Gαi3, and together, these isoforms may exhibit functional redundancy during LPS 115  activation. Indeed, the highly homologous Gαi isoforms are functionally redundant downstream of α2-adrenoreceptors in vivo (263). Whether TLR4 binds to and signals through other Gαi isoforms in endothelial cells remains unclear. However, it has been shown that genetic deletion of the Gαi isoforms results in a decrease in the production of proinflammatory cytokines in macrophages and splenocytes, suggesting that all three isoforms have signaling capacity downstream of TLR4 (254, 255). RNA interference targeting constructs were designed and tested for both of Gαi1 and Gαi3, but knockdown was not detected in either HEK293T or HMECs using three different constructs for each gene. Achieving efficient knockdown of all three Gαi isoforms will be important for testing the hypothesis of functional redundancy of the Gαi subunits following LPS stimulation. Gαi/o proteins have been shown to bind to a consensus motif consisting of a basic residue at the N terminus and the C-terminal structure of BBXXB or BBXB (where B is any basic residue and X is any non-basic residue) (264). This motif is found in the intracellular portion of TLR2, TLR3 and TLR4. Thus it would be of interest to investigate whether this region of the intracellular TIR domain is responsible for binding and activation of Gαi. TLR4 mutants lacking these domains have been created and will be a useful tool to study whether Gαi proteins do indeed interact at these consensus sites. 5.2.1 Is the LPS-induced Gαi/o-mediated signal dependent on CD14/TLR4? The original identification of TLR4 arose from the discovery that C57BL/10ScCr mice, lacking the TLR4 gene, and C3H/HeJ mice, with a mutation in the TLR4 gene, do not respond to LPS (64). C3H/HeJ and CD14-deficient mice exposed to systemic LPS do not exhibit increased circulating leukocyte counts compared to untreated controls (265) and muscle-derived endothelial cells from C3H/HeJ mice do not show increased expression of P- 116  selectin following LPS stimulation, suggesting that endothelial cells do require CD14/TLR4 for endothelial specific functions, such as leukocyte rolling and adhesion (265). Furthermore, a murine bone marrow-derived macrophage cell line isolated from the mouse strain C57BL/10ScNCr, which has a deletion at the Tlr4 locus, does not increase production of TNFα or MIP2 in response to LPS (266). However, the expression of various inflammatory genes, such as IL-1β, has been shown to be TLR4-independent in dendritic cells (267). Thus, it remains controversial whether CD14/TLR4 is required for cellular activation by LPS in all cell types. Consistent with previous reports, we have seen that LPS-induced MAPK activation does indeed require the CD14 receptor molecule in endothelial cells (72, 74). This may suggest that the Gαi/o-mediated signal is downstream of CD14, and likely TLR4, thus making the study of LPS-induced Gαi/o in cells lacking mCD14 relevant in our understanding of the cell-type specific effects of LPS activation. 5.2.2 Cooperation between TLR4 and a GPCR Whether the CD14/TLR4 receptor complex coordinates with another receptor molecule that is capable of activating Gαi/o proteins remains to be determined. CXCR4 has been identified as part of an “LPS sensing apparatus” through association with TLR4 (261, 262). More recently, LPS has been shown to bind to CXCR4 and stimulate IκB phosphorylation and MAPK activation in HEK293-CXCR4 cells (268). RNA interference of CXCR4 in human endothelial cells results in a decrease in cytokine secretion by ~50% (268). This report suggests that LPS may require multiple recognition receptors, but does not explore the role of Gαi/o coupling in LPS signaling. Moreover, Gαi/o do not activate NF-κB in endothelial cells, thus the mechanism of CXCR4-mediated LPS-induced IκB activation may be independent of coupling to Gαi/o. The GPCR, protease-activated receptor 2 (PAR2), 117  has been shown to synergize with TLR4 to increase cytokine production in LPS-stimulated colonic epithelial cells (269). Furthermore, PAR2 was identified as a TLR4-interacting partner that is important for the induction of NO by LPS in macrophages (269). However, the interaction between PAR2 and TLR4 is induced by a PAR2 agonist, but not LPS, and a synergistic effect of PAR2 and TLR4 is not detected in all cell types (269). Moreover, in our hands, overexpression of PAR2 does not increase LPS-induced JNK activation in HMEC (data not shown). The sphingosine 1-phosphate (S1P) receptor family has also been implicated in cooperation with TLR4 to induce cytokine expression, namely IL-6 and IL-8, in human gingival epithelial cells (270). In addition, ERK1/2 activation was synergistically activated by both LPS and S1P (270).  Whether the S1P receptors are important in endothelial activation by LPS is unexplored. 5.2.3 Autocrine stimulation of a GPCR by an LPS-inducible product Another model for the involvement of a GPCR in LPS activation in endothelial cells, is one in which LPS stimulation of TLR4 produces a secreted factor that subsequently engages a GPCR to activate intracellular signaling. We have previously shown that pretreatment of endothelial cells with cyclohexamide does not abolish LPS-induced activation of JNK, suggesting that nascent protein synthesis is not required for activation of JNK downstream of TLR4 (115). However, this finding does not exclude the possibility that, following LPS stimulation, a cytosolic factor may be secreted into the extracellular milieu to activate a GPCR found on the plasma membrane, thereby resulting in the activation of JNK. 5.3 SASH1 AS A NOVEL TLR4 SIGNALING MOLECULE The results presented in chapter 4 outline the identification of a novel TLR4 signaling molecule, SASH1, whose function has yet to be described in the literature. The results 118  presented in this thesis show that SASH1 promotes TLR4 signaling to JNK and NF-κB by acting as a scaffolding molecule to create a molecular complex around TRAF6. 5.3.1 Scaffold molecules in innate immune signaling To date, scaffold molecules in innate immune signaling have not been extensively studied and it remains unclear how the molecular complexes are assembled downstream of TLRs. Scaffold molecules in immune signaling have been primarily characterized in the adaptive immune response downstream of receptors such as the T-cell receptor (TCR) and the B-cell receptor (BCR) (203). Caspase recruitment domain-containing protein 11 (CARD11, also called CARMA1) has been identified as a scaffold molecule important for NF-κB and JNK activation downstream of the TCR (271, 272). In immune signaling, CARD11 functions as a complex with mucosa-associated-lymphoid-tissue-lymphoma- translocation gene 1 (MALT1) and Bcl10 (273). Interestingly, a complex of IRAK1- MALT1-Bcl10-TRAF6-TAK1 has been shown to modulate LPS-induced NF-κB activation in macrophages (274) and CARD11 has been shown to play a role in LPS-induced B cell proliferation and JNK activation (275). Whether the complex of CARD11-Bcl10-MALT1 plays a role in TLR4 signaling in endothelial cells has not been established. However, our preliminary results suggest that overexpressed SASH1 does not bind to Bcl10 in HEK293T cells. As outlined in the introduction, Pellino proteins have been implicated as scaffold molecules downstream of IL-1R/TLRs (208). Yeast two-hybrid analysis identified Pellino2 as a direct interacting partner of IRAK4 (276), and endogenous IRAK-Pellino1 interaction is stimulated by LPS (277). Although Pellino1 has been implicated in IL-1-induced NF-κB signaling, it does not play a role in MAPK activation (211, 278). Pellino2 overexpression increases signaling in reporter assays for NF-κB and AP1 activation, but these findings may 119  be explained by a general role for Pellino2 in transcriptional activation (276). Furthermore, the role of Pellino2 in activation of ERK1/2 and JNK remains controversial due to the lack of reproducibility from different laboratories (276, 278). In contrast, Pellino3 mediates activation of MAPK in HEK293 cells (278, 279), but is thought to be a negative regulator of NF-κB (280), although the exact mechanism remains unknown. What is clear from these studies is that the role of the Pellino family in TLR signaling has not been well characterized and their role in endothelial cells has never been investigated. Thus, SASH1 provides the first example of a TLR scaffolding molecule in endothelial cells. 5.3.2 SASH1 in other TRAF6 signaling cascades In addition to the role of TRAF6 in TLR/IL-1R signaling pathways, TRAF6 also mediates activation of NF-κB and JNK downstream of numerous other receptors, including the TNFR superfamily (106, 281), RANK (282) and TGFβ (283), among others. Recently, the complex of TRAF6/TAK1 has been shown to mediate Smad-independent activation of JNK and p38 downstream of TGFβ (283).  Interestingly, microarray analysis of TGFβ target genes identified SASH1 as a candidate gene downregulated by TGFβ in a Smad4- independent manner (284). Whether SASH1 plays a scaffolding role downstream of other TRAF6 signaling receptors is unknown. 5.3.3 SASH1 as a negative regulator of TLR4 signaling The concentration and localization of scaffold molecules can regulate signaling cascades, allowing scaffold proteins to both amplify and attenuate signals from a receptor (205). Indeed, Pellino3 has been shown to have contrasting roles downstream of IL-1R/TLR for regulation of MAPK and NF-κB activation (278-280), but whether this is due to a spatial or temporal regulation is unknown. The discovery of novel regulators of TLR4 signaling is 120  imperative to our understanding of how the host is protected from inflammation-induced damage. Recently, β-arrestin 1 and β-arrestin 2 have been identified as negative regulators of TRAF6 ubiquitination and subsequently, downstream signaling (184). We have found that SASH1 interacts with both β-arrestin 1 and β-arrestin 2 (Dauphinee et al., unpublished), but whether this interaction plays a role in the regulation of TRAF6 ubiquitination remains unknown. Moreover, whether SASH1 regulates the K63-linked polyubiquitination of other TLR signaling molecules, such as IRAK1, IKKγ and TAK1, is also unknown. Additionally, the recent discovery of SARM as a negative regulator of TRIF-dependent signaling may also be of relevance to the work presented in this thesis (82). SASH1 has two SAM domains that can interact heterotypically with other SAM-domain containing proteins, such as the SAM domain found in SARM. Whether SARM is a binding partner of SASH1 and the biological significance of this potential interaction remains unexplored. 5.3.4 Future directions studying SASH1 in vivo The insertion of the gene-trap vector in intron 14-15 of Sash1 disrupts the SH3 domain, generating a SASH1-LacZ fusion polypeptide lacking most of the SH3 domain, the two SAM domains and the putative TRAF6 binding domain. The loss of these major functional domains suggests that the in vivo function of SASH1 will be lost in these mice. Indeed, we have not identified any viable homozygotes. Since our preliminary studies have failed to reveal any Sash1-/- mice at birth, we are also performing an in-depth genotypic analysis of various embryonic stages to determine the exact age of embryonic lethality. Once we have defined the embryonic stage at which mutant developmental processes are halted, we plan to describe any gross morphological defects and 121  examine the expression of SASH1 at, and before, this stage to aid in the characterization of the cause of embryonic lethality. Furthermore, to ensure that the phenotype of embryonic lethality is a true representation of loss of the Sash1 gene, we are creating a second gene trap mouse using ES cells with an insertion of the gene trap vector within intron 2-3 of the Sash1 locus. This insertion creates a SASH1-LacZ fusion protein that retains only the first two exons of Sash1, and results in loss of all functional domains. To study endothelial TLR signaling in the genetic absence of SASH1 in vitro, it would be useful to obtain SASH1-null endothelial cells. These cells would be a valuable tool for determining if SASH1 is absolutely critical for formation of the molecular complex around TRAF6 and subsequently downstream signaling. Since, we have been unable to decrease the level of endogenous SASH1 below 30% with lentiviral-mediated RNA interference (using three different constructs alone, or in combination with one another), we believe that formation of the complex in these cells may be facilitated by the endogenous SASH1 that is retained after knock-down. Indeed, we have seen that residual FADD, retained after RNA interference, is able to function at a level sufficient to permit signaling in knock- down cells (160). 5.4 GENERAL SUMMARY AND FUTURE PERSPECTIVES The endothelium plays a major role in the pathogenesis of sepsis at the interface between circulating blood and the surrounding tissue, in such processes as vasoregulation, vascular permeability, leukocyte recruitment and adhesion, and hemostasis (256). Therefore, insight into the molecular events that occur in endothelial cells is crucial to our understanding of the pathophysiology of sepsis in the clinical setting. 122  Endothelial cells, through the upregulation of surface adhesion molecules, such as E-selectin and VCAM-1, serve as important mediators of leukocyte recruitment following LPS challenge (285).  However, it remains unclear whether the upregulation of adhesion molecules is an indirect result of LPS activation of immune cells or a direct effect on the endothelium. Using mice that express TLR4 exclusively on the endothelium (EndotheliumTLR4), Andonegui et al. were able to investigate the importance of endothelial TLR4 signaling on adhesion molecule expression and pulmonary neutrophil accumulation (259). EndotheliumTLR4 mice are able to clear a gram-negative infection more efficiently than wild-type counterparts, which the authors explain as an increase in pulmonary neutrophil sequestration without the damaging effects caused by the excessive production of inflammatory mediators by immune cells (259). Similarly, EndotheliumTLR4 mice exhibit increased neutrophil accumulation in cerebral microvessels compared to wild-type controls (260). Recent studies have shown that mice with endothelial-specific blockade of NF-κB, caused by expression of a mutant IκBα (IκBαmt) exclusively in endothelial cells, is sufficient to inhibit expression of adhesion molecules, decrease NO production and subsequently reduce neutrophil infiltration, systemic hypotension and coagulation-events typically associated with a systemic response to LPS (223, 286). However, these mice failed to show a significant difference in the ability to clear bacteria, as compared to wild-type control mice (286). Collectively, these studies highlight the importance of endothelial- specific NF-κB signaling in the pathogenesis of septic shock. Additional studies using endothelial-specific knockout mice with targeted deletions of genes important in innate immune signaling will shed light on the importance of the 123  endothelium following LPS challenge in vivo. Indeed, our laboratory is currently making an endothelial-specific knockout of Sash1 to study the role of endothelial Sash1 in the innate immune response. Over the past few years, the focus in the field of TLRs has evolved from a simple understanding of innate immune signaling to include such concepts as the innate control of the adaptive immune response and endotoxin tolerance. Endotoxin tolerance refers to the transient insensitivity of cells toward LPS following repeated endotoxin challenge (287). First described in vivo as reduced lethality in mice pre-exposed to sublethal doses of LPS, this phenomenon has been most extensively studied in immune cells (288). However, endothelial cells also exhibit signs of endotoxin tolerance, such as decreased NF-κB activation, E-selectin expression and leukocyte adhesion (42, 289). More recently, a clinical experimental study has revealed that endothelial cell activation is reduced in vivo in patients pretreated with multiple, low doses of LPS (290). Although decreased Gαi protein expression and activity has been associated with tolerance in macrophages from LPS-tolerant rats (291, 292), the role of Gαi in endothelial tolerance has not been investigated. Certainly, identification and characterization of endothelial-specific signal regulation will contribute to an understanding of how tolerance develops in vivo. Prolonged cellular activation disturbs the normal physiological functions of the endothelium, leading to endothelial cell dysfunction and the clinical manifestations of various inflammation-associated pathologies. The pathological changes associated with endothelial dysfunction include a loss of vascular integrity and permeability functioning, thrombosis, leukocyte infiltration into the surrounding tissue and increased cytokine production. The position of the endothelium between circulating blood and the surrounding 124  tissues means that they are some of the first cells exposed to invading pathogens, and thus play a critical role in the control of inflammation during bacterial infection (17). In addition, endothelial activation/dysfunction also plays a role in the pathogenesis of various diseases, such as atherosclerosis, diabetes and arthritis. Thus, it would be of interest to investigate if SASH1 and/or Gαi play a role in endothelial activation and dysfunction as it relates to inflammation-associated disease states other than sepsis. In conclusion, this thesis has contributed to the field of innate immune signaling through the identification and characterization of novel TLR4 signaling molecules in endothelial cells. The aim is that the information presented here will further our knowledge of cell context specific signaling.              125  REFERENCES 1. Dudley, D. J. 1992. The immune system in health and disease. Baillieres Clin Obstet Gynaecol 6:393-416. 2. Medzhitov, R., and C. A. Janeway, Jr. 1998. Innate immune recognition and control of adaptive immune responses. Semin Immunol 10:351-353. 3. Pasare, C., and R. Medzhitov. 2004. Toll-like receptors: linking innate and adaptive immunity. Microbes Infect 6:1382-1387. 4. Gallo, R. L., and V. Nizet. 2008. Innate barriers against infection and associated disorders. Drug Discov Today Dis Mech 5:145-152. 5. Schmid-Wendtner, M. H., and H. C. Korting. 2006. The pH of the skin surface and its impact on the barrier function. Skin Pharmacol Physiol 19:296-302. 6. Barak, O., J. R. Treat, and W. D. James. 2005. Antimicrobial peptides: effectors of innate immunity in the skin. Adv Dermatol 21:357-374. 7. Lopez-Vidriero, M. T. 1989. Mucus as a natural barrier. Respiration 55 Suppl 1:28-32. 8. Niedergang, F., and M. N. Kweon. 2005. New trends in antigen uptake in the gut mucosa. Trends Microbiol 13:485-490. 9. Markiewski, M. M., and J. D. Lambris. 2007. The role of complement in inflammatory diseases from behind the scenes into the spotlight. Am J Pathol 171:715-727. 10. Groves, E., A. E. Dart, V. Covarelli, and E. Caron. 2008. Molecular mechanisms of phagocytic uptake in mammalian cells. Cell Mol Life Sci 65:1957-1976. 11. Heidland, A., A. Klassen, P. Rutkowski, and U. Bahner. 2006. The contribution of Rudolf Virchow to the concept of inflammation: what is still of importance? J Nephrol 19 Suppl 10:S102-109. 12. Dinarello, C. A. 2000. Proinflammatory cytokines. Chest 118:503-508. 13. Pinsky, M. R. 2004. Dysregulation of the immune response in severe sepsis. Am J Med Sci 328:220-229. 14. Mogensen, T. H. 2009. Pathogen recognition and inflammatory signaling in innate immune defenses. Clin Microbiol Rev 22:240-273, Table of Contents. 15. Castellheim, A., O. L. Brekke, T. Espevik, M. Harboe, and T. E. Mollnes. 2009. Innate immune responses to danger signals in systemic inflammatory response syndrome and sepsis. Scand J Immunol 69:479-491. 16. Wang, T. S., and J. C. Deng. 2008. Molecular and cellular aspects of sepsis-induced immunosuppression. J Mol Med 86:495-506. 17. Jaffe, E. A. 1987. Cell biology of endothelial cells. Hum Pathol 18:234-239. 18. Grandel, U., and F. Grimminger. 2003. Endothelial responses to bacterial toxins in sepsis. Crit Rev Immunol 23:267-299. 19. Volk, T., and W. J. Kox. 2000. Endothelium function in sepsis. Inflamm Res 49:185-198. 20. Cines, D. B., E. S. Pollak, C. A. Buck, J. Loscalzo, G. A. Zimmerman, R. P. McEver, J. S. Pober, T. M. Wick, B. A. Konkle, B. S. Schwartz, E. S. Barnathan, K. R. McCrae, B. A. Hug, A. M. Schmidt, and D. M. Stern. 1998. Endothelial cells in physiology and in the pathophysiology of vascular disorders. Blood 91:3527-3561. 21. Chatterjee, A., S. M. Black, and J. D. Catravas. 2008. Endothelial nitric oxide (NO) and its pathophysiologic regulation. Vascul Pharmacol 49:134-140. 22. Vo, P. A., B. Lad, J. A. Tomlinson, S. Francis, and A. Ahluwalia. 2005. autoregulatory role of endothelium-derived nitric oxide (NO) on Lipopolysaccharide-induced vascular inducible NO synthase expression and function. J Biol Chem 280:7236-7243. 23. Connelly, L., M. Madhani, and A. J. Hobbs. 2005. Resistance to endotoxic shock in endothelial nitric-oxide synthase (eNOS) knock-out mice: a pro-inflammatory role for eNOS- derived no in vivo. J Biol Chem 280:10040-10046. 126  24. Dejana, E. 1996. Endothelial adherens junctions: implications in the control of vascular permeability and angiogenesis. J Clin Invest 98:1949-1953. 25. Yuan, S. Y., M. H. Wu, E. E. Ustinova, M. Guo, J. H. Tinsley, P. De Lanerolle, and W. Xu. 2002. Myosin light chain phosphorylation in neutrophil-stimulated coronary microvascular leakage. Circ Res 90:1214-1221. 26. London, N. R., K. J. Whitehead, and D. Y. Li. 2009. Endogenous endothelial cell signaling systems maintain vascular stability. Angiogenesis 12:149-158. 27. Gong, P., D. J. Angelini, S. Yang, G. Xia, A. S. Cross, D. Mann, D. D. Bannerman, S. N. Vogel, and S. E. Goldblum. 2008. TLR4 signaling is coupled to SRC family kinase activation, tyrosine phosphorylation of zonula adherens proteins, and opening of the paracellular pathway in human lung microvascular endothelia. J Biol Chem 283:13437- 13449. 28. Bannerman, D. D., M. Sathyamoorthy, and S. E. Goldblum. 1998. Bacterial lipopolysaccharide disrupts endothelial monolayer integrity and survival signaling events through caspase cleavage of adherens junction proteins. J Biol Chem 273:35371-35380. 29. Weber, C., L. Fraemohs, and E. Dejana. 2007. The role of junctional adhesion molecules in vascular inflammation. Nat Rev Immunol 7:467-477. 30. Ostermann, G., L. Fraemohs, T. Baltus, A. Schober, M. Lietz, A. Zernecke, E. A. Liehn, and C. Weber. 2005. Involvement of JAM-A in mononuclear cell recruitment on inflamed or atherosclerotic endothelium: inhibition by soluble JAM-A. Arterioscler Thromb Vasc Biol 25:729-735. 31. Ozaki, H., K. Ishii, H. Horiuchi, H. Arai, T. Kawamoto, K. Okawa, A. Iwamatsu, and T. Kita. 1999. Cutting edge: combined treatment of TNF-alpha and IFN-gamma causes redistribution of junctional adhesion molecule in human endothelial cells. J Immunol 163:553-557. 32. Johnstone, S., B. Isakson, and D. Locke. 2009. Biological and biophysical properties of vascular connexin channels. Int Rev Cell Mol Biol 278:69-118. 33. Straub, A. C., S. R. Johnstone, K. R. Heberlein, M. J. Rizzo, A. K. Best, S. Boitano, and B. E. Isakson. 2009. Site-Specific Connexin Phosphorylation Is Associated with Reduced Heterocellular Communication between Smooth Muscle and Endothelium. J Vasc Res 47:277-286. 34. Figueroa, X. F., and B. R. Duling. 2009. Gap junctions in the control of vascular function. Antioxid Redox Signal 11:251-266. 35. Stan, R. V. 2007. Endothelial stomatal and fenestral diaphragms in normal vessels and angiogenesis. J Cell Mol Med 11:621-643. 36. Satchell, S. C., and F. Braet. 2009. Glomerular endothelial cell fenestrations: an integral component of the glomerular filtration barrier. Am J Physiol Renal Physiol 296:F947-956. 37. Levick, J. R., and L. H. Smaje. 1987. An analysis of the permeability of a fenestra. Microvasc Res 33:233-256. 38. Wisse, E., R. B. De Zanger, K. Charels, P. Van Der Smissen, and R. S. McCuskey. 1985. The liver sieve: considerations concerning the structure and function of endothelial fenestrae, the sinusoidal wall and the space of Disse. Hepatology 5:683-692. 39. Braet, F., R. De Zanger, T. Sasaoki, M. Baekeland, P. Janssens, B. Smedsrod, and E. Wisse. 1994. Assessment of a method of isolation, purification, and cultivation of rat liver sinusoidal endothelial cells. Lab Invest 70:944-952. 40. Kubes, P. 2002. The complexities of leukocyte recruitment. Semin Immunol 14:65-72. 41. Ulbrich, H., E. E. Eriksson, and L. Lindbom. 2003. Leukocyte and endothelial cell adhesion molecules as targets for therapeutic interventions in inflammatory disease. Trends Pharmacol Sci 24:640-647. 42. Lush, C. W., G. Cepinskas, and P. R. Kvietys. 2000. LPS tolerance in human endothelial cells: reduced PMN adhesion, E-selectin expression, and NF-kappaB mobilization. Am J Physiol Heart Circ Physiol 278:H853-861. 127  43. Boyle, E. M., Jr., J. C. Kovacich, T. G. Canty, Jr., E. N. Morgan, E. Chi, E. D. Verrier, and T. H. Pohlman. 1998. Inhibition of nuclear factor-kappa B nuclear localization reduces human E-selectin expression and the systemic inflammatory response. Circulation 98:II282-288. 44. Collins, T., M. A. Read, A. S. Neish, M. Z. Whitley, D. Thanos, and T. Maniatis. 1995. Transcriptional regulation of endothelial cell adhesion molecules: NF-kappa B and cytokine- inducible enhancers. Faseb J 9:899-909. 45. Coisne, C., C. Faveeuw, Y. Delplace, L. Dehouck, F. Miller, R. Cecchelli, and B. Dehouck. 2006. Differential expression of selectins by mouse brain capillary endothelial cells in vitro in response to distinct inflammatory stimuli. Neurosci Lett 392:216-220. 46. Matsukawa, A., N. W. Lukacs, C. M. Hogaboam, R. N. Knibbs, D. C. Bullard, S. L. Kunkel, and L. M. Stoolman. 2002. Mice genetically lacking endothelial selectins are resistant to the lethality in septic peritonitis. Exp Mol Pathol 72:68-76. 47. Wu, K. K., and P. Thiagarajan. 1996. Role of endothelium in thrombosis and hemostasis. Annu Rev Med 47:315-331. 48. Dahlback, B., and B. O. Villoutreix. 2005. The anticoagulant protein C pathway. FEBS Lett 579:3310-3316. 49. Kume, M., T. Hayashi, H. Yuasa, H. Tanaka, J. Nishioka, M. Ido, E. C. Gabazza, Y. Kawarada, and K. Suzuki. 2003. Bacterial lipopolysaccharide decreases thrombomodulin expression in the sinusoidal endothelial cells of rats -- a possible mechanism of intrasinusoidal microthrombus formation and liver dysfunction. J Hepatol 38:9-17. 50. Moore, K. L., S. P. Andreoli, N. L. Esmon, C. T. Esmon, and N. U. Bang. 1987. Endotoxin enhances tissue factor and suppresses thrombomodulin expression of human vascular endothelium in vitro. J Clin Invest 79:124-130. 51. Pawlinski, R., and N. Mackman. 2004. Tissue factor, coagulation proteases, and protease- activated receptors in endotoxemia and sepsis. Crit Care Med 32:S293-297. 52. Rietschel, E. T., H. Brade, O. Holst, L. Brade, S. Muller-Loennies, U. Mamat, U. Zahringer, F. Beckmann, U. Seydel, K. Brandenburg, A. J. Ulmer, T. Mattern, H. Heine, J. Schletter, H. Loppnow, U. Schonbeck, H. D. Flad, S. Hauschildt, U. F. Schade, F. Di Padova, S. Kusumoto, and R. R. Schumann. 1996. Bacterial endotoxin: Chemical constitution, biological recognition, host response, and immunological detoxification. Curr Top Microbiol Immunol 216:39-81. 53. Kawahara, K., U. Seydel, M. Matsuura, H. Danbara, E. T. Rietschel, and U. Zahringer. 1991. Chemical structure of glycosphingolipids isolated from Sphingomonas paucimobilis. FEBS Lett 292:107-110. 54. Nikaido, H., and M. Vaara. 1985. Molecular basis of bacterial outer membrane permeability. Microbiol Rev 49:1-32. 55. O'Neill, L. A. 2008. The interleukin-1 receptor/Toll-like receptor superfamily: 10 years of progress. Immunol Rev 226:10-18. 56. Yoneyama, M., and T. Fujita. 2008. Structural mechanism of RNA recognition by the RIG-I- like receptors. Immunity 29:178-181. 57. Shaw, M. H., T. Reimer, Y. G. Kim, and G. Nunez. 2008. NOD-like receptors (NLRs): bona fide intracellular microbial sensors. Curr Opin Immunol 20:377-382. 58. Lemaitre, B., E. Nicolas, L. Michaut, J. M. Reichhart, and J. A. Hoffmann. 1996. The dorsoventral regulatory gene cassette spatzle/Toll/cactus controls the potent antifungal response in Drosophila adults. Cell 86:973-983. 59. Gay, N. J., and F. J. Keith. 1991. Drosophila Toll and IL-1 receptor. Nature 351:355-356. 60. Lauw, F. N., D. R. Caffrey, and D. T. Golenbock. 2005. Of mice and man: TLR11 (finally) finds profilin. Trends Immunol 26:509-511. 61. Wu, H., H. Wang, W. Xiong, S. Chen, H. Tang, and D. Han. 2008. Expression patterns and functions of toll-like receptors in mouse sertoli cells. Endocrinology 149:4402-4412. 128  62. Barton, G. M., J. C. Kagan, and R. Medzhitov. 2006. Intracellular localization of Toll-like receptor 9 prevents recognition of self DNA but facilitates access to viral DNA. Nat Immunol 7:49-56. 63. Poltorak, A., I. Smirnova, X. He, M. Y. Liu, C. Van Huffel, O. McNally, D. Birdwell, E. Alejos, M. Silva, X. Du, P. Thompson, E. K. Chan, J. Ledesma, B. Roe, S. Clifton, S. N. Vogel, and B. Beutler. 1998. Genetic and physical mapping of the Lps locus: identification of the toll-4 receptor as a candidate gene in the critical region. Blood Cells Mol Dis 24:340-355. 64. Poltorak, A., X. He, I. Smirnova, M. Y. Liu, C. Van Huffel, X. Du, D. Birdwell, E. Alejos, M. Silva, C. Galanos, M. Freudenberg, P. Ricciardi-Castagnoli, B. Layton, and B. Beutler. 1998. Defective LPS signaling in C3H/HeJ and C57BL/10ScCr mice: mutations in Tlr4 gene. Science 282:2085-2088. 65. Schumann, R. R., S. R. Leong, G. W. Flaggs, P. W. Gray, S. D. Wright, J. C. Mathison, P. S. Tobias, and R. J. Ulevitch. 1990. Structure and function of lipopolysaccharide binding protein. Science 249:1429-1431. 66. Herzum, I., and H. Renz. 2008. Inflammatory markers in SIRS, sepsis and septic shock. Curr Med Chem 15:581-587. 67. Dunzendorfer, S., H. K. Lee, K. Soldau, and P. S. Tobias. 2004. TLR4 is the signaling but not the lipopolysaccharide uptake receptor. J Immunol 173:1166-1170. 68. da Silva Correia, J., K. Soldau, U. Christen, P. S. Tobias, and R. J. Ulevitch. 2001. Lipopolysaccharide is in close proximity to each of the proteins in its membrane receptor complex. transfer from CD14 to TLR4 and MD-2. J Biol Chem 276:21129-21135. 69. Wright, S. D. 1995. CD14 and innate recognition of bacteria. J Immunol 155:6-8. 70. Arditi, M., J. Zhou, R. Dorio, G. W. Rong, S. M. Goyert, and K. S. Kim. 1993. Endotoxin- mediated endothelial cell injury and activation: role of soluble CD14. Infect Immun 61:3149- 3156. 71. Jersmann, H. P., C. S. Hii, G. L. Hodge, and A. Ferrante. 2001. Synthesis and surface expression of CD14 by human endothelial cells. Infect Immun 69:479-485. 72. Lloyd, K. L., and P. Kubes. 2006. GPI-linked endothelial CD14 contributes to the detection of LPS. Am J Physiol Heart Circ Physiol 291:H473-481. 73. Pugin, J., C. C. Schurer-Maly, D. Leturcq, A. Moriarty, R. J. Ulevitch, and P. S. Tobias. 1993. Lipopolysaccharide activation of human endothelial and epithelial cells is mediated by lipopolysaccharide-binding protein and soluble CD14. Proc Natl Acad Sci U S A 90:2744- 2748. 74. Lloyd-Jones, K. L., M. M. Kelly, and P. Kubes. 2008. Varying importance of soluble and membrane CD14 in endothelial detection of lipopolysaccharide. J Immunol 181:1446-1453. 75. Nagai, Y., S. Akashi, M. Nagafuku, M. Ogata, Y. Iwakura, S. Akira, T. Kitamura, A. Kosugi, M. Kimoto, and K. Miyake. 2002. Essential role of MD-2 in LPS responsiveness and TLR4 distribution. Nat Immunol 3:667-672. 76. Matsushima, N., T. Tanaka, P. Enkhbayar, T. Mikami, M. Taga, K. Yamada, and Y. Kuroki. 2007. Comparative sequence analysis of leucine-rich repeats (LRRs) within vertebrate toll- like receptors. BMC Genomics 8:124. 77. Jin, M. S., and J. O. Lee. 2008. Structures of the toll-like receptor family and its ligand complexes. Immunity 29:182-191. 78. Werling, D., O. C. Jann, V. Offord, E. J. Glass, and T. J. Coffey. 2009. Variation matters: TLR structure and species-specific pathogen recognition. Trends Immunol 30:124-130. 79. Lee, H. K., S. Dunzendorfer, and P. S. Tobias. 2004. Cytoplasmic domain-mediated dimerizations of toll-like receptor 4 observed by beta-lactamase enzyme fragment complementation. J Biol Chem 279:10564-10574. 80. Nunez Miguel, R., J. Wong, J. F. Westoll, H. J. Brooks, L. A. O'Neill, N. J. Gay, C. E. Bryant, and T. P. Monie. 2007. A dimer of the Toll-like receptor 4 cytoplasmic domain 129  provides a specific scaffold for the recruitment of signalling adaptor proteins. PLoS One 2:e788. 81. O'Neill, L. A., and A. G. Bowie. 2007. The family of five: TIR-domain-containing adaptors in Toll-like receptor signalling. Nat Rev Immunol 7:353-364. 82. Carty, M., R. Goodbody, M. Schroder, J. Stack, P. N. Moynagh, and A. G. Bowie. 2006. The human adaptor SARM negatively regulates adaptor protein TRIF-dependent Toll-like receptor signaling. Nat Immunol 7:1074-1081. 83. Lord, K. A., B. Hoffman-Liebermann, and D. A. Liebermann. 1990. Complexity of the immediate early response of myeloid cells to terminal differentiation and growth arrest includes ICAM-1, Jun-B and histone variants. Oncogene 5:387-396. 84. Bonnert, T. P., K. E. Garka, P. Parnet, G. Sonoda, J. R. Testa, and J. E. Sims. 1997. The cloning and characterization of human MyD88: a member of an IL-1 receptor related family. FEBS Lett 402:81-84. 85. Wesche, H., W. J. Henzel, W. Shillinglaw, S. Li, and Z. Cao. 1997. MyD88: an adapter that recruits IRAK to the IL-1 receptor complex. Immunity 7:837-847. 86. Muzio, M., J. Ni, P. Feng, and V. M. Dixit. 1997. IRAK (Pelle) family member IRAK-2 and MyD88 as proximal mediators of IL-1 signaling. Science 278:1612-1615. 87. Kagan, J. C., and R. Medzhitov. 2006. Phosphoinositide-mediated adaptor recruitment controls Toll-like receptor signaling. Cell 125:943-955. 88. Gray, P., A. Dunne, C. Brikos, C. A. Jefferies, S. L. Doyle, and L. A. O'Neill. 2006. MyD88 adapter-like (Mal) is phosphorylated by Bruton's tyrosine kinase during TLR2 and TLR4 signal transduction. J Biol Chem 281:10489-10495. 89. Jefferies, C. A., S. Doyle, C. Brunner, A. Dunne, E. Brint, C. Wietek, E. Walch, T. Wirth, and L. A. O'Neill. 2003. Bruton's tyrosine kinase is a Toll/interleukin-1 receptor domain- binding protein that participates in nuclear factor kappaB activation by Toll-like receptor 4. J Biol Chem 278:26258-26264. 90. Kawai, T., O. Adachi, T. Ogawa, K. Takeda, and S. Akira. 1999. Unresponsiveness of MyD88-deficient mice to endotoxin. Immunity 11:115-122. 91. Yamamoto, M., S. Sato, H. Hemmi, H. Sanjo, S. Uematsu, T. Kaisho, K. Hoshino, O. Takeuchi, M. Kobayashi, T. Fujita, K. Takeda, and S. Akira. 2002. Essential role for TIRAP in activation of the signalling cascade shared by TLR2 and TLR4. Nature 420:324-329. 92. Horng, T., G. M. Barton, R. A. Flavell, and R. Medzhitov. 2002. The adaptor molecule TIRAP provides signalling specificity for Toll-like receptors. Nature 420:329-333. 93. Cao, Z., W. J. Henzel, and X. Gao. 1996. IRAK: a kinase associated with the interleukin-1 receptor. Science 271:1128-1131. 94. Ringwood, L., and L. Li. 2008. The involvement of the interleukin-1 receptor-associated kinases (IRAKs) in cellular signaling networks controlling inflammation. Cytokine 42:1-7. 95. Li, S., A. Strelow, E. J. Fontana, and H. Wesche. 2002. IRAK-4: a novel member of the IRAK family with the properties of an IRAK-kinase. Proc Natl Acad Sci U S A 99:5567- 5572. 96. Suzuki, N., S. Suzuki, G. S. Duncan, D. G. Millar, T. Wada, C. Mirtsos, H. Takada, A. Wakeham, A. Itie, S. Li, J. M. Penninger, H. Wesche, P. S. Ohashi, T. W. Mak, and W. C. Yeh. 2002. Severe impairment of interleukin-1 and Toll-like receptor signalling in mice lacking IRAK-4. Nature 416:750-756. 97. Oganesyan, G., S. K. Saha, B. Guo, J. Q. He, A. Shahangian, B. Zarnegar, A. Perry, and G. Cheng. 2006. Critical role of TRAF3 in the Toll-like receptor-dependent and -independent antiviral response. Nature 439:208-211. 98. Song, Y. J., K. Y. Jen, V. Soni, E. Kieff, and E. Cahir-McFarland. 2006. IL-1 receptor- associated kinase 1 is critical for latent membrane protein 1-induced p65/RelA serine 536 phosphorylation and NF-kappaB activation. Proc Natl Acad Sci U S A 103:2689-2694. 130  99. Keating, S. E., G. M. Maloney, E. M. Moran, and A. G. Bowie. 2007. IRAK-2 participates in multiple toll-like receptor signaling pathways to NFkappaB via activation of TRAF6 ubiquitination. J Biol Chem 282:33435-33443. 100. Kobayashi, K., L. D. Hernandez, J. E. Galan, C. A. Janeway, Jr., R. Medzhitov, and R. A. Flavell. 2002. IRAK-M is a negative regulator of Toll-like receptor signaling. Cell 110:191- 202. 101. Gan, L., and L. Li. 2006. Regulations and roles of the interleukin-1 receptor associated kinases (IRAKs) in innate and adaptive immunity. Immunol Res 35:295-302. 102. Cao, Z., J. Xiong, M. Takeuchi, T. Kurama, and D. V. Goeddel. 1996. TRAF6 is a signal transducer for interleukin-1. Nature 383:443-446. 103. Ye, H., J. R. Arron, B. Lamothe, M. Cirilli, T. Kobayashi, N. K. Shevde, D. Segal, O. K. Dzivenu, M. Vologodskaia, M. Yim, K. Du, S. Singh, J. W. Pike, B. G. Darnay, Y. Choi, and H. Wu. 2002. Distinct molecular mechanism for initiating TRAF6 signalling. Nature 418:443-447. 104. Chung, J. Y., Y. C. Park, H. Ye, and H. Wu. 2002. All TRAFs are not created equal: common and distinct molecular mechanisms of TRAF-mediated signal transduction. J Cell Sci 115:679-688. 105. Xu, L. G., L. Y. Li, and H. B. Shu. 2004. TRAF7 potentiates MEKK3-induced AP1 and CHOP activation and induces apoptosis. J Biol Chem 279:17278-17282. 106. Kobayashi, T., M. C. Walsh, and Y. Choi. 2004. The role of TRAF6 in signal transduction and the immune response. Microbes Infect 6:1333-1338. 107. Lamothe, B., A. D. Campos, W. K. Webster, A. Gopinathan, L. Hur, and B. G. Darnay. 2008. The RING domain and first zinc finger of TRAF6 coordinate signaling by interleukin-1, lipopolysaccharide, and RANKL. J Biol Chem 283:24871-24880. 108. Morita, Y., C. Kanei-Ishii, T. Nomura, and S. Ishii. 2005. TRAF7 sequesters c-Myb to the cytoplasm by stimulating its sumoylation. Mol Biol Cell 16:5433-5444. 109. Mendoza, H., D. G. Campbell, K. Burness, J. Hastie, N. Ronkina, J. H. Shim, J. S. Arthur, R. J. Davis, M. Gaestel, G. L. Johnson, S. Ghosh, and P. Cohen. 2008. Roles for TAB1 in regulating the IL-1-dependent phosphorylation of the TAB3 regulatory subunit and activity of the TAK1 complex. Biochem J 409:711-722. 110. Takaesu, G., S. Kishida, A. Hiyama, K. Yamaguchi, H. Shibuya, K. Irie, J. Ninomiya-Tsuji, and K. Matsumoto. 2000. TAB2, a novel adaptor protein, mediates activation of TAK1 MAPKKK by linking TAK1 to TRAF6 in the IL-1 signal transduction pathway. Mol Cell 5:649-658. 111. Kishida, S., H. Sanjo, S. Akira, K. Matsumoto, and J. Ninomiya-Tsuji. 2005. TAK1-binding protein 2 facilitates ubiquitination of TRAF6 and assembly of TRAF6 with IKK in the IL-1 signaling pathway. Genes Cells 10:447-454. 112. Kanayama, A., R. B. Seth, L. Sun, C. K. Ea, M. Hong, A. Shaito, Y. H. Chiu, L. Deng, and Z. J. Chen. 2004. TAB2 and TAB3 activate the NF-kappaB pathway through binding to polyubiquitin chains. Mol Cell 15:535-548. 113. Besse, A., B. Lamothe, A. D. Campos, W. K. Webster, U. Maddineni, S. C. Lin, H. Wu, and B. G. Darnay. 2007. TAK1-dependent signaling requires functional interaction with TAB2/TAB3. J Biol Chem 282:3918-3928. 114. Jiang, Z., J. Ninomiya-Tsuji, Y. Qian, K. Matsumoto, and X. Li. 2002. Interleukin-1 (IL-1) receptor-associated kinase-dependent IL-1-induced signaling complexes phosphorylate TAK1 and TAB2 at the plasma membrane and activate TAK1 in the cytosol. Mol Cell Biol 22:7158-7167. 115. Deng, L., C. Wang, E. Spencer, L. Yang, A. Braun, J. You, C. Slaughter, C. Pickart, and Z. J. Chen. 2000. Activation of the IkappaB kinase complex by TRAF6 requires a dimeric ubiquitin-conjugating enzyme complex and a unique polyubiquitin chain. Cell 103:351-361. 116. Chen, Z. J. 2005. Ubiquitin signalling in the NF-kappaB pathway. Nat Cell Biol 7:758-765. 131  117. Yamin, T. T., and D. K. Miller. 1997. The interleukin-1 receptor-associated kinase is degraded by proteasomes following its phosphorylation. J Biol Chem 272:21540-21547. 118. Wang, C., L. Deng, M. Hong, G. R. Akkaraju, J. Inoue, and Z. J. Chen. 2001. TAK1 is a ubiquitin-dependent kinase of MKK and IKK. Nature 412:346-351. 119. Scheidereit, C. 2006. IkappaB kinase complexes: gateways to NF-kappaB activation and transcription. Oncogene 25:6685-6705. 120. Chen, Z., J. Hagler, V. J. Palombella, F. Melandri, D. Scherer, D. Ballard, and T. Maniatis. 1995. Signal-induced site-specific phosphorylation targets I kappa B alpha to the ubiquitin- proteasome pathway. Genes Dev 9:1586-1597. 121. O'Neill, L. A., A. Dunne, M. Edjeback, P. Gray, C. Jefferies, and C. Wietek. 2003. Mal and MyD88: adapter proteins involved in signal transduction by Toll-like receptors. J Endotoxin Res 9:55-59. 122. Medvedev, A. E., W. Piao, J. Shoenfelt, S. H. Rhee, H. Chen, S. Basu, L. M. Wahl, M. J. Fenton, and S. N. Vogel. 2007. Role of TLR4 tyrosine phosphorylation in signal transduction and endotoxin tolerance. J Biol Chem 282:16042-16053. 123. Aki, D., R. Mashima, K. Saeki, Y. Minoda, M. Yamauchi, and A. Yoshimura. 2005. Modulation of TLR signalling by the C-terminal Src kinase (Csk) in macrophages. Genes Cells 10:357-368. 124. Anand, A. R., M. Cucchiarini, E. F. Terwilliger, and R. K. Ganju. 2008. The tyrosine kinase Pyk2 mediates lipopolysaccharide-induced IL-8 expression in human endothelial cells. J Immunol 180:5636-5644. 125. Anand, A. R., R. Bradley, and R. K. Ganju. 2009. LPS-induced MCP-1 expression in human microvascular endothelial cells is mediated by the tyrosine kinase, Pyk2 via the p38 MAPK/NF-kappaB-dependent pathway. Mol Immunol 46:962-968. 126. Aki, D., Y. Minoda, H. Yoshida, S. Watanabe, R. Yoshida, G. Takaesu, T. Chinen, T. Inaba, M. Hikida, T. Kurosaki, K. Saeki, and A. Yoshimura. 2008. Peptidoglycan and lipopolysaccharide activate PLCgamma2, leading to enhanced cytokine production in macrophages and dendritic cells. Genes Cells 13:199-208. 127. Huang, Q., J. Yang, Y. Lin, C. Walker, J. Cheng, Z. G. Liu, and B. Su. 2004. Differential regulation of interleukin 1 receptor and Toll-like receptor signaling by MEKK3. Nat Immunol 5:98-103. 128. Hull, C., G. McLean, F. Wong, P. J. Duriez, and A. Karsan. 2002. Lipopolysaccharide signals an endothelial apoptosis pathway through TNF receptor-associated factor 6-mediated activation of c-Jun NH2-terminal kinase. J Immunol 169:2611-2618. 129. Matsuzawa, A., K. Saegusa, T. Noguchi, C. Sadamitsu, H. Nishitoh, S. Nagai, S. Koyasu, K. Matsumoto, K. Takeda, and H. Ichijo. 2005. ROS-dependent activation of the TRAF6-ASK1- p38 pathway is selectively required for TLR4-mediated innate immunity. Nat Immunol 6:587-592. 130. Banerjee, A., R. Gugasyan, M. McMahon, and S. Gerondakis. 2006. Diverse Toll-like receptors utilize Tpl2 to activate extracellular signal-regulated kinase (ERK) in hemopoietic cells. Proc Natl Acad Sci U S A 103:3274-3279. 131. Loniewski, K. J., S. Patial, and N. Parameswaran. 2007. Sensitivity of TLR4- and -7-induced NF kappa B1 p105-TPL2-ERK pathway to TNF-receptor-associated-factor-6 revealed by RNAi in mouse macrophages. Mol Immunol 44:3715-3723. 132. Matsuguchi, T., A. Masuda, K. Sugimoto, Y. Nagai, and Y. Yoshikai. 2003. JNK-interacting protein 3 associates with Toll-like receptor 4 and is involved in LPS-mediated JNK activation. Embo J 22:4455-4464. 133. Cabanski, M., M. Steinmuller, L. M. Marsh, E. Surdziel, W. Seeger, and J. Lohmeyer. 2008. PKR regulates TLR2/TLR4-dependent signaling in murine alveolar macrophages. Am J Respir Cell Mol Biol 38:26-31. 132  134. Arndt, P. G., N. Suzuki, N. J. Avdi, K. C. Malcolm, and G. S. Worthen. 2004. Lipopolysaccharide-induced c-Jun NH2-terminal kinase activation in human neutrophils: role of phosphatidylinositol 3-Kinase and Syk-mediated pathways. J Biol Chem 279:10883- 10891. 135. Wymann, M. P., and L. Pirola. 1998. Structure and function of phosphoinositide 3-kinases. Biochim Biophys Acta 1436:127-150. 136. Franke, T. F., S. I. Yang, T. O. Chan, K. Datta, A. Kazlauskas, D. K. Morrison, D. R. Kaplan, and P. N. Tsichlis. 1995. The protein kinase encoded by the Akt proto-oncogene is a target of the PDGF-activated phosphatidylinositol 3-kinase. Cell 81:727-736. 137. Koyasu, S. 2003. The role of PI3K in immune cells. Nat Immunol 4:313-319. 138. Aksoy, E., W. Vanden Berghe, S. Detienne, Z. Amraoui, K. A. Fitzgerald, G. Haegeman, M. Goldman, and F. Willems. 2005. Inhibition of phosphoinositide 3-kinase enhances TRIF- dependent NF-kappa B activation and IFN-beta synthesis downstream of Toll-like receptor 3 and 4. Eur J Immunol 35:2200-2209. 139. Reddy, S. A., J. H. Huang, and W. S. Liao. 1997. Phosphatidylinositol 3-kinase in interleukin 1 signaling. Physical interaction with the interleukin 1 receptor and requirement in NFkappaB and AP-1 activation. J Biol Chem 272:29167-29173. 140. Ishii, K. J., F. Takeshita, I. Gursel, M. Gursel, J. Conover, A. Nussenzweig, and D. M. Klinman. 2002. Potential role of phosphatidylinositol 3 kinase, rather than DNA-dependent protein kinase, in CpG DNA-induced immune activation. J Exp Med 196:269-274. 141. Rhee, S. H., H. Kim, M. P. Moyer, and C. Pothoulakis. 2006. Role of MyD88 in phosphatidylinositol 3-kinase activation by flagellin/toll-like receptor 5 engagement in colonic epithelial cells. J Biol Chem 281:18560-18568. 142. Arbibe, L., J. P. Mira, N. Teusch, L. Kline, M. Guha, N. Mackman, P. J. Godowski, R. J. Ulevitch, and U. G. Knaus. 2000. Toll-like receptor 2-mediated NF-kappa B activation requires a Rac1-dependent pathway. Nat Immunol 1:533-540. 143. Fruman, D. A., S. B. Snapper, C. M. Yballe, L. Davidson, J. Y. Yu, F. W. Alt, and L. C. Cantley. 1999. Impaired B cell development and proliferation in absence of phosphoinositide 3-kinase p85alpha. Science 283:393-397. 144. Li, X., J. C. Tupper, D. D. Bannerman, R. K. Winn, C. J. Rhodes, and J. M. Harlan. 2003. Phosphoinositide 3 kinase mediates Toll-like receptor 4-induced activation of NF-kappa B in endothelial cells. Infect Immun 71:4414-4420. 145. Ojaniemi, M., V. Glumoff, K. Harju, M. Liljeroos, K. Vuori, and M. Hallman. 2003. Phosphatidylinositol 3-kinase is involved in Toll-like receptor 4-mediated cytokine expression in mouse macrophages. Eur J Immunol 33:597-605. 146. Laird, M. H., S. H. Rhee, D. J. Perkins, A. E. Medvedev, W. Piao, M. J. Fenton, and S. N. Vogel. 2009. TLR4/MyD88/PI3K interactions regulate TLR4 signaling. J Leukoc Biol 85:966-977. 147. Schabbauer, G., M. Tencati, B. Pedersen, R. Pawlinski, and N. Mackman. 2004. PI3K-Akt pathway suppresses coagulation and inflammation in endotoxemic mice. Arterioscler Thromb Vasc Biol 24:1963-1969. 148. Fukao, T., M. Tanabe, Y. Terauchi, T. Ota, S. Matsuda, T. Asano, T. Kadowaki, T. Takeuchi, and S. Koyasu. 2002. PI3K-mediated negative feedback regulation of IL-12 production in DCs. Nat Immunol 3:875-881. 149. Guha, M., and N. Mackman. 2002. The phosphatidylinositol 3-kinase-Akt pathway limits lipopolysaccharide activation of signaling pathways and expression of inflammatory mediators in human monocytic cells. J Biol Chem 277:32124-32132. 150. Luyendyk, J. P., G. A. Schabbauer, M. Tencati, T. Holscher, R. Pawlinski, and N. Mackman. 2008. Genetic analysis of the role of the PI3K-Akt pathway in lipopolysaccharide-induced cytokine and tissue factor gene expression in monocytes/macrophages. J Immunol 180:4218- 4226. 133  151. McCudden, C. R., M. D. Hains, R. J. Kimple, D. P. Siderovski, and F. S. Willard. 2005. G- protein signaling: back to the future. Cell Mol Life Sci 62:551-577. 152. Willars, G. B. 2006. Mammalian RGS proteins: multifunctional regulators of cellular signalling. Semin Cell Dev Biol 17:363-376. 153. Ma, L., and G. Pei. 2007. Beta-arrestin signaling and regulation of transcription. J Cell Sci 120:213-218. 154. Lefkowitz, R. J., and S. K. Shenoy. 2005. Transduction of receptor signals by beta-arrestins. Science 308:512-517. 155. Solomon, K. R., E. A. Kurt-Jones, R. A. Saladino, A. M. Stack, I. F. Dunn, M. Ferretti, D. Golenbock, G. R. Fleisher, and R. W. Finberg. 1998. Heterotrimeric G proteins physically associated with the lipopolysaccharide receptor CD14 modulate both in vivo and in vitro responses to lipopolysaccharide. J Clin Invest 102:2019-2027. 156. Ferlito, M., O. G. Romanenko, K. Guyton, S. Ashton, F. Squadrito, P. V. Halushka, and J. A. Cook. 2002. Implication of Galpha i proteins and Src tyrosine kinases in endotoxin-induced signal transduction events and mediator production. J Endotoxin Res 8:427-435. 157. Fan, H., O. M. Peck, G. E. Tempel, P. V. Halushka, and J. A. Cook. 2004. Toll-like receptor 4 coupled GI protein signaling pathways regulate extracellular signal-regulated kinase phosphorylation and AP-1 activation independent of NFkappaB activation. Shock 22:57-62. 158. Fan, H., B. Zingarelli, O. M. Peck, G. Teti, G. E. Tempel, P. V. Halushka, K. Spicher, G. Boulay, L. Birnbaumer, and J. A. Cook. 2005. Lipopolysaccharide- and gram-positive bacteria-induced cellular inflammatory responses: role of heterotrimeric Galpha(i) proteins. Am J Physiol Cell Physiol 289:C293-301. 159. Hoshino, K., T. Kaisho, T. Iwabe, O. Takeuchi, and S. Akira. 2002. Differential involvement of IFN-beta in Toll-like receptor-stimulated dendritic cell activation. Int Immunol 14:1225- 1231. 160. Taniguchi, T., K. Ogasawara, A. Takaoka, and N. Tanaka. 2001. IRF family of transcription factors as regulators of host defense. Annu Rev Immunol 19:623-655. 161. Oshiumi, H., M. Sasai, K. Shida, T. Fujita, M. Matsumoto, and T. Seya. 2003. TIR- containing adapter molecule (TICAM)-2, a bridging adapter recruiting to toll-like receptor 4 TICAM-1 that induces interferon-beta. J Biol Chem 278:49751-49762. 162. Rowe, D. C., A. F. McGettrick, E. Latz, B. G. Monks, N. J. Gay, M. Yamamoto, S. Akira, L. A. O'Neill, K. A. Fitzgerald, and D. T. Golenbock. 2006. The myristoylation of TRIF-related adaptor molecule is essential for Toll-like receptor 4 signal transduction. Proc Natl Acad Sci U S A 103:6299-6304. 163. Kagan, J. C., T. Su, T. Horng, A. Chow, S. Akira, and R. Medzhitov. 2008. TRAM couples endocytosis of Toll-like receptor 4 to the induction of interferon-beta. Nat Immunol 9:361- 368. 164. Harari, O. A., P. Alcaide, D. Ahl, F. W. Luscinskas, and J. K. Liao. 2006. Absence of TRAM restricts Toll-like receptor 4 signaling in vascular endothelial cells to the MyD88 pathway. Circ Res 98:1134-1140. 165. Yamamoto, M., S. Sato, H. Hemmi, K. Hoshino, T. Kaisho, H. Sanjo, O. Takeuchi, M. Sugiyama, M. Okabe, K. Takeda, and S. Akira. 2003. Role of adaptor TRIF in the MyD88- independent toll-like receptor signaling pathway. Science 301:640-643. 166. Hacker, H., V. Redecke, B. Blagoev, I. Kratchmarova, L. C. Hsu, G. G. Wang, M. P. Kamps, E. Raz, H. Wagner, G. Hacker, M. Mann, and M. Karin. 2006. Specificity in Toll-like receptor signalling through distinct effector functions of TRAF3 and TRAF6. Nature 439:204-207. 167. Fitzgerald, K. A., S. M. McWhirter, K. L. Faia, D. C. Rowe, E. Latz, D. T. Golenbock, A. J. Coyle, S. M. Liao, and T. Maniatis. 2003. IKKepsilon and TBK1 are essential components of the IRF3 signaling pathway. Nat Immunol 4:491-496. 134  168. Sasai, M., H. Oshiumi, M. Matsumoto, N. Inoue, F. Fujita, M. Nakanishi, and T. Seya. 2005. Cutting Edge: NF-kappaB-activating kinase-associated protein 1 participates in TLR3/Toll- IL-1 homology domain-containing adapter molecule-1-mediated IFN regulatory factor 3 activation. J Immunol 174:27-30. 169. Kenny, E. F., and L. A. O'Neill. 2008. Signalling adaptors used by Toll-like receptors: an update. Cytokine 43:342-349. 170. Kaiser, W. J., and M. K. Offermann. 2005. Apoptosis induced by the toll-like receptor adaptor TRIF is dependent on its receptor interacting protein homotypic interaction motif. J Immunol 174:4942-4952. 171. Chinnaiyan, A. M., K. O'Rourke, M. Tewari, and V. M. Dixit. 1995. FADD, a novel death domain-containing protein, interacts with the death domain of Fas and initiates apoptosis. Cell 81:505-512. 172. Bannerman, D. D., J. C. Tupper, J. D. Kelly, R. K. Winn, and J. M. Harlan. 2002. The Fas- associated death domain protein suppresses activation of NF-kappa B by LPS and IL-1 beta. J Clin Invest 109:419-425. 173. Zhande, R., S. M. Dauphinee, J. A. Thomas, M. Yamamoto, S. Akira, and A. Karsan. 2007. FADD negatively regulates lipopolysaccharide signaling by impairing interleukin-1 receptor- associated kinase 1-MyD88 interaction. Mol Cell Biol 27:7394-7404. 174. Opipari, A. W., Jr., H. M. Hu, R. Yabkowitz, and V. M. Dixit. 1992. The A20 zinc finger protein protects cells from tumor necrosis factor cytotoxicity. J Biol Chem 267:12424-12427. 175. Wertz, I. E., K. M. O'Rourke, H. Zhou, M. Eby, L. Aravind, S. Seshagiri, P. Wu, C. Wiesmann, R. Baker, D. L. Boone, A. Ma, E. V. Koonin, and V. M. Dixit. 2004. De- ubiquitination and ubiquitin ligase domains of A20 downregulate NF-kappaB signalling. Nature 430:694-699. 176. Evans, P. C., H. Ovaa, M. Hamon, P. J. Kilshaw, S. Hamm, S. Bauer, H. L. Ploegh, and T. S. Smith. 2004. Zinc-finger protein A20, a regulator of inflammation and cell survival, has de- ubiquitinating activity. Biochem J 378:727-734. 177. Heyninck, K., and R. Beyaert. 1999. The cytokine-inducible zinc finger protein A20 inhibits IL-1-induced NF-kappaB activation at the level of TRAF6. FEBS Lett 442:147-150. 178. Boone, D. L., E. E. Turer, E. G. Lee, R. C. Ahmad, M. T. Wheeler, C. Tsui, P. Hurley, M. Chien, S. Chai, O. Hitotsumatsu, E. McNally, C. Pickart, and A. Ma. 2004. The ubiquitin- modifying enzyme A20 is required for termination of Toll-like receptor responses. Nat Immunol 5:1052-1060. 179. O'Reilly, S. M., and P. N. Moynagh. 2003. Regulation of Toll-like receptor 4 signalling by A20 zinc finger protein. Biochem Biophys Res Commun 303:586-593. 180. Hu, X., E. Yee, J. M. Harlan, F. Wong, and A. Karsan. 1998. Lipopolysaccharide induces the antiapoptotic molecules, A1 and A20, in microvascular endothelial cells. Blood 92:2759- 2765. 181. Lee, E. G., D. L. Boone, S. Chai, S. L. Libby, M. Chien, J. P. Lodolce, and A. Ma. 2000. Failure to regulate TNF-induced NF-kappaB and cell death responses in A20-deficient mice. Science 289:2350-2354. 182. Lefkowitz, R. J., and E. J. Whalen. 2004. beta-arrestins: traffic cops of cell signaling. Curr Opin Cell Biol 16:162-168. 183. Lefkowitz, R. J., K. Rajagopal, and E. J. Whalen. 2006. New roles for beta-arrestins in cell signaling: not just for seven-transmembrane receptors. Mol Cell 24:643-652. 184. Wang, Y., Y. Tang, L. Teng, Y. Wu, X. Zhao, and G. Pei. 2006. Association of beta-arrestin and TRAF6 negatively regulates Toll-like receptor-interleukin 1 receptor signaling. Nat Immunol 7:139-147. 185. Mink, M., B. Fogelgren, K. Olszewski, P. Maroy, and K. Csiszar. 2001. A novel human gene (SARM) at chromosome 17q11 encodes a protein with a SAM motif and structural similarity 135  to Armadillo/beta-catenin that is conserved in mouse, Drosophila, and Caenorhabditis elegans. Genomics 74:234-244. 186. Belinda, L. W., W. X. Wei, B. T. Hanh, L. X. Lei, H. Bow, and D. J. Ling. 2008. SARM: a novel Toll-like receptor adaptor, is functionally conserved from arthropod to human. Mol Immunol 45:1732-1742. 187. Liberati, N. T., K. A. Fitzgerald, D. H. Kim, R. Feinbaum, D. T. Golenbock, and F. M. Ausubel. 2004. Requirement for a conserved Toll/interleukin-1 resistance domain protein in the Caenorhabditis elegans immune response. Proc Natl Acad Sci U S A 101:6593-6598. 188. Couillault, C., N. Pujol, J. Reboul, L. Sabatier, J. F. Guichou, Y. Kohara, and J. J. Ewbank. 2004. TLR-independent control of innate immunity in Caenorhabditis elegans by the TIR domain adaptor protein TIR-1, an ortholog of human SARM. Nat Immunol 5:488-494. 189. Burns, K., J. Clatworthy, L. Martin, F. Martinon, C. Plumpton, B. Maschera, A. Lewis, K. Ray, J. Tschopp, and F. Volpe. 2000. Tollip, a new component of the IL-1RI pathway, links IRAK to the IL-1 receptor. Nat Cell Biol 2:346-351. 190. Zhang, G., and S. Ghosh. 2002. Negative regulation of toll-like receptor-mediated signaling by Tollip. J Biol Chem 277:7059-7065. 191. Bulut, Y., E. Faure, L. Thomas, O. Equils, and M. Arditi. 2001. Cooperation of Toll-like receptor 2 and 6 for cellular activation by soluble tuberculosis factor and Borrelia burgdorferi outer surface protein A lipoprotein: role of Toll-interacting protein and IL-1 receptor signaling molecules in Toll-like receptor 2 signaling. J Immunol 167:987-994. 192. Didierlaurent, A., B. Brissoni, D. Velin, N. Aebi, A. Tardivel, E. Kaslin, J. C. Sirard, G. Angelov, J. Tschopp, and K. Burns. 2006. Tollip regulates proinflammatory responses to interleukin-1 and lipopolysaccharide. Mol Cell Biol 26:735-742. 193. Brint, E. K., D. Xu, H. Liu, A. Dunne, A. N. McKenzie, L. A. O'Neill, and F. Y. Liew. 2004. ST2 is an inhibitor of interleukin 1 receptor and Toll-like receptor 4 signaling and maintains endotoxin tolerance. Nat Immunol 5:373-379. 194. Liew, F. Y., H. Liu, and D. Xu. 2005. A novel negative regulator for IL-1 receptor and Toll- like receptor 4. Immunol Lett 96:27-31. 195. Qin, J., Y. Qian, J. Yao, C. Grace, and X. Li. 2005. SIGIRR inhibits interleukin-1 receptor- and toll-like receptor 4-mediated signaling through different mechanisms. J Biol Chem 280:25233-25241. 196. Kumar, S., M. D. Minnich, and P. R. Young. 1995. ST2/T1 protein functionally binds to two secreted proteins from Balb/c 3T3 and human umbilical vein endothelial cells but does not bind interleukin 1. J Biol Chem 270:27905-27913. 197. Wald, D., J. Qin, Z. Zhao, Y. Qian, M. Naramura, L. Tian, J. Towne, J. E. Sims, G. R. Stark, and X. Li. 2003. SIGIRR, a negative regulator of Toll-like receptor-interleukin 1 receptor signaling. Nat Immunol 4:920-927. 198. Janssens, S., K. Burns, E. Vercammen, J. Tschopp, and R. Beyaert. 2003. MyD88S, a splice variant of MyD88, differentially modulates NF-kappaB- and AP-1-dependent gene expression. FEBS Lett 548:103-107. 199. Mendoza-Barbera, E., M. A. Corral-Rodriguez, A. Soares-Schanoski, M. Velarde, S. Macieira, A. Messerschmidt, E. Lopez-Collazo, and P. Fuentes-Prior. 2009. Contribution of globular death domains and unstructured linkers to MyD88.IRAK-4 heterodimer formation: an explanation for the antagonistic activity of MyD88s. Biochem Biophys Res Commun 380:183-187. 200. Wesche, H., X. Gao, X. Li, C. J. Kirschning, G. R. Stark, and Z. Cao. 1999. IRAK-M is a novel member of the Pelle/interleukin-1 receptor-associated kinase (IRAK) family. J Biol Chem 274:19403-19410. 201. Lioubin, M. N., P. A. Algate, S. Tsai, K. Carlberg, A. Aebersold, and L. R. Rohrschneider. 1996. p150Ship, a signal transduction molecule with inositol polyphosphate-5-phosphatase activity. Genes Dev 10:1084-1095. 136  202. An, H., H. Xu, M. Zhang, J. Zhou, T. Feng, C. Qian, R. Qi, and X. Cao. 2005. Src homology 2 domain-containing inositol-5-phosphatase 1 (SHIP1) negatively regulates TLR4-mediated LPS response primarily through a phosphatase activity- and PI-3K-independent mechanism. Blood 105:4685-4692. 203. Shaw, A. S., and E. L. Filbert. 2009. Scaffold proteins and immune-cell signalling. Nat Rev Immunol 9:47-56. 204. Levchenko, A., J. Bruck, and P. W. Sternberg. 2000. Scaffold proteins may biphasically affect the levels of mitogen-activated protein kinase signaling and reduce its threshold properties. Proc Natl Acad Sci U S A 97:5818-5823. 205. Locasale, J. W., A. S. Shaw, and A. K. Chakraborty. 2007. Scaffold proteins confer diverse regulatory properties to protein kinase cascades. Proc Natl Acad Sci U S A 104:13307-13312. 206. Bacher, S., J. Grosshans, W. Droge, and M. L. Schmitz. 2001. The Drosophila proteins Pelle and Tube induce JNK/AP-1 activity in mammalian cells. FEBS Lett 497:153-158. 207. Lin, C. C., Y. S. Huoh, K. R. Schmitz, L. E. Jensen, and K. M. Ferguson. 2008. Pellino proteins contain a cryptic FHA domain that mediates interaction with phosphorylated IRAK1. Structure 16:1806-1816. 208. Moynagh, P. N. 2009. The Pellino family: IRAK E3 ligases with emerging roles in innate immune signalling. Trends Immunol 30:33-42. 209. Conze, D. B., C. J. Wu, J. A. Thomas, A. Landstrom, and J. D. Ashwell. 2008. Lys63-linked polyubiquitination of IRAK-1 is required for interleukin-1 receptor- and toll-like receptor- mediated NF-kappaB activation. Mol Cell Biol 28:3538-3547. 210. Windheim, M., M. Stafford, M. Peggie, and P. Cohen. 2008. Interleukin-1 (IL-1) induces the Lys63-linked polyubiquitination of IL-1 receptor-associated kinase 1 to facilitate NEMO binding and the activation of IkappaBalpha kinase. Mol Cell Biol 28:1783-1791. 211. Jiang, Z., H. J. Johnson, H. Nie, J. Qin, T. A. Bird, and X. Li. 2003. Pellino 1 is required for interleukin-1 (IL-1)-mediated signaling through its interaction with the IL-1 receptor- associated kinase 4 (IRAK4)-IRAK-tumor necrosis factor receptor-associated factor 6 (TRAF6) complex. J Biol Chem 278:10952-10956. 212. Nagase, T., K. Ishikawa, M. Suyama, R. Kikuno, N. Miyajima, A. Tanaka, H. Kotani, N. Nomura, and O. Ohara. 1998. Prediction of the coding sequences of unidentified human genes. XI. The complete sequences of 100 new cDNA clones from brain which code for large proteins in vitro. DNA Res 5:277-286. 213. Zeller, C., B. Hinzmann, S. Seitz, H. Prokoph, E. Burkhard-Goettges, J. Fischer, B. Jandrig, L. E. Schwarz, A. Rosenthal, and S. Scherneck. 2003. SASH1: a candidate tumor suppressor gene on chromosome 6q24.3 is downregulated in breast cancer. Oncogene 22:2972-2983. 214. Qiao, F., and J. U. Bowie. 2005. The many faces of SAM. Sci STKE 2005:re7. 215. Pawson, T. 1995. Protein modules and signalling networks. Nature 373:573-580. 216. Alcock, H. E., T. J. Stephenson, J. A. Royds, and D. W. Hammond. 2003. Analysis of colorectal tumor progression by microdissection and comparative genomic hybridization. Genes Chromosomes Cancer 37:369-380. 217. Rimkus, C., M. Martini, J. Friederichs, R. Rosenberg, D. Doll, J. R. Siewert, B. Holzmann, and K. P. Janssen. 2006. Prognostic significance of downregulated expression of the candidate tumour suppressor gene SASH1 in colon cancer. Br J Cancer 95:1419-1423. 218. Beer, S., T. Scheikl, B. Reis, N. Huser, K. Pfeffer, and B. Holzmann. 2005. Impaired immune responses and prolonged allograft survival in Sly1 mutant mice. Mol Cell Biol 25:9646-9660. 219. Claudio, J. O., Y. X. Zhu, S. J. Benn, A. H. Shukla, C. J. McGlade, N. Falcioni, and A. K. Stewart. 2001. HACS1 encodes a novel SH3-SAM adaptor protein differentially expressed in normal and malignant hematopoietic cells. Oncogene 20:5373-5377. 220. Uchida, T., A. Nakao, N. Nakano, A. Kuramasu, H. Saito, K. Okumura, C. Ra, and H. Ogawa. 2001. Identification of Nash1, a novel protein containing a nuclear localization 137  signal, a sterile alpha motif, and an SH3 domain preferentially expressed in mast cells. Biochem Biophys Res Commun 288:137-141. 221. Beer, S., A. B. Simins, A. Schuster, and B. Holzmann. 2001. Molecular cloning and characterization of a novel SH3 protein (SLY) preferentially expressed in lymphoid cells. Biochim Biophys Acta 1520:89-93. 222. Zhu, Y. X., S. Benn, Z. H. Li, E. Wei, E. Masih-Khan, Y. Trieu, M. Bali, C. J. McGlade, J. O. Claudio, and A. K. Stewart. 2004. The SH3-SAM adaptor HACS1 is up-regulated in B cell activation signaling cascades. J Exp Med 200:737-747. 223. Ding, J., D. Song, X. Ye, and S. F. Liu. 2009. A pivotal role of endothelial-specific NF- kappaB signaling in the pathogenesis of septic shock and septic vascular dysfunction. J Immunol 183:4031-4038. 224. Larrivee, B., and A. Karsan. 2005. Isolation and culture of primary endothelial cells. Methods Mol Biol 290:315-329. 225. Lowry, O. H., N. J. Rosebrough, A. L. Farr, and R. J. Randall. 1951. Protein measurement with the Folin phenol reagent. J Biol Chem 193:265-275. 226. Blonder, J., K. C. Chan, H. J. Issaq, and T. D. Veenstra. 2006. Identification of membrane proteins from mammalian cell/tissue using methanol-facilitated solubilization and tryptic digestion coupled with 2D-LC-MS/MS. Nat Protoc 1:2784-2790. 227. Puntervoll, P., R. Linding, C. Gemund, S. Chabanis-Davidson, M. Mattingsdal, S. Cameron, D. M. Martin, G. Ausiello, B. Brannetti, A. Costantini, F. Ferre, V. Maselli, A. Via, G. Cesareni, F. Diella, G. Superti-Furga, L. Wyrwicz, C. Ramu, C. McGuigan, R. Gudavalli, I. Letunic, P. Bork, L. Rychlewski, B. Kuster, M. Helmer-Citterich, W. N. Hunter, R. Aasland, and T. J. Gibson. 2003. ELM server: A new resource for investigating short functional sites in modular eukaryotic proteins. Nucleic Acids Res 31:3625-3630. 228. Lentschat, A., H. Karahashi, K. S. Michelsen, L. S. Thomas, W. Zhang, S. N. Vogel, and M. Arditi. 2005. Mastoparan, a G protein agonist peptide, differentially modulates TLR4- and TLR2-mediated signaling in human endothelial cells and murine macrophages. J Immunol 174:4252-4261. 229. Yibin, G., Z. Jiang, Z. Hong, L. Gengfa, W. Liangxi, W. Guo, and L. Yongling. 2005. A synthesized cationic tetradecapeptide from hornet venom kills bacteria and neutralizes lipopolysaccharide in vivo and in vitro. Biochem Pharmacol 70:209-219. 230. Kerfoot, S. M., E. M. Long, M. J. Hickey, G. Andonegui, B. M. Lapointe, R. C. Zanardo, C. Bonder, W. G. James, S. M. Robbins, and P. Kubes. 2004. TLR4 contributes to disease- inducing mechanisms resulting in central nervous system autoimmune disease. J Immunol 173:7070-7077. 231. Hawiger, J. 2001. Innate immunity and inflammation: a transcriptional paradigm. Immunol Res 23:99-109. 232. Zen, K., A. Karsan, T. Eunson, E. Yee, and J. M. Harlan. 1998. Lipopolysaccharide-induced NF-kappaB activation in human endothelial cells involves degradation of IkappaBalpha but not IkappaBbeta. Exp Cell Res 243:425-433. 233. Zhang, F. X., C. J. Kirschning, R. Mancinelli, X. P. Xu, Y. Jin, E. Faure, A. Mantovani, M. Rothe, M. Muzio, and M. Arditi. 1999. Bacterial lipopolysaccharide activates nuclear factor- kappaB through interleukin-1 signaling mediators in cultured human dermal endothelial cells and mononuclear phagocytes. J Biol Chem 274:7611-7614. 234. Wong, F., C. Hull, R. Zhande, J. Law, and A. Karsan. 2004. Lipopolysaccharide initiates a TRAF6-mediated endothelial survival signal. Blood 103:4520-4526. 235. Alexopoulou, L., A. C. Holt, R. Medzhitov, and R. A. Flavell. 2001. Recognition of double- stranded RNA and activation of NF-kappaB by Toll-like receptor 3. Nature 413:732-738. 236. Patel, T. B. 2004. Single transmembrane spanning heterotrimeric g protein-coupled receptors and their signaling cascades. Pharmacol Rev 56:371-385. 138  237. Goren, H. J., J. K. Northup, and M. D. Hollenberg. 1985. Action of insulin modulated by pertussis toxin in rat adipocytes. Can J Physiol Pharmacol 63:1017-1022. 238. Jo, H., B. Y. Cha, H. W. Davis, and J. M. McDonald. 1992. Identification, partial purification, and characterization of two guanosine triphosphate-binding proteins associated with insulin receptors. Endocrinology 131:2855-2862. 239. Okamoto, T., T. Okamoto, Y. Murayama, Y. Hayashi, E. Ogata, and I. Nishimoto. 1993. GTP-binding protein-activator sequences in the insulin receptor. FEBS Lett 334:143-148. 240. Naiki, Y., K. S. Michelsen, W. Zhang, S. Chen, T. M. Doherty, and M. Arditi. 2005. Transforming growth factor-beta differentially inhibits MyD88-dependent, but not TRAM- and TRIF-dependent, lipopolysaccharide-induced TLR4 signaling. J Biol Chem 280:5491- 5495. 241. Mikami, F., J. H. Lim, H. Ishinaga, U. H. Ha, H. Gu, T. Koga, H. Jono, H. Kai, and J. D. Li. 2006. The transforming growth factor-beta-Smad3/4 signaling pathway acts as a positive regulator for TLR2 induction by bacteria via a dual mechanism involving functional cooperation with NF-kappaB and MAPK phosphatase 1-dependent negative cross-talk with p38 MAPK. J Biol Chem 281:22397-22408. 242. Ogawa, S., J. Lozach, C. Benner, G. Pascual, R. K. Tangirala, S. Westin, A. Hoffmann, S. Subramaniam, M. David, M. G. Rosenfeld, and C. K. Glass. 2005. Molecular determinants of crosstalk between nuclear receptors and toll-like receptors. Cell 122:707-721. 243. Fan, H., G. Teti, S. Ashton, K. Guyton, G. E. Tempel, P. V. Halushka, and J. A. Cook. 2003. Involvement of G(i) proteins and Src tyrosine kinase in TNFalpha production induced by lipopolysaccharide, group B Streptococci and Staphylococcus aureus. Cytokine 22:126-133. 244. Boraschi, D., and A. Tagliabue. 2006. The interleukin-1 receptor family. Vitam Horm 74:229- 254. 245. O'Neill, L. A., T. A. Bird, A. J. Gearing, and J. Saklatvala. 1990. Interleukin-1 signal transduction. Increased GTP binding and hydrolysis in membranes of a murine thymoma line (EL4). J Biol Chem 265:3146-3152. 246. Singer, S. J., and G. L. Nicolson. 1972. The fluid mosaic model of the structure of cell membranes. Science 175:720-731. 247. Lajoie, P., J. G. Goetz, J. W. Dennis, and I. R. Nabi. 2009. Lattices, rafts, and scaffolds: domain regulation of receptor signaling at the plasma membrane. J Cell Biol 185:381-385. 248. Triantafilou, M., F. G. Gamper, P. M. Lepper, M. A. Mouratis, C. Schumann, E. Harokopakis, R. E. Schifferle, G. Hajishengallis, and K. Triantafilou. 2007. Lipopolysaccharides from atherosclerosis-associated bacteria antagonize TLR4, induce formation of TLR2/1/CD36 complexes in lipid rafts and trigger TLR2-induced inflammatory responses in human vascular endothelial cells. Cell Microbiol 9:2030-2039. 249. Triantafilou, M., K. Miyake, D. T. Golenbock, and K. Triantafilou. 2002. Mediators of innate immune recognition of bacteria concentrate in lipid rafts and facilitate lipopolysaccharide- induced cell activation. J Cell Sci 115:2603-2611. 250. Kawai, T., and S. Akira. 2007. Signaling to NF-kappaB by Toll-like receptors. Trends Mol Med 13:460-469. 251. Darnay, B. G., J. Ni, P. A. Moore, and B. B. Aggarwal. 1999. Activation of NF-kappaB by RANK requires tumor necrosis factor receptor-associated factor (TRAF) 6 and NF-kappaB- inducing kinase. Identification of a novel TRAF6 interaction motif. J Biol Chem 274:7724- 7731. 252. Lamothe, B., A. Besse, A. D. Campos, W. K. Webster, H. Wu, and B. G. Darnay. 2007. Site- specific Lys-63-linked tumor necrosis factor receptor-associated factor 6 auto-ubiquitination is a critical determinant of I kappa B kinase activation. J Biol Chem 282:4102-4112. 253. Wooff, J., L. Pastushok, M. Hanna, Y. Fu, and W. Xiao. 2004. The TRAF6 RING finger domain mediates physical interaction with Ubc13. FEBS Lett 566:229-233. 139  254. Bradley, J. R., and J. S. Pober. 2001. Tumor necrosis factor receptor-associated factors (TRAFs). Oncogene 20:6482-6491. 255. Verstak, B., K. Nagpal, S. P. Bottomley, D. T. Golenbock, P. J. Hertzog, and A. Mansell. 2009. MyD88 adapter-like (Mal)/TIRAP interaction with TRAF6 is critical for TLR2- and TLR4-mediated NF-kappaB proinflammatory responses. J Biol Chem 284:24192-24203. 256. Dauphinee, S. M., and A. Karsan. 2006. Lipopolysaccharide signaling in endothelial cells. Lab Invest 86:9-22. 257. Medzhitov, R., P. Preston-Hurlburt, and C. A. Janeway, Jr. 1997. A human homologue of the Drosophila Toll protein signals activation of adaptive immunity. Nature 388:394-397. 258. Yoneyama, M., and T. Fujita. 2007. RIG-I family RNA helicases: cytoplasmic sensor for antiviral innate immunity. Cytokine Growth Factor Rev 18:545-551. 259. Andonegui, G., H. Zhou, D. Bullard, M. M. Kelly, S. C. Mullaly, B. McDonald, E. M. Long, S. M. Robbins, and P. Kubes. 2009. Mice that exclusively express TLR4 on endothelial cells can efficiently clear a lethal systemic Gram-negative bacterial infection. J Clin Invest 119:1921-1930. 260. Zhou, H., G. Andonegui, C. H. Wong, and P. Kubes. 2009. Role of endothelial TLR4 for neutrophil recruitment into central nervous system microvessels in systemic inflammation. J Immunol 183:5244-5250. 261. Triantafilou, K., M. Triantafilou, and R. L. Dedrick. 2001. A CD14-independent LPS receptor cluster. Nat Immunol 2:338-345. 262. Triantafilou, M., and K. Triantafilou. 2002. Lipopolysaccharide recognition: CD14, TLRs and the LPS-activation cluster. Trends Immunol 23:301-304. 263. Albarran-Juarez, J., R. Gilsbach, R. P. Piekorz, K. Pexa, N. Beetz, J. Schneider, B. Nurnberg, and L. Hein. 2009. Modulation of alpha2-adrenoceptor functions by heterotrimeric Galphai protein isoforms. J Pharmacol Exp Ther 331:35-44. 264. Okamoto, T., T. Katada, Y. Murayama, M. Ui, E. Ogata, and I. Nishimoto. 1990. A simple structure encodes G protein-activating function of the IGF-II/mannose 6-phosphate receptor. Cell 62:709-717. 265. Andonegui, G., S. M. Goyert, and P. Kubes. 2002. Lipopolysaccharide-induced leukocyte- endothelial cell interactions: a role for CD14 versus toll-like receptor 4 within microvessels. J Immunol 169:2111-2119. 266. Lorenz, E., D. D. Patel, T. Hartung, and D. A. Schwartz. 2002. Toll-like receptor 4 (TLR4)- deficient murine macrophage cell line as an in vitro assay system to show TLR4-independent signaling of Bacteroides fragilis lipopolysaccharide. Infect Immun 70:4892-4896. 267. Na, H. Y., K. Mazumdar, H. J. Moon, S. Chang, and S. Y. Seong. 2009. TLR4-independent and PKR-dependent interleukin 1 receptor antagonist expression upon LPS stimulation. Cell Immunol 259:33-40. 268. Triantafilou, M., P. M. Lepper, C. D. Briault, M. A. Ahmed, J. M. Dmochowski, C. Schumann, and K. Triantafilou. 2008. Chemokine receptor 4 (CXCR4) is part of the lipopolysaccharide "sensing apparatus". Eur J Immunol 38:192-203. 269. Rallabhandi, P., Q. M. Nhu, V. Y. Toshchakov, W. Piao, A. E. Medvedev, M. D. Hollenberg, A. Fasano, and S. N. Vogel. 2008. Analysis of proteinase-activated receptor 2 and TLR4 signal transduction: a novel paradigm for receptor cooperativity. J Biol Chem 283:24314- 24325. 270. Eskan, M. A., B. G. Rose, M. R. Benakanakere, Q. Zeng, D. Fujioka, M. H. Martin, M. J. Lee, and D. F. Kinane. 2008. TLR4 and S1P receptors cooperate to enhance inflammatory cytokine production in human gingival epithelial cells. Eur J Immunol 38:1138-1147. 271. Wang, D., Y. You, S. M. Case, L. M. McAllister-Lucas, L. Wang, P. S. DiStefano, G. Nunez, J. Bertin, and X. Lin. 2002. A requirement for CARMA1 in TCR-induced NF-kappa B activation. Nat Immunol 3:830-835. 140  272. Blonska, M., B. P. Pappu, R. Matsumoto, H. Li, B. Su, D. Wang, and X. Lin. 2007. The CARMA1-Bcl10 signaling complex selectively regulates JNK2 kinase in the T cell receptor- signaling pathway. Immunity 26:55-66. 273. Lin, X., and D. Wang. 2004. The roles of CARMA1, Bcl10, and MALT1 in antigen receptor signaling. Semin Immunol 16:429-435. 274. Dong, W., Y. Liu, J. Peng, L. Chen, T. Zou, H. Xiao, Z. Liu, W. Li, Y. Bu, and Y. Qi. 2006. The IRAK-1-BCL10-MALT1-TRAF6-TAK1 cascade mediates signaling to NF-kappaB from Toll-like receptor 4. J Biol Chem 281:26029-26040. 275. Hara, H., T. Wada, C. Bakal, I. Kozieradzki, S. Suzuki, N. Suzuki, M. Nghiem, E. K. Griffiths, C. Krawczyk, B. Bauer, F. D'Acquisto, S. Ghosh, W. C. Yeh, G. Baier, R. Rottapel, and J. M. Penninger. 2003. The MAGUK family protein CARD11 is essential for lymphocyte activation. Immunity 18:763-775. 276. Strelow, A., C. Kollewe, and H. Wesche. 2003. Characterization of Pellino2, a substrate of IRAK1 and IRAK4. FEBS Lett 547:157-161. 277. Choi, K. C., Y. S. Lee, S. Lim, H. K. Choi, C. H. Lee, E. K. Lee, S. Hong, I. H. Kim, S. J. Kim, and S. H. Park. 2006. Smad6 negatively regulates interleukin 1-receptor-Toll-like receptor signaling through direct interaction with the adaptor Pellino-1. Nat Immunol 7:1057- 1065. 278. Jensen, L. E., and A. S. Whitehead. 2003. Pellino2 activates the mitogen activated protein kinase pathway. FEBS Lett 545:199-202. 279. Butler, M. P., J. A. Hanly, and P. N. Moynagh. 2005. Pellino3 is a novel upstream regulator of p38 MAPK and activates CREB in a p38-dependent manner. J Biol Chem 280:27759- 27768. 280. Xiao, H., W. Qian, K. Staschke, Y. Qian, G. Cui, L. Deng, M. Ehsani, X. Wang, Y. W. Qian, Z. J. Chen, R. Gilmour, Z. Jiang, and X. Li. 2008. Pellino 3b negatively regulates interleukin- 1-induced TAK1-dependent NF kappaB activation. J Biol Chem 283:14654-14664. 281. Hostager, B. S. 2007. Roles of TRAF6 in CD40 signaling. Immunol Res 39:105-114. 282. Darnay, B. G., A. Besse, A. T. Poblenz, B. Lamothe, and J. J. Jacoby. 2007. TRAFs in RANK signaling. Adv Exp Med Biol 597:152-159. 283. Yamashita, M., K. Fatyol, C. Jin, X. Wang, Z. Liu, and Y. E. Zhang. 2008. TRAF6 mediates Smad-independent activation of JNK and p38 by TGF-beta. Mol Cell 31:918-924. 284. Levy, L., and C. S. Hill. 2005. Smad4 dependency defines two classes of transforming growth factor {beta} (TGF-{beta}) target genes and distinguishes TGF-{beta}-induced epithelial-mesenchymal transition from its antiproliferative and migratory responses. Mol Cell Biol 25:8108-8125. 285. Kelly, M., J. M. Hwang, and P. Kubes. 2007. Modulating leukocyte recruitment in inflammation. J Allergy Clin Immunol 120:3-10. 286. Ye, X., J. Ding, X. Zhou, G. Chen, and S. F. Liu. 2008. Divergent roles of endothelial NF- kappaB in multiple organ injury and bacterial clearance in mouse models of sepsis. J Exp Med 205:1303-1315. 287. West, M. A., and W. Heagy. 2002. Endotoxin tolerance: A review. Crit Care Med 30:S64- S73. 288. Greer, G. G., and E. T. Rietschel. 1978. Inverse relationship between the susceptibility of lipopolysaccharide (lipid A)-pretreated mice to the hypothermic and lethal effect of lipopolysaccharide. Infect Immun 20:366-374. 289. Ogawa, H., P. Rafiee, J. Heidemann, P. J. Fisher, N. A. Johnson, M. F. Otterson, B. Kalyanaraman, K. A. Pritchard, Jr., and D. G. Binion. 2003. Mechanisms of endotoxin tolerance in human intestinal microvascular endothelial cells. J Immunol 170:5956-5964. 290. Draisma, A., P. Pickkers, M. P. Bouw, and J. G. van der Hoeven. 2009. Development of endotoxin tolerance in humans in vivo. Crit Care Med 37:1261-1267. 141  291. Makhlouf, M., S. H. Ashton, J. Hildebrandt, N. Mehta, T. W. Gettys, P. V. Halushka, and J. A. Cook. 1996. Alterations in macrophage G proteins are associated with endotoxin tolerance. Biochim Biophys Acta 1312:163-168. 292. Makhlouf, M., B. Zingarelli, P. V. Halushka, and J. A. Cook. 1998. Endotoxin tolerance alters macrophage membrane regulatory G proteins. Prog Clin Biol Res 397:217-226.                                              142                 APPENDIX I                                 143   APPENDIX I   The following are a list of publications that I have co-authored, which are relevant to this thesis. For each publication, I have summarized the major findings and have indicated my contribution. A copy of these publications is included at the end of this thesis.  Dauphinee SM and Karsan A. Lipopolysaccharide signaling in endothelial cells. Lab Invest. 2006 Jan;86(1):9-22. Review. PMID 16357886  • This review summarizes the molecular signaling pathways associated with lipopolysaccharide binding to its cognate receptor, TLR4, on endothelial cells. • I wrote this review  Zhande R, Dauphinee SM, Thomas JA, Yamamoto M, Akira S and Karsan A. FADD negatively regulates lipopolysaccharide signaling by impairing interleukin-1 receptor- associated kinase 1-MyD88 interaction. Mol Cell Biol. 2007 Nov;27(21):7394-404. Epub 2007 Sep 4. PMID 17785432  • This peer-reviewed article demonstrates that FADD inhibits LPS activation of endothelial cells through inhibiting interaction between key molecules in the signaling cascade. This work provides the starting point for chapter 3 of this thesis. • I was involved in generating the following data Figure 2C, 2D. Mutant FADD inhibits LPS signaling Figure 8A, 8B. FADD suppresses endothelial sprouting in vitro Lipopolysaccharide signaling in endothelial cells Shauna M Dauphinee1,2 and Aly Karsan1,2,3,4 1Department of Medical Biophysics, British Columbia Cancer Agency, Vancouver, British Columbia, Canada; 2Experimental Medicine Program, University of British Columbia, Vancouver, British Columbia, Canada; 3Department of Pathology and Laboratory Medicine, British Columbia Cancer Agency, Vancouver, British Columbia, Canada and 4Department of Pathology and Laboratory Medicine, University of British Columbia, Vancouver, British Columbia, Canada Sepsis is the systemic immune response to severe bacterial infection. The innate immune recognition of bacterial and viral products is mediated by a family of transmembrane receptors known as Toll-like receptors (TLRs). In endothelial cells, exposure to lipopolysaccharide (LPS), a major cell wall constituent of Gram- negative bacteria, results in endothelial activation through a receptor complex consisting of TLR4, CD14 and MD2. Recruitment of the adaptor protein myeloid differentiation factor (MyD88) initiates an MyD88-dependent pathway that culminates in the early activation of nuclear factor-jB (NF-jB) and the mitogen-activated protein kinases. In parallel, a MyD88-independent pathway results in a late-phase activation of NF-jB. The outcome is the production of various proinflammatory mediators and ultimately cellular injury, leading to the various vascular sequelae of sepsis. This review will focus on the signaling pathways initiated by LPS binding to the TLR4 receptor in endothelial cells and the coordinated regulation of this pathway. Laboratory Investigation (2006) 86, 9–22. doi:10.1038/labinvest.3700366; published online 7 November 2005 Keywords: lipopolysaccharide; Toll-like receptors; innate immunity; endothelial cell; inflammation Sepsis is the leading cause of mortality in critically ill patients.1 The development of sepsis occurs as a result of a systemic inflammatory response to a severe bacterial infection.2 Under normal condi- tions, a controlled cellular response to bacterial products protects the host from infection. In sepsis, hyperactivation of the immune response leads to the excessive production of various proinflammatory cytokines and cellular injury.2 In mammals, the innate immune system is the first line of host defense involved in detecting the wide variety of invading microbial pathogens.3 Receptors of the innate immune system are activated by microbial components such as lipopolysacchar- ide (LPS) (also known as endotoxin), which is a key molecule involved in the initiation of the sepsis syndrome.3 Endothelial dysfunction in sepsis The endothelium plays a major role in the patho- genesis of sepsis. Endothelial cells line the inner wall of blood vessels, lying at the interface between circulating blood and the surrounding tissue.4 Although these cells are potentially highly suscep- tible to injury given that they are the first cells exposed to invading pathogens circulating in the bloodstream, endothelial cells have a remarkable capacity to protect themselves.4 The endothelium serves a multitude of functions that help to maintain organ homeostasis, including vasoregulation, selec- tive vascular permeability and providing an anti- coagulant surface.5 However, during infection, the normal physiological functions of the endothelium are perturbed, contributing to the organ failure characteristic of sepsis.6 Although there is debate on the extent of vascular dysfunction that is due to the direct effects of LPS on the endothelium, relative to the effects that are secondary to the release of inflammatory mediators, such as tumor necrosis factor a (TNFa), interleukin-1b (IL-1b), interferons (IFNs) and others, from macrophages and immune cells,7 this review will concentrate on the direct cellular and molecular effects of LPS on the endothelium. Received 1 September 2005; accepted 2 October 2005; published online 7 November 2005 Correspondence: Dr A Karsan, MD, Department of Medical Biophysics, British Columbia Cancer Research Centre, 675 West 10th Avenue, Vancouver, British Columbia, Canada V5Z 1L3. E-mail: akarsan@bccrc.ca Funding: CIHR Grant # MOP-64409 Laboratory Investigation (2006) 86, 9–22 & 2006 USCAP, Inc All rights reserved 0023-6837/06 $30.00 www.laboratoryinvestigation.org Vasoregulation The endothelium secretes a diversity of paracrine agents that mediate vascular tone.8 Vasodilating compounds such as nitric oxide (NO) and prostacy- clin increase the diameter of the intravascular space by relaxing adjacent smooth muscle cells within the vessel wall.9 These mediators counteract vasocon- strictors such as angiotensin to maintain a balance in vascular resistance.10 Upon exposure to LPS, endothelial cells upregulate the expression of inducible NO synthase (NOS2), which leads to an increase in the production of NO.11 The use of pharmacological NOS inhibitors in patients with septic shock increases blood pressure and restores vascular resistance.12 Furthermore, NOS2-deficient mice do not experience the impaired vasoconstric- tion associated with endotoxemia.13 This excessive production of vasoactive mediators, such as NO, results in the impaired vasoconstriction and asso- ciated hypotension seen in patients with sepsis.14 Vascular Permeability Endothelial cells serve as a selective barrier for the exchange of fluid and macromolecules from the vascular compartment to the tissue.5 An increase in vascular permeability is mediated by retraction of the endothelium and phosphorylation of the light chain of nonmuscle myosin.15 This phosphorylation event induces a conformational change in the myosin light chain that promotes the interaction of actin and myosin, thereby supporting the contractile state.15 A recombinant form of LPS derived from the horseshoe crab, Limulus polyphemus, that is designed to neutralize endotoxin action, has been shown to prevent LPS-induced actin depolymeriza- tion, thereby preventing an LPS-induced increase in endothelial permeability.16 LPS also directly con- tributes to endothelial barrier dysfunction through a caspase-mediated cleavage of junctional proteins involved in regulating transport of material between the vascular space and tissue.17 Leukocyte Recruitment and Adhesion During the healing process, circulating leukocytes are recruited to the endothelium, where they adhere and traverse between the endothelial cells to enter the site of inflammation.18 The initial step in this process involves recruitment of the leukocyte to the endothelial surface through the association between selectin molecules, which are found on the surface of both the circulating leukocytes (L-selectin) and the endothelium (E- and P-selectin), and sialylated, carbohydrate moieties on the contacting cell.18 The initial selectin-mediated rolling followed by tighter adhesion to the endothelium is also mediated by the integrin family. Integrins are heterodimeric mole- cules expressed on the surface of leukocytes that facilitate a strong association with cell-adhesion molecules (CAMs) on the surface of cytokine- stimulated endothelial cells (ICAM-1, VCAM-1).19 LPS directly increases expression of E-selectin and integrin counter receptors,20 and the upregulation of this adhesion molecule expression requires the nuclear localization of nuclear factor-kappa B (NF- kB).21,22 In septic patients, elevated levels of endo- thelial selectins correlate with poor prognosis.23 Furthermore, mice deficient for endothelial selec- tins show increased survival in an animal model of sepsis.24 Hemostasis Under normal physiological conditions, the en- dothelium is maintained in an anticoagulatory state by the action of thrombin on the surface of the endothelial cell.25 Thrombin, bound to thrombomo- dulin, cleaves protein C into its active form, activated protein C, which inhibits the activated coagulation factors, Va and VIIIa.26 During sepsis, the endothelium shifts from an anticoagulant sur- face to a procoagulant surface by reduced expression of anticoagulatory molecules such as thrombo- modulin, thereby shifting the action of thrombin towards the cleavage of fibrinogen and the genera- tion of fibrin clots.27,28 Furthermore, LPS may directly induce the prothrombotic state by upregu- lating the endothelial expression of tissue factor through an NF-kB-dependent mechanism.29 These changes contribute to the disseminated intravascu- lar coagulation characteristic of sepsis. Toll-like receptor (TLR)4 signaling LPS is a key component of the cell wall of Gram- negative bacteria.30 It is composed of three structural elements: a core oligosaccharide, an O-specific chain made up of repeating sequences of polysac- charides and a lipid A component, which is responsible for the proinflammatory properties of LPS.30 The binding of LPS to the surface of endothelial cells results in endothelial activation, as demonstrated by the expression of proinflamma- tory cytokines and adhesion molecules, and, in some cases, endothelial apoptosis.31 LPS also acti- vates monocytes and macrophages to stimulate the production of proinflammatory mediators, which in turn modulate endothelial function. Collectively, this initiates a parallel cascade of events that contribute to the clinical manifestations of sepsis. The TLRs are a family of pattern recognition receptors that are classified on the basis of homology of the cytoplasmic domain with that of the inter- leukin-1 receptor (IL-1R) family, which is known as the Toll/IL-1R (TIR) domain.32 To date, there have been 12 TLRs identified in mice33 and 10 TLRs in humans.34–36 TLR4 was established as the LPS signaling receptor based on genetic evidence from LPS signaling in endothelial cells SM Dauphinee and A Karsan 10 Laboratory Investigation (2006) 86, 9–22 the LPS-insensitive mouse strain, C3H/HeJ, which has a single point mutation in the TIR domain of TLR4. In addition, the C57BL/10ScCr strain of mice has a null mutation in the TLR4 gene that also confers resistance to LPS.37 However, it should be noted that LPS from some bacterial species, such as Porphyromonas gingivalis, activate cells through TLR2.38 The first host protein involved in the recognition of LPS is LPS-binding protein (LBP).39 LBP is an acute-phase protein, the role of which is to bring LPS to the cell surface by binding to LPS and forming a ternary complex with the LPS receptor molecule, CD14.39 In endothelial cells, LBP also serves to enhance LPS uptake.40 Formation of the complex between LPS and CD14 facilitates the transfer of LPS to the LPS receptor complex composed of TLR4 and MD2.41 CD14 is found in two forms: a membrane-bound glycosylphosphatidyl- inositol (GPI)-anchored protein (mCD14) and as a soluble proteolytic fragment lacking the GPI anchor (sCD14).42 Endothelial cells lack mCD14 found on many cell types, such as macrophages and mono- cytes, and thus require sCD14 cleaved off cells bearing mCD14 and found circulating in the plas- ma.43 In addition to the production of sCD14 by proteolysis of the membrane-bound receptor, mono- cytic cell lines also secrete sCD14 molecules that never acquire a GPI anchor.44 Although it is widely accepted that endothelial cells do not express mCD14, early-passage human umbilical vein endo- thelial cells (HUVECs) have been reported to synthesize and express mCD14 at levels that are capable of supporting LPS-induced cell activation.45 However, LPS activation of endothelial cells is primarily achieved using sCD14.46 Nevertheless, in vivo expression of mCD14 on endothelial cells has not been examined. Since CD14 lacks a trans- membrane domain, it does not have the capability to initiate intracellular signaling events.42 Concerted efforts over several years have led to the discovery of the TLR4/MD2 receptor complex as the signaling entity for LPS. MD2 is a secreted glycoprotein that functions as an indispensable extracellular adaptor molecule for LPS-initiated signaling events,47 per- haps by aiding in ligand recognition.48 In addition to CD14, TLR4 and MD2, Triantifilou et al49,50 have proposed that other molecules may be involved in CD14-independent signal initiation in macrophages. Studies using affinity chromatogra- phy, and later confirmed by fluorescence resonance energy transfer (FRET), revealed that LPS associates with the heat shock proteins, Hsp70 and Hsp90, chemokine receptor 4 (CXCR4) and growth differ- entiation factor 5 (GDF5).49,50 In human coronary artery endothelial cells, TLR4 functions intracellularly,51 suggesting that LPS uptake may be necessary for optimal signal trans- duction. Similarly, TLR4 colocalizes with LPS within the Golgi of intestinal epithelial cells52 and functions within this intracellular compartment to recognize internalized LPS.53 Indeed, TLR4 is the primary receptor for transduction of the LPS signal, but it is not responsible for the internalization of LPS, pointing to the existence of another receptor for the uptake of LPS.40 LPS internalization in endothe- lial cells via scavenger receptor-dependent path- ways is suggested to be important in the clearance of LPS.54 However, overexpression of a fluorescent TLR4 in human embryonic kidney (HEK293) cells showed that TLR4 cycled between the plasma membrane and the Golgi complex, but the initiation of LPS signaling occurred at the membrane, as evidenced by an inability to disrupt signaling using brefeldin A, a pharmacological agent that disrupts the Golgi.55 Thus, further studies are required to definitively elucidate the subcellular location in which LPS activates endothelial cells. In contrast, in mononcytes that express mCD14, TLR4 is found exclusively on the cell surface localized within discrete lipid microdomains following LPS stimulation.56 The TLR4 signaling cascade initiated following LPS binding is enhanced by homodimerization of the receptor and subsequent recruitment of TIR- domain-containing adaptor molecules (TIRAP) to the cytoplasmic domain of the receptor.57,58 These adaptors include myeloid differentiation factor 88 (MyD88), MyD88 adaptor-like protein (Mal), also called TIRAP, TIR-containing adaptor inducing IFNb (TRIF), also known as TIRAP-1 (TICAM-1), and TRIF-related adaptor molecule (TRAM), also called TIRAP-2 (TICAM-2).59 Activation of TLR4 leads to stimulation of both a MyD88-dependent and a MyD88-independent path- way.60 The major players involved in eliciting the functional effects of LPS within endothelial cells are activated through the NF-kB, mitogen-activated protein kinase (MAPK) and phosphatidylinositol 3-kinase (PI3K)/Akt pathways. These pathways reg- ulate the balance between cell viability and inflam- mation. MyD88-Dependent Signaling NF-kB activation MyD88 was originally cloned as an adaptor mole- cule within the IL-1R complex and shown to possess a C-terminal TIR domain and an N-terminal death domain (DD) that recruits the serine/threonine kinase, IL-1 receptor-associated kinase (IRAK).61 During MyD88-dependent signaling, MyD88 is recruited to the TLR4 receptor through interaction with the TIR domain of TLR4.62 This complex in turn facilitates the recruitment of IRAK1 and IRAK4 via a DD present in both molecules.61,63 Upon LPS stimulation, TIRAP also associates with TLR4 via a TIR–TIR interaction, and is essential for MyD88- dependent signaling.64 This is apparent in TIRAP- deficient mice, which exhibit delayed kinetics with respect to NF-kB and MAPK activation.65 In LPS signaling in endothelial cells SM Dauphinee and A Karsan 11 Laboratory Investigation (2006) 86, 9–22 endothelial cells, TIRAP has been shown to mediate LPS-induced NF-kB activation and apoptosis.66 However, the precise role of TIRAP has not been fully elucidated. The binding of IRAK4 to the receptor complex facilitates the transphosphoryla- tion of IRAK1, inducing IRAK1 kinase activity.67 Ectopic expression of a kinase-deficient IRAK4 dramatically reduced both IRAK1 phosphorylation and downstream NF-kB activation.63 Furthermore, IRAK4 knockout mice have severely impaired TLR signaling, indicating that IRAK4 kinase activity is essential for LPS signaling.68 The autophosphoryla- tion and activation of IRAK1 results in the ability to bind TNF receptor-associated factor-6 (TRAF6).69 TRAF6 contains an N-terminal domain comprised of a RING finger and several zinc fingers, which mediate downstream signaling, and a C-terminal domain involved in self-association and hetero- logous protein interactions.70 The complex of IRAK1 and TRAF6 dissociates from the receptor to form a complex at the membrane with transforming growth factor-b (TGF- b)-activated kinase 1 (TAK1) and the adaptor molecules, TAK1-binding protein 1 and 2 (TAB1 and TAB2).71 TAB1 functions as an activator of TAK1,72 while TAB2 links TAK1 to TRAF6 and facilitates the ubiquitination of TRAF6.73,74 More recently, a third adaptor molecule has been identi- fied, TAB3, which also interacts with TAK1 and mediates activation of NF-kB.75 Although TAK1 phosphorylation occurs at the membrane and is dependent on the formation of the TRAF6–TAK1– TAB1–TAB2–TAB3 complex, it does not become active until the complex translocates to the cyto- plasm.71 Here, it forms a complex with the ubiquitin conjugating enzyme Ubc13 and the Ubc-like protein, Uev1A, to catalyze the formation of a polyubiquitin chain linked through lysine 63 (K63) of ubiquitin.76 When TAK1 is activated, IRAK1 is released from the complex and eventually degraded by the ubiquitin– proteasome system.77 TRAF6 is recycled through the process of deubiquitination.78 The formation of K63-linked polyubiquitin chains on TRAF6 results in TAK1-mediated phosphorylation of the IkB kinase (IKK) complex, composed of two catalytic subunits, IKKa and IKKb, and a regulatory subunit, IKKg, also known as NF-kB essential modulator (NEMO).79 Activation of IKK involves the TRAF- interacting protein with a forkhead-associated (FHA) domain (TIFA) protein.80 TIFA promotes the oligomerization of TRAF6 and enhances the auto- ubiquitinating activity of TRAF6, thereby facilitat- ing downstream activation of NF-kB and JNK.80 Activation of the IKKs leads to downstream phos- phorylation of members of the inhibitor of NF-kB (IkB) family, resulting in ubiquitin-directed protea- some-mediated degradation of the IkB members, thus permitting the release and nuclear transloca- tion of the transcription factor, NF-kB.81 In endothe- lial cells, degradation of the IkB protein is dependent on its tyrosine phosphorylation and is specific to the IkBa isoform, while IkBb and IkBg protein levels remain unchanged following LPS stimulation.82 As will be discussed below, several other parallel pathways have an impact on TLR4- mediated NF-kB activation, including PI3K, recep- tor-interacting protein (RIP) and the MAPKs. In non- endothelial cells, various PKC isoforms have also been implicated in NF-kB activation83,84 (Figure 1). PI3K activation PI3Ks are a family of kinases that catalyze the phosphorylation of phosphoinositides.85 The phos- phorylated lipid products, phosphatidylinositol (PtdIns) 3,4-bisphosphate and PtdIns 3,4,5-tripho- sphate, then act as second messengers to activate downstream events, including activation of Akt.86 B cells isolated from PI3K-deficient mice fail to Figure 1 MyD88-dependent signaling pathway. In endothelial cells, LPS binds to the TLR4 receptor complex consisting of soluble CD14 (sCD14) and MD2. This results in the recruitment of the adaptor molecules MyD88 and TIRAP. Then, IRAK1 and 4 are recruited to the receptor complex via interactions between the DDs of MyD88 and IRAKs. IRAK1 recruits and activates TRAF6, leading to the downstream activation of the IKK complex and the MAPKs. Activation of the IKK complex results in phosphoryla- tion and degradation of IkB, permitting the nuclear translocation of the transcription factor, NF-kB. This results in the expression of proinflammatory molecules. Concurrently, activation of the MAPKs, predominantly JNK, results in activation of the trans- cription factor AP-1, leading to a proapoptotic state. PI3K, through association with MyD88 or TRAF6, is also involved in NF-kB activation through an Akt-dependent mechanism. Mole- cules which are known to be involved in endothelial TLR4- mediated signaling are highlighted in red. LPS signaling in endothelial cells SM Dauphinee and A Karsan 12 Laboratory Investigation (2006) 86, 9–22 proliferate in response to LPS, indicating that PI3K plays a role in LPS-induced signaling events.87 Following LPS challenge, PI3K has been shown to exist in a complex with TLR4 and MyD88 in murine macrophages.88 Consistent with this, enforced expression of various dominant negative mutants has shown that PI3K activation is downstream of MyD88/IRAK/TRAF6, but not TIRAP.89,90 Further, PI3K and its downstream target kinase, Akt, appear to be an important component of LPS-induced NF-kB activity following its translocation to the nucleus.88,90 RIP is a serine/threonine kinase that plays a critical role in LPS-induced activation of NF-kB.91 RIP/ splenocytes exhibit decreased LPS-induced Akt phosphorylation, suggesting that RIP also connects TLR4 signaling to PI3K activation.91 RIP2 is a kinase related to RIP that contains a caspase recruitment domain (CARD) at its N-terminus.92 RIP2 knockout mice are resistant to the lethal effects of LPS-induced septic shock93 and RIP2 participates in LPS signaling in a kinase indepen- dent manner, likely functioning as an adaptor molecule94 (Figure 1). Mitogen-activated protein kinases In addition to the activation of NF-kB and PI3K, activation of TAK1 also leads to activation of a family of MAPKs consisting of p38, extracellular signal-regulated kinase (ERK) and c-jun NH2-termi- nal kinase (JNK)79 (Figure 1). Activation of the MAPKs by TLR4 involves MyD88 and TIRAP.62 However, both MyD88 and TIRAP knockout mice are able to activate MAPKs albeit with delayed kinetics, pointing to the existence of a MyD88/ TIRAP-independent pathway for the activation of MAPKs.65 The mechanism of TLR4-mediated activa- tion of MAPKs is not fully understood but it has been recognized that their activation occurs differ- entially during exposure to LPS. Activation of p38 is critically dependent on MEKK3.95 MEKK3 activates both NF-kB and p38 kinase, but not ERK, by binding to TRAF6 in mouse embryonic fibroblasts (MEFs).95 However, a domi- nant negative mutant of TRAF6 does not inhibit LPS activation of p38 in endothelial cells, suggesting that p38 does not lie downstream of TRAF6.96 Similarly, TRAF6 is not required for LPS-induced activation of ERK in endothelial cells.90 Nonetheless, in immune cells, the activation of p38 involves TRAF6 and apoptosis signal-regulating kinase 1 (ASK1).97 LPS- induced intracellular reactive oxygen species (ROS) mediate the formation of a complex between TRAF6 and ASK1 to facilitate activation of p38 but not JNK.97 Furthermore, ASK1 is required for LPS- induced cytokine production and ASK1-null mice are resistant to endotoxic shock.97 The activation of ERK in response to LPS has not been well characterized. However, the MAP kinase kinase, Tpl2 (also known as Cot) has been impli- cated in LPS-induced ERK activation in macro- phages.98 Tpl2 has been shown to coordinate with the Ras pathway to induce activation of ERK in COS- 1 monkey kidney cells99 and the Ras pathway has been implicated in LPS-induced ERK activation.100 In endothelial cells, JNK activation has been shown to require TRAF6 activation.96 In macro- phages, however, activation of JNK is dependent on interaction of the scaffolding protein JNK-interact- ing protein 3 (JIP3) with the cytoplasmic domain of TLR4.101 Inhibition of JIP3 function, either by expression of a dominant negative form of the protein or using RNA interference, inhibits JNK activation in macrophages.101 JIP3 is physically associated with MEKK1 in RAW264.7 cells.101 In neutrophils, LPS-induced JNK activation involves the participation of the protein tyrosine kinase, Syk, and PI3K.102 Taken together, these findings suggest that the receptor complex may include many more proteins than that shown in Figures 1 and 2. Functional effects of MyD88/TIRAP-dependent signaling LPS activates both NF-kB and JNK through TRAF6, leading to proinflammatory and apoptotic signaling pathways, respectively.96 TRAF6 is also the bifurca- tion point, downstream of which both death and survival signals are induced.90 We have shown that LPS induces a proapoptotic signal through TRAF6- mediated activation of JNK, which in turn leads to caspase activation following mitochondrial depolari- zation.96 Additionally, PI3K-dependent survival signals are mediated downstream of TRAF6 in endothelial cells.90 The TRAF6-interacting protein, RIP, is essential for cell survival following LPS stimulation in splenocytes and provides a link between TLR4 and the PI3K-Akt survival pathway.91 LPS has also been shown to directly stimulate endothelial sprouting in vitro and angiogenesis in vivo.103 The initiation of angiogenesis mediated by LPS is dependent on TRAF6 and the downstream effector molecules, NF-kB and JNK.103 The balance between endothelial survival and death pathways is crucial in the pathogenesis of sepsis as endothelial cell apoptosis could potentially lead to severe vascular collapse, as demonstrated by the dissemi- nated endothelial apoptosis seen in C57BL/6 mice challenged with LPS104 (Figure 2). MyD88-Independent Signaling The MyD88-independent signaling pathway was identified in studies using MyD88 deficient mice.105 Although MyD88/ mice were resistant to LPS- induced death, delayed activation of both NF-kB and MAPK pathways was observed.105 Furthermore, MyD88-deficient cells failed to release proinflam- matory cytokines in response to LPS suggesting direct activation of NF-kB and the MAPKs by LPS.106 Collectively, these results suggested the existence of a MyD88-independent pathway downstream of LPS signaling in endothelial cells SM Dauphinee and A Karsan 13 Laboratory Investigation (2006) 86, 9–22 ligand engagement of the TLR4 receptor. The MyD88-dependent pathway is largely responsible for controlling the expression of inflammatory cytokines, such as TNFa, IL-6 and IL-12, through activation of NF-kB, whereas the MyD88-indepen- dent pathway induces expression of IFN-inducible genes, such as IP10 and glucocorticoid-attenuated response gene 16 (GARG16), through activation of the transcription factor, IFN regulatory factor 3 (IRF3).107 MyD88-independent signaling begins with recruitment of the adaptor molecule, TRAM, to the cytoplasmic domain of TLR4.108 TRAM is specific to the TLR4 signaling pathway and was cloned as a TLR4-binding protein in yeast that facilitates the binding of TRIF to the receptor complex by forming heterodimers with TRIF.108,109 TRIF has been shown to play an essential role in IRF3 activation and production of IFNb.110 The recruitment of these adaptor molecules to the receptor complex results in binding of TRAF6 and RIP1 to TRIF, leading to the late-phase activation of NF-kB, which contributes to the induction of IFNb.111 In a parallel pathway, TRAF family member-associated NF-kB activator (TANK) binding kinase 1 (TBK1) and IKKe (also known as IKKi) interact with TRIF and mediate activation of IRF3, leading to the induction of IFNb.112 IRF3 is a member of a family of transcrip- tion factors that are involved in the induction of type I IFNs that translocate to the nucleus upon phosphorylation and dimerization.113 Once inside the nucleus, IRF3 controls IFNb transcription by recruiting the coactivators, p300 and CBP.113 NF-kB- activating kinase associating protein (NAP1) also interacts with TRIF and TBK1 to stimulate IFNb production through activation of IRF3.114 Although MyD88-independent pathways have not been di- rectly studied in the endothelium, recent studies have demonstrated LPS-dependent induction of MyD88-independent genes (eg, IFNb, IP10, IL-6 and iNOS) in endothelial cells115 (Figure 3). Negative regulation of TLR4 signaling Modulation of signaling cascades involved in immune regulation is imperative for the prevention Figure 2 LPS-induced signaling events in endothelial cells. Stimulation of endothelial cells with LPS results in the activation of TRAF6, leading to activation of several downstream signaling pathways, including activation of JNK and NF-kB, leading to the production of proinflammatory signaling molecules. Activation of TRAF6 also results in endothelial sprouting through JNK and NF- kB-dependent signaling pathways. Furthermore, activation of TRAF6 also results in the activation of PI3K and Akt, which results in stimulation of inflammation, but inhibition of apopto- sis. The activation of ERK in endothelial cells is TRAF6- independent. Figure 3 MyD88-independent signaling pathway. The late-phase activation of NF-kB occurs through the MyD88-independent pathway involving the adaptor molecules, TRIF and TRAM. TRIF associates with TRAF6 and RIP1, as well as NAP1 and TBK1, to lead to the activation of the transcription factors, NF-kB and IRF3, respectively. MyD88-independent signaling has not been exten- sively studied in endothelial cells. LPS signaling in endothelial cells SM Dauphinee and A Karsan 14 Laboratory Investigation (2006) 86, 9–22 of an aberrant inflammatory response. Several molecules have been identified that regulate the TLR4 signaling pathway (Figure 4). Downstream of Tyrosine Kinases (Dok) Dok proteins are a family of adaptor proteins, consisting of Dok-1–5.116–119 Dok1 and Dok2 have been reported to be involved in the negative regulation of LPS signaling.120 Mice that are defi- cient in Dok1 and Dok2 show an increase in TNFa and NO production in peritoneal and bone-marrow derived macrophages, as compared to wild-type cells.120 Furthermore, these mice exhibit an in- creased sensitivity to LPS, as demonstrated by an increase in TNFa production and increased lethality compared to wild-type mice.120 Overexpression of Dok1 or Dok2 inhibits LPS-induced ERK activa- tion.120 Dok4 is highly expressed in endothelial cells121 and has recently been implicated as a positive regulator of TNFa-mediated NF-kB activa- tion in endothelial cells.122 However, the Dok family has not been studied in the LPS response of endothelial cells. Phosphatidylinositol 3-Kinase PI3K plays a dual role in LPS signaling events as it is both positively and negatively involved in TLR signal transduction. PI3K has been shown to negatively regulate the production of inflammatory mediators important in the pathogenesis of sepsis, such as proinflammatory cytokines and NO.123,124 In endothelial cells and monocytes, inhibition of the PI3K–Akt pathway has been shown to enhance LPS- induced cytokine production, suggesting a negative regulatory role for PI3K.125 Activation of PI3K also reduces MyD88-dependent production of the pro- inflammatory molecules IL-12 in dendritic cells and TNFa in monocytes.126,127 In C6 glial cells stimulated with LPS, inhibition of PI3K results in the induction of NOS.123 Moreover, the enforced expression of a constitutively active form of PI3K has been shown to inhibit the expression of NOS2 in human astro- cytes.128 Collectively, these results demonstrate that PI3K is a negative regulator of NO production in several cell types. Conversely, PI3K has been shown to positively regulate MyD88-dependent signaling through the TLR2 receptor, although whether this is true in endothelial cells has not been studied. Inhibition of PI3K leads to a reduction in the activation of NF-kB and the MAPKs, p38 and p44/42.129 PI3K also plays a role in MyD88-independent signaling. In the TRIF- dependent signaling cascade, PI3K physically asso- ciates with TRIF in response to LPS and inhibits TRIF-dependent NF-kB transcriptional activity, but has no effect on IRF3 activation in dendritic cells.130 A20 A20 is a zinc-finger protein that was initially identified as an antiapoptotic protein, but has subsequently been shown to have both ubiquitinase and deubiquitinase activities.131–133 In endothelial cells, expression of A20 is induced by LPS in a sCD14- and NF-kB-dependent manner.134 Mice that are deficient for the expression of A20 exhibit increased responsiveness to LPS and develop severe inflammation.135 Expression of A20 inhibits TLR4- mediated NF-kB activation by regulating MEKK-1 kinase activity.136 The regulatory action imparted by A20 is dependent on its ability to act as a TRAF6 deubiquitinase, thereby reversing TRAF6 Figure 4 Modulation of TLR4-mediated signaling pathway. TLR4 signaling is regulated by several molecules. In endothelial cells, A20 functions as a deubiquitinase enzyme to inhibit activation of TRAF6 upstream of NF-kB. A1 also inhibits NF-kB activa- tion through an unknown mechanism. Both A1 and A20 are upregulated by LPS through NF-kB, suggesting a negative feed- back loop to reduce the signaling by LPS. SHIP inhibits PI3K activity. PI3K itself plays a dual role in regulating LPS-induced signaling events, acting as both a positive and a negative regulator. FADD attenuates the activation of NF-kB, perhaps by sequestering MyD88 or IRAK to prevent downstream signaling. ST2, an orphan receptor expressed in endothelial cells, sequesters the adaptor molecules MyD88 and TIRAP, preventing down- stream signaling. In nonendothelial cells, a splice variant of MyD88, MyD88s, inhibits the association of MyD88 and IRAK. IRAKM prevents the association of IRAK and TRAF6. The orphan receptor, SIGIRR, negatively regulates TLR signaling, possibly by attenuating the recruitment of downstream signaling molecules. Dok1 and Dok2 inhibit LPS-induced ERK activation. Molecules that are involved in endothelial TLR4-mediated signaling, or implicated in the regulation of this pathway, are highlighted in red. LPS signaling in endothelial cells SM Dauphinee and A Karsan 15 Laboratory Investigation (2006) 86, 9–22 activation.137 A20 also acts as a negative regulator of TNFa-induced NF-kB activation through ubiquitina- tion of RIP, targeting RIP for degradation.132 Whether A20 also regulates LPS signaling by ubiquitinating RIP remains to be shown. The upregulation of A20 by LPS suggests that A20 functions as part of a negative feedback mechanism to regulate the innate immune response to LPS. A1 A1 is a homolog of the antiapoptotic molecule, Bcl- 2, that also functions as a cytoprotective protein in response to TNF stimulation.134,138 Similar to A20, LPS upregulates mRNA expression of A1 in endothelial cells through an NF-kB-dependent mechanism,134 and protects cells from LPS-induced apoptosis. In addition to the antiapoptotic effects of A1, it has also been shown that the Bcl homology 4 (BH4) domain of A1 inhibits endothelial activation by inhibiting NF-kB activity, and thus one of the major proinflammatory pathways activated by LPS.139,140 Thus, the NF-kB-dependent upregulation of A1 by LPS acts as a negative regulator in a feedback loop to dampen the LPS signal. Fas-Associated Death Domain (FADD) FADD is a proapoptotic adaptor molecule that couples the cytoplasmic domain of death receptor molecules to effector caspases.141 Prior to identifica- tion of mammalian TLRs, we had shown that LPS regulates apoptosis through a FADD-dependent pathway that was independent of death receptor signaling.142 More recently, FADD has been shown to play an inhibitory role in LPS-induced NF-kB- dependent gene expression. FADD/ MEFs stimu- lated with LPS demonstrate increased IkB-b degra- dation compared to FADDþ /þ cells, indicating that the role of FADD is upstream of IkB degradation.143 Although a definitive role for FADD in LPS signal- ing has been established, the mechanism through which FADD exerts its effects remains unclear. MyD88 is a mediator of TLR2-induced apoptosis through a FADD-dependent pathway through the binding of FADD to the DD of MyD88 and sub- sequent recruitment of caspase 8.144 As such, it has been suggested that, during LPS signaling, FADD may bind to the DD of MyD88 and thereby prevent the obligatory interaction between MyD88 and IRAK.143 Similarly, it is also possible that FADD binds to IRAK to sequester it in the cytosol, thus preventing the interaction with MyD88 and subse- quently abrogating downstream signaling events. Toll-Interacting Protein (Tollip) Tollip was identified as a protein partner of the IL- 1R complex following IL-1 stimulation. In unstimu- lated cells, Tollip forms a complex with IRAK and is recruited to the receptor following stimulation.145 Tollip has also been shown to associate with TLR4 and is phosphorylated by IRAK following stimula- tion with LPS.146 Tollip plays a negative role in LPS signaling in macrophages by suppressing the acti- vity of IRAK before receptor activation.146 Further- more, overexpression of Tollip results in the inhibi- tion of NF-kB activity downstream of IRAK.145 Tollip is expressed in human dermal endothelial cells and ectopic expression of Tollip in these cells inhibits LPS-induced NF-kB activation.147 Src Homology 2 Domain-Containing Inositol-5-Phosphate (SHIP) SHIP is a cellular phosphatase that catalyzes the removal of a phosphate group from PtdIns 3,4,5- triphosphate, a product of PI3K activity.148 Upon initial exposure to LPS, intracellular levels of SHIP are increased, which promotes endotoxin toler- ance.149 SHIP1 negatively regulates LPS-induced activation of Akt downstream of PI3K. In addition, SHIP1 also inhibits LPS-induced MAPK activation and IkB-a degradation, although this is independent of PI3K involvement. However, this function is thought to occur by SHIP inhibiting formation of a complex between TLR4 and MyD88.150 SHIP1 is expressed in endothelial cells,151 and thus may play a role in the regulation of LPS signaling in these cells. ST2 ST2 (also known as T1) is a TIR-domain-containing orphan receptor expressed on TH2 cells and mast cells.152,153 Mice that are deficient for ST2 exhibit an increase in LPS-induced cytokine production.154 Although ST2 contains a TIR domain and is able to activate JNK, ERK and p38, it is unable to activate NF-kB.154 Forced overexpression of ST2 in HEK293 cells results in a decrease in LPS-induced NF-kB activation.155 The inhibitory effect of ST2 is the result of the interaction between its TIR domain and the TIR domains of MyD88 and TIRAP, thus sequestering these critical adaptors during LPS signaling.154 A soluble ST2 receptor was immuno- precipitated from endothelial cells, indicating that ST2 is expressed in these cells.156 The presence of ST2 in endothelial cells suggests that this molecule may also be involved in the regulation of TLR4- mediated signaling in these cells. Additional Inhibitors of TLR4 Signaling The suppressors of cytokine signaling are a family of intracellular proteins that regulate cytokine signal- ing in a classical feedback mechanism.157 LPS- induced upregulation of the SOCS1 protein has LPS signaling in endothelial cells SM Dauphinee and A Karsan 16 Laboratory Investigation (2006) 86, 9–22 been observed in a macrophage cell line and SOCS1/ mice are hyperresponsive to LPS, show- ing increased serum TNFa levels and increased lethality.158 Furthermore, ectopic expression of SOCS1 in a macrophage cell line inhibits LPS- induced NF-kB activation.158,159 SOCS1 is expressed in endothelial cells,160 but its involvement in regulating LPS signaling in these cells has not been examined. RP105 is a TLR homolog that lacks an intracellular TIR domain.161 Expression of RP105 is induced in the presence of the MD2 homolog, MD1.162 The complex of RP105 and MD1 interacts with TLR4/ MD2, impairing its ability to bind LPS.161 Moreover, dendritic cells derived from RP105-deficient mice produce higher levels of proinflammatory cytokines following LPS stimulation, compared to wild-type controls.161 Enforced expression of TLR4, MD1 and MD2 in HEK293 cells showed that RP105-mediated inhibition of IL-8 production was the result of inhibition of NF-kB activity.161 RP105 expression in endothelial cells is low, suggesting that RP105 may play only a minor role in TLR4-mediated signaling in the endothelium.163 The chemokine receptor CXCR4 plays a putative role in LPS signaling as part of a proposed LPS activation cluster.49 Inhibition of CXCR4 in HEK293 cells expressing TLR4, MD2 and CD14, using anti- CXCR4 antibodies, resulted in an increase in NF-kB luciferase activity in response to LPS, suggesting that CXCR4 inhibits signaling through TLR4.164 CXCR4 is expressed in endothelial cells165 and thus may be involved in regulating endothelial responses to LPS. Single immunoglobulin IL-1 receptor-related molecule (SIGIRR) (also known as TIR8) is another member of the TIR-domain-containing receptor family that is unable to induce inflammatory responses. SIGIRR is thought to play a role in regulation of inflammatory events as its expression is downregulated in tissues of mice injected with LPS.166 Furthermore, the role of SIGIRR in LPS signaling is thought to be negative, since it interacts with both TLR4 and TRAF6.166 SIGIRR has recently been shown to interact with TLR4 through its TIR domain to attenuate recruitment of downstream signaling molecules.167 Although SIGIRR functions as a modulator of TLR signaling, expression of SIGIRR is relatively low in endothelial compared with epithelial cell lines, suggesting that it may play only a minimal role in signaling in the endothe- lium.166 Using a yeast two-hybrid approach, TRIAD3 was identified as a RING-finger protein that interacts with the cytoplasmic tail of several TLRs, including TLR4.168 The most abundant isoform identified in a range of cell lines is TRIAD3A, which contains structural features reminiscent of an E3 ubiquitin- protein ligase and promotes degradation of the receptor by targeting the receptor for ubiquitination and proteolytic degradation.168 Furthermore, deple- tion of intracellular TRIAD3A results in an increase in TLR4-stimulated NF-kB luciferase activity.168 TRIAD3A expression has not been examined in endothelial cells. MyD88s is a splice variant of MyD88 that lacks the intermediate domain required for recruitment of IRAK4.169 MyD88s inhibits TLR4 signaling by pre- venting the recruitment of IRAK4 to MyD88 in the receptor complex, thus abolishing phosphorylation and activation of IRAK1.67 As a result, MyD88s functions as a dominant-negative inhibitor and is unable to activate NF-kB. However, activation of JNK is unaffected by MyD88s expression, suggesting that pathways leading to NF-kB and MAPK activa- tion may diverge at the level of MyD88.170 MyD88s expression has not been determined in endothelial cells. IRAK-M is a functional member of the IRAK/Pelle family that is expressed exclusively in monocytic cell lines. IRAK-M can interact with both MyD88 and TRAF6 and possesses intrinsic kinase activity, although it is incapable of autophosphorylation.171 IRAK-M plays a negative role in TLR4 signaling by preventing the dissociation of IRAK4 and MyD88 and thus preventing the formation of the IRAK– TRAF6 complex. Furthermore, IRAK-M knockout mice show an increased responsiveness to bacterial LPS.172 Conclusion The mechanism of LPS-induced signaling events mediated by TLR4 has been extensively studied over recent years. Much work has been carried out to elucidate the role of specific molecules comprising the signaling pathways and to identify the negative regulators of the LPS signaling cascades such that therapeutic approaches may be derived from these discoveries. Novel players involved in TLR4 signal transduction, such as Nod1 and Nod2,173 as well as the demonstration of crosstalk interactions with other signaling pathways, such as those seen between TGFb174 and TLR4 signaling cascades, will provide further insight into the molecular mecha- nisms of innate immunity. Although research in the field of TLR signaling has predominantly focused on the pathways in- itiated in macrophages and monocytes, it is evident that the endothelium plays a critical role during sepsis. A deeper understanding of the regulatory elements of TLR4-mediated survival and death pathways in endothelial cells may facilitate the development of novel therapies to avoid the patho- logical events associated with endothelial injury during sepsis. References 1 Angus DC, Linde-Zwirble WT, Lidicker J, et al. Epidemiology of severe sepsis in the United States: LPS signaling in endothelial cells SM Dauphinee and A Karsan 17 Laboratory Investigation (2006) 86, 9–22 analysis of incidence, outcome, and associated costs of care. Crit Care Med 2001;29:1303–1310. 2 Pinsky MR. Dysregulation of the immune response in severe sepsis. Am J Med Sci 2004;328:220–229. 3 Medzhitov R. Toll-like receptors and innate immu- nity. Nat Rev Immunol 2001;1:135–145. 4 Jaffe EA. Cell biology of endothelial cells. Hum Pathol 1987;18:234–239. 5 Grandel U, Grimminger F. Endothelial responses to bacterial toxins in sepsis. Crit Rev Immunol 2003; 23:267–299. 6 Volk T, Kox WJ. Endothelium function in sepsis. Inflamm Res 2000;49:185–198. 7 Salgado A, Boveda JL, Monasterio J, et al. Inflamma- tory mediators and their influence on haemostasis. Haemostasis 1994;24:132–138. 8 Sumpio BE, Riley JT, Dardik A. Cells in focus: endothelial cell. Int J Biochem Cell Biol 2002;34: 1508–1512. 9 Rongen GA, Smits P, Thien T. Endothelium and the regulation of vascular tone with emphasis on the role of nitric oxide. Physiology, pathophysiology and clinical implications. Neth J Med 1994;44:26–35. 10 Ruschitzka F, Corti R, Noll G, et al. A rationale for treatment of endothelial dysfunction in hypertension. J Hypertens Suppl 1999;17:S25–S35. 11 Morikawa A, Koide N, Kato Y, et al. Augmentation of nitric oxide production by gamma interferon in a mouse vascular endothelial cell line and its modula- tion by tumor necrosis factor alpha and lipopolysac- charide. Infect Immun 2000;68:6209–6214. 12 Petros A, Bennett D, Vallance P. Effect of nitric oxide synthase inhibitors on hypotension in patients with septic shock. Lancet 1991;338:1557–1558. 13 Spohr F, Cornelissen AJ, Busch C, et al. Role of endogenous nitric oxide in endotoxin-induced alteration of hypoxic pulmonary vasoconstriction in mice. Am J Physiol Heart Circ Physiol 2005;289: H823–H831. 14 Thiemermann C. Nitric oxide and septic shock. Gen Pharmacol 1997;29:159–166. 15 Yuan SY. Protein kinase signaling in the modulation of microvascular permeability. Vascul Pharmacol 2002;39:213–223. 16 Bannerman DD, Fitzpatrick MJ, Anderson DY, et al. Endotoxin-neutralizing protein protects against en- dotoxin-induced endothelial barrier dysfunction. In- fect Immun 1998;66:1400–1407. 17 Bannerman DD, Sathyamoorthy M, Goldblum SE. Bacterial lipopolysaccharide disrupts endothelial monolayer integrity and survival signaling events through caspase cleavage of adherens junction pro- teins. J Biol Chem 1998;273:35371–35380. 18 Kubes P. The complexities of leukocyte recruitment. Semin Immunol 2002;14:65–72. 19 Ulbrich H, Eriksson EE, Lindbom L. Leukocyte and endothelial cell adhesion molecules as targets for therapeutic interventions in inflammatory disease. Trends Pharmacol Sci 2003;24:640–647. 20 Lush CW, Cepinskas G, Kvietys PR. LPS tolerance in human endothelial cells: reduced PMN adhesion, E-selectin expression, and NF-kappaB mobilization. Am J Physiol Heart Circ Physiol 2000;278:H853– H861. 21 Boyle Jr EM, Kovacich JC, Canty Jr TG, et al. Inhibition of nuclear factor-kappa B nuclear localiza- tion reduces human E-selectin expression and the systemic inflammatory response. Circulation 1998; 98:II282–II288. 22 Collins T, Read MA, Neish AS, et al. Transcriptional regulation of endothelial cell adhesion molecules: NF-kappa B and cytokine-inducible enhancers. FAS- EB J 1995;9:899–909. 23 Cummings CJ, Sessler CN, Beall LD, et al. Soluble E- selectin levels in sepsis and critical illness. Correla- tion with infection and hemodynamic dysfunction. Am J Respir Crit Care Med 1997;156:431–437. 24 Matsukawa A, Lukacs NW, Hogaboam CM, et al. Mice genetically lacking endothelial selectins are resistant to the lethality in septic peritonitis. Exp Mol Pathol 2002;72:68–76. 25 Wu KK, Thiagarajan P. Role of endothelium in thrombosis and hemostasis. Annu Rev Med 1996;47: 315–331. 26 Dahlback B, Villoutreix BO. The anticoagulant pro- tein C pathway. FEBS Lett 2005;579:3310–3316. 27 Moore KL, Andreoli SP, Esmon NL, et al. Endotoxin enhances tissue factor and suppresses thrombomo- dulin expression of human vascular endothelium in vitro. J Clin Invest 1987;79:124–130. 28 Pawlinski R, Mackman N. Tissue factor, coagulation proteases, and protease-activated receptors in endo- toxemia and sepsis. Crit Care Med 2004;32:S293– S297. 29 Parry GC, Mackman N. Transcriptional regulation of tissue factor expression in human endothelial cells. Arterioscler Thromb Vasc Biol 1995;15:612–621. 30 Alexander C, Rietschel ET. Bacterial lipopolysacchar- ides and innate immunity. J Endotoxin Res 2001;7: 167–202. 31 Hack CE, Zeerleder S. The endothelium in sepsis: source of and a target for inflammation. Crit Care Med 2001;29:S21–S27. 32 Slack JL, Schooley K, Bonnert TP, et al. Identification of two major sites in the type I interleukin-1 receptor cytoplasmic region responsible for coupling to pro- inflammatory signaling pathways. J Biol Chem 2000; 275:4670–4678. 33 Tabeta K, Georgel P, Janssen E, et al. Toll-like receptors 9 and 3 as essential components of innate immune defense against mouse cytomegalovirus infection. Proc Natl Acad Sci USA 2004;101: 3516–3521. 34 Chuang TH, Ulevitch RJ. Cloning and characteriza- tion of a sub-family of human toll-like receptors: hTLR7, hTLR8 and hTLR9. Eur Cytokine Netw 2000;11:372–378. 35 Chuang T, Ulevitch RJ. Identification of hTLR10: a novel human Toll-like receptor preferentially expressed in immune cells. Biochim Biophys Acta 2001;1518:157–161. 36 Du X, Poltorak A, Wei Y, et al. Three novel mammalian toll-like receptors: gene structure, ex- pression, and evolution. Eur Cytokine Netw 2000; 11:362–371. 37 Poltorak A, He X, Smirnova I, et al. Defective LPS signaling in C3H/HeJ and C57BL/10ScCr mice: mutations in Tlr4 gene. Science 1998;282: 2085–2088. 38 Darveau RP, Pham TT, Lemley K, et al. Porphyromo- nas gingivalis lipopolysaccharide contains multiple lipid A species that functionally interact with both toll-like receptors 2 and 4. Infect Immun 2004;72: 5041–5051. LPS signaling in endothelial cells SM Dauphinee and A Karsan 18 Laboratory Investigation (2006) 86, 9–22 39 Schumann RR, Leong SR, Flaggs GW, et al. Structure and function of lipopolysaccharide binding protein. Science 1990;249:1429–1431. 40 Dunzendorfer S, Lee HK, Soldau K, et al. TLR4 is the signaling but not the lipopolysaccharide uptake receptor. J Immunol 2004;173:1166–1170. 41 da Silva Correia J, Soldau K, Christen U, et al. Lipopolysaccharide is in close proximity to each of the proteins in its membrane receptor complex. transfer from CD14 to TLR4 and MD-2. J Biol Chem 2001;276:21129–21135. 42 Wright SD. CD14 and innate recognition of bacteria. J Immunol 1995;155:6–8. 43 Arditi M, Zhou J, Dorio R, et al. Endotoxin-mediated endothelial cell injury and activation: role of soluble CD14. Infect Immun 1993;61:3149–3156. 44 Bufler P, Stiegler G, Schuchmann M, et al. Soluble lipopolysaccharide receptor (CD14) is released via two different mechanisms from human monocytes and CD14 transfectants. Eur J Immunol 1995;25: 604–610. 45 Jersmann HP, Hii CS, Hodge GL, et al. Synthesis and surface expression of CD14 by human endothelial cells. Infect Immun 2001;69:479–485. 46 Pugin J, Schurer-Maly CC, Leturcq D, et al. Lipopo- lysaccharide activation of human endothelial and epithelial cells is mediated by lipopolysaccharide- binding protein and soluble CD14. Proc Natl Acad Sci USA 1993;90:2744–2748. 47 Nagai Y, Akashi S, Nagafuku M, et al. Essential role of MD-2 in LPS responsiveness and TLR4 distribution. Nat Immunol 2002;3:667–672. 48 Visintin A, Latz E, Monks BG, et al. Lysines 128 and 132 enable lipopolysaccharide binding to MD-2, leading to Toll-like receptor-4 aggregation and signal transduction. J Biol Chem 2003;278:48313–48320. 49 Triantafilou M, Triantafilou K. Lipopolysaccharide recognition: CD14, TLRs and the LPS-activation cluster. Trends Immunol 2002;23:301–304. 50 Triantafilou K, Triantafilou M, Dedrick RL. A CD14- independent LPS receptor cluster. Nat Immunol 2001;2:338–345. 51 Dunzendorfer S, Lee HK, Soldau K, et al. Toll-like receptor 4 functions intracellularly in human coro- nary artery endothelial cells: roles of LBP and sCD14 in mediating LPS responses. FASEB J 2004;18: 1117–1119. 52 Hornef MW, Frisan T, Vandewalle A, et al. Toll-like receptor 4 resides in the Golgi apparatus and colocalizes with internalized lipopolysaccharide in intestinal epithelial cells. J Exp Med 2002;195: 559–570. 53 Hornef MW, Normark BH, Vandewalle A, et al. Intracellular recognition of lipopolysaccharide by toll-like receptor 4 in intestinal epithelial cells. J Exp Med 2003;198:1225–1235. 54 Shnyra A, Lindberg AA. Scavenger receptor pathway for lipopolysaccharide binding to Kupffer and en- dothelial liver cells in vitro. Infect Immun 1995; 63:865–873. 55 Latz E, Visintin A, Lien E, et al. Lipopolysaccharide rapidly traffics to and from the Golgi apparatus with the toll-like receptor 4-MD-2–CD14 complex in a process that is distinct from the initiation of signal transduction. J Biol Chem 2002;277:47834–47843. 56 Triantafilou M, Miyake K, Golenbock DT, et al. Mediators of innate immune recognition of bacteria concentrate in lipid rafts and facilitate lipopolysac- charide-induced cell activation. J Cell Sci 2002;115: 2603–2611. 57 Zhang H, Tay PN, Cao W, et al. Integrin-nucleated Toll-like receptor (TLR) dimerization reveals subcel- lular targeting of TLRs and distinct mechanisms of TLR4 activation and signaling. FEBS Lett 2002;532: 171–176. 58 Lee HK, Dunzendorfer S, Tobias PS. Cytoplasmic domain-mediated dimerizations of toll-like receptor 4 observed by beta-lactamase enzyme fragment com- plementation. J Biol Chem 2004;279:10564–10574. 59 Akira S, Takeda K. Toll-like receptor signalling. Nat Rev Immunol 2004;4:499–511. 60 Takeda K, Akira S. TLR signaling pathways. Semin Immunol 2004;16:3–9. 61 Wesche H, Henzel WJ, Shillinglaw W, et al. MyD88: an adapter that recruits IRAK to the IL-1 receptor complex. Immunity 1997;7:837–847. 62 O’Neill LA, Dunne A, Edjeback M, et al. Mal and MyD88: adapter proteins involved in signal transduc- tion by Toll-like receptors. J Endotoxin Res 2003;9: 55–59. 63 Li S, Strelow A, Fontana EJ, et al. IRAK-4: a novel member of the IRAK family with the properties of an IRAK-kinase. Proc Natl Acad Sci USA 2002;99: 5567–5572. 64 Fitzgerald KA, Palsson-McDermott EM, Bowie AG, et al. Mal (MyD88-adapter-like) is required for Toll- like receptor-4 signal transduction. Nature 2001;413: 78–83. 65 Yamamoto M, Sato S, Hemmi H, et al. Essential role for TIRAP in activation of the signalling cascade shared by TLR2 and TLR4. Nature 2002;420:324–329. 66 Bannerman DD, Erwert RD, Winn RK, et al. TIRAP mediates endotoxin-induced NF-kappaB activation and apoptosis in endothelial cells. Biochem Biophys Res Commun 2002;295:157–162. 67 Burns K, Janssens S, Brissoni B, et al. Inhibition of interleukin 1 receptor/Toll-like receptor signaling through the alternatively spliced, short form of MyD88 is due to its failure to recruit IRAK-4. J Exp Med 2003;197:263–268. 68 Suzuki N, Suzuki S, Duncan GS, et al. Severe impairment of interleukin-1 and Toll-like receptor signalling in mice lacking IRAK-4. Nature 2002;416: 750–756. 69 Cao Z, Xiong J, Takeuchi M, et al. TRAF6 is a signal transducer for interleukin-1. Nature 1996;383: 443–446. 70 Chung JY, Park YC, Ye H, et al. All TRAFs are not created equal: common and distinct molecular mechanisms of TRAF-mediated signal transduction. J Cell Sci 2002;115:679–688. 71 Jiang Z, Ninomiya-Tsuji J, Qian Y, et al. Interleukin-1 (IL-1) receptor-associated kinase-dependent IL-1- induced signaling complexes phosphorylate TAK1 and TAB2 at the plasma membrane and activate TAK1 in the cytosol. Mol Cell Biol 2002;22:7158–7167. 72 Shibuya H, Yamaguchi K, Shirakabe K, et al. TAB1: an activator of the TAK1 MAPKKK in TGF-beta signal transduction. Science 1996;272:1179–1182. 73 Takaesu G, Kishida S, Hiyama A, et al. TAB2, a novel adaptor protein, mediates activation of TAK1 MAPKKK by linking TAK1 to TRAF6 in the IL-1 signal transduction pathway. Mol Cell 2000;5: 649–658. LPS signaling in endothelial cells SM Dauphinee and A Karsan 19 Laboratory Investigation (2006) 86, 9–22 74 Kishida S, Sanjo H, Akira S, et al. TAK1-binding protein 2 facilitates ubiquitination of TRAF6 and assembly of TRAF6 with IKK in the IL-1 signaling pathway. Genes Cells 2005;10:447–454. 75 Cheung PC, Nebreda AR, Cohen P. TAB3, a new binding partner of the protein kinase TAK1. Biochem J 2004;378:27–34. 76 Deng L, Wang C, Spencer E, et al. Activation of the IkappaB kinase complex by TRAF6 requires a dimeric ubiquitin-conjugating enzyme complex and a unique polyubiquitin chain. Cell 2000;103: 351–361. 77 Yamin TT, Miller DK. The interleukin-1 receptor- associated kinase is degraded by proteasomes following its phosphorylation. J Biol Chem 1997;272: 21540–21547. 78 Jensen LE, Whitehead AS. Ubiquitin activated tumor necrosis factor receptor associated factor-6 (TRAF6) is recycled via deubiquitination. FEBS Lett 2003;553: 190–194. 79 Wang C, Deng L, Hong M, et al. TAK1 is a ubiquitin- dependent kinase of MKK and IKK. Nature 2001;412: 346–351. 80 Ea CK, Sun L, Inoue J, et al. TIFA activates IkappaB kinase (IKK) by promoting oligomerization and ubiquitination of TRAF6. Proc Natl Acad Sci USA 2004;101:15318–15323. 81 Chen Z, Hagler J, Palombella VJ, et al. Signal-induced site-specific phosphorylation targets I kappa B alpha to the ubiquitin–proteasome pathway. Genes Dev 1995;9:1586–1597. 82 Zen K, Karsan A, Eunson T, et al. Lipopolysacchar- ide-inducedNFkappaB activation in human endothelial cells involves degradation of IkappaBalpha but not IkappaBbeta. Exp Cell Res 1998;243:425–433. 83 Asehnoune K, Strassheim D, Mitra S, et al. Involve- ment of PKCalpha/beta in TLR4 and TLR2 dependent activation of NF-kappaB. Cell Signal 2005;17: 385–394. 84 Dallot E, Mehats C, Oger S, et al. A role for PKCzeta in the LPS-induced translocation NF-kappaB p65 sub- unit in cultured myometrial cells. Biochimie 2005;87: 513–521. 85 Wymann MP, Pirola L. Structure and function of phosphoinositide 3-kinases. Biochim Biophys Acta 1998;1436:127–150. 86 Franke TF, Yang SI, Chan TO, et al. The protein kinase encoded by the Akt proto-oncogene is a target of the PDGF-activated phosphatidylinositol 3-kinase. Cell 1995;81:727–736. 87 Fruman DA, Snapper SB, Yballe CM, et al. Impaired B cell development and proliferation in absence of phosphoinositide 3-kinase p85alpha. Science 1999; 283:393–397. 88 Ojaniemi M, Glumoff V, Harju K, et al. Phosphatidy- linositol 3-kinase is involved in Toll-like receptor 4- mediated cytokine expression in mouse macrophages. Eur J Immunol 2003;33:597–605. 89 Li X, Tupper JC, Bannerman DD, et al. Phosphoinosi- tide 3 kinase mediates Toll-like receptor 4-induced activation of NF-kappa B in endothelial cells. Infect Immun 2003;71:4414–4420. 90 Wong F, Hull C, Zhande R, et al. Lipopolysaccharide initiates a TRAF6-mediated endothelial survival signal. Blood 2004;103:4520–4526. 91 Vivarelli MS, McDonald D, Miller M, et al. RIP links TLR4 to Akt and is essential for cell survival in response to LPS stimulation. J Exp Med 2004;200: 399–404. 92 McCarthy JV, Ni J, Dixit VM. RIP2 is a novel NF- kappaB-activating and cell death-inducing kinase. J Biol Chem 1998;273:16968–16975. 93 Chin AI, Dempsey PW, Bruhn K, et al. Involvement of receptor-interacting protein 2 in innate and adaptive immune responses. Nature 2002;416:190–194. 94 Lu C, Wang A, Dorsch M, et al. Participation of Rip2 in lipopolysaccharide signaling is independent of its kinase activity. J Biol Chem 2005;280:16278–16283. 95 Huang Q, Yang J, Lin Y, et al. Differential regulation of interleukin 1 receptor and Toll-like receptor signaling by MEKK3. Nat Immunol 2004;5:98–103. 96 Hull C, McLean G, Wong F, et al. Lipopolysaccharide signals an endothelial apoptosis pathway through TNF receptor-associated factor 6-mediated activation of c-Jun NH2-terminal kinase. J Immunol 2002;169: 2611–2618. 97 Matsuzawa A, Saegusa K, Noguchi T, et al. ROS- dependent activation of the TRAF6–ASK1–p38 path- way is selectively required for TLR4-mediated innate immunity. Nat Immunol 2005;6:587–592. 98 Dumitru CD, Ceci JD, Tsatsanis C, et al. TNF-alpha induction by LPS is regulated posttranscriptionally via a Tpl2/ERK-dependent pathway. Cell 2000;103: 1071–1083. 99 Patriotis C, Makris A, Chernoff J, et al. Tpl-2 acts in concert with Ras and Raf-1 to activate mitogen- activated protein kinase. Proc Natl Acad Sci USA 1994;91:9755–9759. 100 Guha M, O’Connell MA, Pawlinski R, et al. Lipopo- lysaccharide activation of the MEK-ERK1/2 pathway in human monocytic cells mediates tissue factor and tumor necrosis factor alpha expression by inducing Elk-1 phosphorylation and Egr-1 expression. Blood 2001;98:1429–1439. 101 Matsuguchi T, Masuda A, Sugimoto K, et al. JNK- interacting protein 3 associates with Toll-like recep- tor 4 and is involved in LPS-mediated JNK activation. EMBO J 2003;22:4455–4464. 102 Arndt PG, Suzuki N, Avdi NJ, et al. Lipopolysacchar- ide-induced c-Jun NH2-terminal kinase activation in human neutrophils: role of phosphatidylinositol 3- kinase and Syk-mediated pathways. J Biol Chem 2004;279:10883–10891. 103 Pollet I, Opina CJ, Zimmerman C, et al. Bacterial lipopolysaccharide directly induces angiogenesis through TRAF6-mediated activation of NF-kappaB and c-Jun N-terminal kinase. Blood 2003;102: 1740–1742. 104 Haimovitz-Friedman A, Cordon-Cardo C, Bayoumy S, et al. Lipopolysaccharide induces disseminated endothelial apoptosis requiring ceramide generation. J Exp Med 1997;186:1831–1841. 105 Kawai T, Adachi O, Ogawa T, et al. Unresponsiveness of MyD88-deficient mice to endotoxin. Immunity 1999;11:115–122. 106 Kawai T, Takeuchi O, Fujita T, et al. Lipopolysacchar- ide stimulates the MyD88-independent pathway and results in activation of IFN-regulatory factor 3 and the expression of a subset of lipopolysaccharide-induci- ble genes. J Immunol 2001;167:5887–5894. 107 Hoshino K, Kaisho T, Iwabe T, et al. Differential involvement of IFN-beta in Toll-like receptor-stimu- lated dendritic cell activation. Int Immunol 2002; 14:1225–1231. LPS signaling in endothelial cells SM Dauphinee and A Karsan 20 Laboratory Investigation (2006) 86, 9–22 108 Oshiumi H, Sasai M, Shida K, et al. TIR-containing adapter molecule (TICAM)-2, a bridging adapter recruiting to toll-like receptor 4 TICAM-1 that induces interferon-beta. J Biol Chem 2003;278: 49751–49762. 109 Yamamoto M, Sato S, Hemmi H, et al. TRAM is specifically involved in the Toll-like receptor 4- mediated MyD88-independent signaling pathway. Nat Immunol 2003;4:1144–1150. 110 Yamamoto M, Sato S, Hemmi H, et al. Role of adaptor TRIF in the MyD88-independent toll-like receptor signaling pathway. Science 2003;301:640–643. 111 Kaiser WJ, Offermann MK. Apoptosis induced by the toll-like receptor adaptor TRIF is dependent on its receptor interacting protein homotypic interaction motif. J Immunol 2005;174:4942–4952. 112 Fitzgerald KA, McWhirter SM, Faia KL, et al. IKKepsilon and TBK1 are essential components of the IRF3 signaling pathway. Nat Immunol 2003;4: 491–496. 113 Taniguchi T, Ogasawara K, Takaoka A, et al. IRF family of transcription factors as regulators of host defense. Annu Rev Immunol 2001;19:623–655. 114 Sasai M, Oshiumi H, Matsumoto M, et al. Cutting edge: NF-kappaB-activating kinase-associated protein 1 participates in TLR3/Toll-IL-1 homology domain- containing adapter molecule-1-mediated IFN regula- tory factor 3 activation. J Immunol 2005;174:27–30. 115 Lentschat A, Karahashi H, Michelsen KS, et al. Mastoparan, a G protein agonist peptide, differen- tially modulates TLR4- and TLR2-mediated signaling in human endothelial cells and murine macrophages. J Immunol 2005;174:4252–4261. 116 Grimm J, Sachs M, Britsch S, et al. Novel p62dok family members, dok-4 and dok-5, are substrates of the c-Ret receptor tyrosine kinase and mediate neuronal differentiation. J Cell Biol 2001;154: 345–354. 117 Yamanashi Y, Baltimore D. Identification of the Abl- and rasGAP-associated 62 kDa protein as a docking protein, Dok. Cell 1997;88:205–211. 118 Di Cristofano A, Carpino N, Dunant N, et al. Molecular cloning and characterization of p56dok-2 defines a new family of RasGAP-binding proteins. J Biol Chem 1998;273:4827–4830. 119 Lemay S, Davidson D, Latour S, et al. Dok-3, a novel adapter molecule involved in the negative regulation of immunoreceptor signaling. Mol Cell Biol 2000; 20:2743–2754. 120 Shinohara H, Inoue A, Toyama-Sorimachi N, et al. Dok-1 and Dok-2 are negative regulators of lipopoly- saccharide-induced signaling. J Exp Med 2005;201: 333–339. 121 Bedirian A, Baldwin C, Abe J, et al. Pleckstrin homology and phosphotyrosine-binding domain- dependent membrane association and tyrosine phos- phorylation of Dok-4, an inhibitory adapter molecule expressed in epithelial cells. J Biol Chem 2004;279: 19335–19349. 122 Itoh S, Lemay S, Osawa M, et al. Mitochondrial Dok-4 recruits Src kinase and regulates NF-{kappa}B activa- tion in endothelial cells. J Biol Chem 2005;280: 26383–26396. 123 Pahan K, Raymond JR, Singh I. Inhibition of phos- phatidylinositol 3-kinase induces nitric-oxide syn- thase in lipopolysaccharide- or cytokine-stimulated C6 glial cells. J Biol Chem 1999;274:7528–7536. 124 Martin M, Schifferle RE, Cuesta N, et al. Role of the phosphatidylinositol 3 kinase–Akt pathway in the regulation of IL-10 and IL-12 by Porphyromonas gingivalis lipopolysaccharide. J Immunol 2003;171: 717–725. 125 Schabbauer G, Tencati M, Pedersen B, et al. PI3K–Akt pathway suppresses coagulation and inflammation in endotoxemic mice. Arterioscler Thromb Vasc Biol 2004;24:1963–1969. 126 Fukao T, Tanabe M, Terauchi Y, et al. PI3K-mediated negative feedback regulation of IL-12 production in DCs. Nat Immunol 2002;3:875–881. 127 Guha M, Mackman N. The phosphatidylinositol 3- kinase–Akt pathway limits lipopolysaccharide acti- vation of signaling pathways and expression of inflammatory mediators in human monocytic cells. J Biol Chem 2002;277:32124–32132. 128 Pahan K, Liu X, Wood C, et al. Expression of a constitutively active form of phosphatidylinositol 3- kinase inhibits the induction of nitric oxide synthase in human astrocytes. FEBS Lett 2000;472:203–207. 129 Strassheim D, Asehnoune K, Park JS, et al. Phospho- inositide 3-kinase and Akt occupy central roles in inflammatory responses of Toll-like receptor 2-stimu- lated neutrophils. J Immunol 2004;172:5727–5733. 130 Aksoy E, Vanden Berghe W, Detienne S, et al. Inhibition of phosphoinositide 3-kinase enhances TRIF-dependent NF-kappaB activation and IFN-beta synthesis downstream of Toll-like receptor 3 and 4. Eur J Immunol 2005;35:2200–2209. 131 Opipari Jr AW, Hu HM, Yabkowitz R, et al. The A20 zinc finger protein protects cells from tumor necrosis factor cytotoxicity. J Biol Chem 1992;267: 12424–12427. 132 Wertz IE, O’Rourke KM, Zhou H, et al. De-ubiquitina- tion and ubiquitin ligase domains of A20 down- regulate NF-kappaB signalling. Nature 2004;430: 694–699. 133 Evans PC, Ovaa H, Hamon M, et al. Zinc-finger protein A20, a regulator of inflammation and cell survival, has de-ubiquitinating activity. Biochem J 2004;378:727–734. 134 Hu X, Yee E, Harlan JM, et al. Lipopolysaccharide induces the antiapoptotic molecules, A1 and A20, in microvascular endothelial cells. Blood 1998;92: 2759–2765. 135 Lee EG, Boone DL, Chai S, et al. Failure to regulate TNF-induced NF-kappaB and cell death responses in A20-deficient mice. Science 2000;289:2350–2354. 136 O’Reilly SM, Moynagh PN. Regulation of Toll-like receptor 4 signalling by A20 zinc finger protein. Biochem Biophys Res Commun 2003;303:586–593. 137 Boone DL, Turer EE, Lee EG, et al. The ubiquitin- modifying enzyme A20 is required for termination of Toll-like receptor responses. Nat Immunol 2004;5: 1052–1060. 138 Chuang PI, Yee E, Karsan A, et al. A1 is a constitutive and inducible Bcl-2 homologue in mature human neutrophils. Biochem Biophys Res Commun 1998; 249:361–365. 139 Stroka DM, Badrichani AZ, Bach FH, et al. Over- expression of A1, an NF-kappaB-inducible anti- apoptotic bcl gene, inhibits endothelial cell activa- tion. Blood 1999;93:3803–3810. 140 Rocha E, Mahiou J, Badrichani AZ, et al. The BH4 domain of A1, an anti-apoptotic bcl family gene, is necessary and sufficient for its antiinflammatory LPS signaling in endothelial cells SM Dauphinee and A Karsan 21 Laboratory Investigation (2006) 86, 9–22 function in endothelial cells. Transplant Proc 2001; 33:314. 141 Chinnaiyan AM, O’Rourke K, Tewari M, et al. FADD, a novel death domain-containing protein, interacts with the death domain of Fas and initiates apoptosis. Cell 1995;81:505–512. 142 Choi KB, Wong F, Harlan JM, et al. Lipopolysacchar- ide mediates endothelial apoptosis by a FADD- dependent pathway. J Biol Chem 1998;273:20185– 20188. 143 Bannerman DD, Tupper JC, Kelly JD, et al. The Fas- associated death domain protein suppresses activa- tion of NF-kappa B by LPS and IL-1 beta. J Clin Invest 2002;109:419–425. 144 Aliprantis AO, Yang RB, Weiss DS, et al. The apoptotic signaling pathway activated by Toll-like receptor-2. EMBO J 2000;19:3325–3336. 145 Burns K, Clatworthy J, Martin L, et al. Tollip, a new component of the IL-1RI pathway, links IRAK to the IL-1 receptor. Nat Cell Biol 2000;2:346–351. 146 Zhang G, Ghosh S. Negative regulation of toll-like receptor-mediated signaling by Tollip. J Biol Chem 2002;277:7059–7065. 147 Bulut Y, Faure E, Thomas L, et al. Cooperation of Toll- like receptor 2 and 6 for cellular activation by soluble tuberculosis factor and Borrelia burgdorferi outer surface protein A lipoprotein: role of Toll-interacting protein and IL-1 receptor signaling molecules in Toll-like receptor 2 signaling. J Immunol 2001;167: 987–994. 148 Lioubin MN, Algate PA, Tsai S, et al. p150Ship, a signal transduction molecule with inositol polyphos- phate-5-phosphatase activity. Genes Dev 1996;10: 1084–1095. 149 Sly LM, Rauh MJ, Kalesnikoff J, et al. LPS-induced upregulation of SHIP is essential for endotoxin tolerance. Immunity 2004;21:227–239. 150 An H, Xu H, Zhang M, et al. Src homology 2 domain- containing inositol-5-phosphatase 1 (SHIP1) nega- tively regulates TLR4-mediated LPS response pr- imarily through a phosphatase activity- and PI-3K- independent mechanism. Blood 2005;105:4685–4692. 151 Zippo A, De Robertis A, Bardelli M, et al. Identifica- tion of Flk-1 target genes in vasculogenesis: Pim-1 is required for endothelial and mural cell differentiation in vitro. Blood 2004;103:4536–4544. 152 Moritz DR, Rodewald HR, Gheyselinck J, et al. The IL-1 receptor-related T1 antigen is expressed on immature and mature mast cells and on fetal blood mast cell progenitors. J Immunol 1998;161:4866–4874. 153 Xu D, Chan WL, Leung BP, et al. Selective expression of a stable cell surface molecule on type 2 but not type 1 helper T cells. J Exp Med 1998;187:787–794. 154 Brint EK, Xu D, Liu H, et al. ST2 is an inhibitor of interleukin 1 receptor and Toll-like receptor 4 signal- ing and maintains endotoxin tolerance. Nat Immunol 2004;5:373–379. 155 Liew FY, Liu H, Xu D. A novel negative regulator for IL-1 receptor and Toll-like receptor 4. Immunol Lett 2005;96:27–31. 156 Kumar S, Minnich MD, Young PR. ST2/T1 protein functionally binds to two secreted proteins from Balb/c 3T3 and human umbilical vein endothelial cells but does not bind interleukin 1. J Biol Chem 1995;270:27905–27913. 157 Yoshimura A, Nishinakamura H, Matsumura Y, et al. Negative regulation of cytokine signaling and immune responses by SOCS proteins. Arthritis Res Ther 2005;7:100–110. 158 Nakagawa R, Naka T, Tsutsui H, et al. SOCS-1 participates in negative regulation of LPS responses. Immunity 2002;17:677–687. 159 Kinjyo I, Hanada T, Inagaki-Ohara K, et al. SOCS1/ JAB is a negative regulator of LPS-induced macro- phage activation. Immunity 2002;17:583–591. 160 Wang X, Athayde N, Trudinger B. Fetal plasma stimulates endothelial cell production of cytokines and the family of suppressor of cytokine signaling in umbilical placental vascular disease. Am J Obstet Gynecol 2003;188:510–516. 161 Divanovic S, Trompette A, Atabani SF, et al. Negative regulation of Toll-like receptor 4 signaling by the Toll- like receptor homolog RP105. Nat Immunol 2005; 6:571–578. 162 Miyake K, Shimazu R, Kondo J, et al. Mouse MD-1, a molecule that is physically associated with RP105 and positively regulates its expression. J Immunol 1998;161:1348–1353. 163 Hijiya N, Miyake K, Akashi S, et al. Possible involvement of toll-like receptor 4 in endothelial cell activation of larger vessels in response to lipopoly- saccharide. Pathobiology 2002;70:18–25. 164 Kishore SP, Bungum MK, Platt JL, et al. Selec- tive suppression of Toll-like receptor 4 activation by chemokine receptor 4. FEBS Lett 2005;579: 699–704. 165 Murdoch C, Monk PN, Finn A. Cxc chemokine receptor expression on human endothelial cells. Cytokine 1999;11:704–712. 166 Wald D, Qin J, Zhao Z, et al. SIGIRR, a negative regulator of Toll-like receptor-interleukin 1 receptor signaling. Nat Immunol 2003;4:920–927. 167 Qin J, Qian Y, Yao J, et al. SIGIRR inhibits interleukin- 1 receptor- and toll-like receptor 4-mediated signaling through different mechanisms. J Biol Chem 2005;280: 25233–25241. 168 Chuang TH, Ulevitch RJ. Triad3A, an E3 ubiquitin- protein ligase regulating Toll-like receptors. Nat Immunol 2004;5:495–502. 169 Janssens S, Burns K, Tschopp J, et al. Regulation of interleukin-1- and lipopolysaccharide-induced NF- kappaB activation by alternative splicing of MyD88. Curr Biol 2002;12:467–471. 170 Janssens S, Burns K, Vercammen E, et al. MyD88S, a splice variant of MyD88, differentially modulates NF- kappaB- and AP-1-dependent gene expression. FEBS Lett 2003;548:103–107. 171 Wesche H, Gao X, Li X, et al. IRAK-M is a novel member of the Pelle/interleukin-1 receptor-associated kinase (IRAK) family. J Biol Chem 1999;274:19403– 19410. 172 Kobayashi K, Hernandez LD, Galan JE, et al. IRAK-M is a negative regulator of Toll-like receptor signaling. Cell 2002;110:191–202. 173 Fritz JH, Girardin SE, Fitting C, et al. Synergistic stimulation of human monocytes and dendritic cells by Toll-like receptor 4 and NOD1- and NOD2- activating agonists. Eur J Immunol 2005;35:2459– 2470. 174 Naiki Y, Michelsen KS, Zhang W, et al. Transforming growth factor-beta differentially inhibits MyD88- dependent, but not TRAM- and TRIF-dependent, lipopolysaccharide-induced TLR4 signaling. J Biol Chem 2005;280:5491–5495. LPS signaling in endothelial cells SM Dauphinee and A Karsan 22 Laboratory Investigation (2006) 86, 9–22 MOLECULAR AND CELLULAR BIOLOGY, Nov. 2007, p. 7394–7404 Vol. 27, No. 21 0270-7306/07/$08.000 doi:10.1128/MCB.00600-07 Copyright © 2007, American Society for Microbiology. All Rights Reserved. FADD Negatively Regulates Lipopolysaccharide Signaling by Impairing Interleukin-1 Receptor-Associated Kinase 1–MyD88 Interaction Rachel Zhande,1,2 Shauna M. Dauphinee,1,4 James A. Thomas,5 Masahiro Yamamoto,6 Shizuo Akira,6 and Aly Karsan1,2,3,4* Department of Medical Biophysics, British Columbia Cancer Agency,1 Department of Pathology and Laboratory Medicine, University of British Columbia,2 Department of Pathology and Laboratory Medicine, British Columbia Cancer Agency,3 and Experimental Medicine Program, University of British Columbia,4 Vancouver, BC, Canada; Departments of Pediatrics, Molecular Biology, and Oncology, University of Texas Southwestern Medical Center, Dallas, Texas5; and Department of Host Defense, Research Institute for Microbial Diseases, Osaka University, and ERATO, Japan Science and Technology Agency, Osaka, Japan6 Received 5 April 2007/Returned for modification 11 May 2007/Accepted 25 August 2007 Lipopolysaccharide (LPS) engages Toll-like receptor 4 (TLR4) on various cells to initiate inflammatory and angiogenic pathways. FADD is an adaptor protein involved in death receptor-mediated apoptosis. Here we report a role for FADD in regulation of TLR4 signals in endothelial cells. FADD specifically attenuates LPS-induced activation of c-Jun NH2-terminal kinase and phosphatidylinositol 3-kinase in a death domain- dependent manner. In contrast, FADD-null cells show hyperactivation of these kinases. Examining physical associations of endogenous proteins, we show that FADD interacts with interleukin-1 receptor-associated kinase 1 (IRAK1) and MyD88. LPS stimulation increases IRAK1-FADD interaction and recruitment of the IRAK1-FADD complex to activated MyD88. IRAK1 is required for FADD-MyD88 interaction, as FADD does not associate with MyD88 in IRAK1-null cells. By shuttling FADD to MyD88, IRAK1 provides a mechanism for controlled and limited activation of the TLR4 signaling pathway. Functionally, enforced FADD expression inhibited LPS- but not vascular endothelial growth factor-induced endothelial cell sprouting, while FADD deficiency led to enhanced production of proinflammatory cytokines induced by stimulation of TLR4 and TLR2, but not TLR3. Reconstitution of FADD reversed the enhanced production of proinflammatory cytokines. Thus, FADD is a physiological negative regulator of IRAK1/MyD88-dependent responses in innate immune signaling. Bacterial lipopolysaccharide (LPS) is a potent inflammatory molecule that evokes immune responses by activating various cell types, including endothelial cells, through Toll-like recep- tor 4 (TLR4). The cytoplasmic portions of all TLRs as well as that of the interleukin-1 receptor (IL-1R) share a highly con- served region known as the Toll–IL-1R (TIR) domain. Upon receptor occupancy, the TIR domain recruits several TIR do- main-containing intracellular proteins (MyD88, TIRAP/Mal (MyD88 adaptor-like protein), TRIF (Toll/IL-1R domain-con- taining adaptor-inducing beta interferon)/TICAM-1 (TIR-con- taining adapter molecule 2 [TICAM-2]), and TRAM (TRIF- related adaptor molecule)/TICAM-2 (13, 15, 27, 30, 34) via homophilic interactions. Signaling downstream from most TLRs is dependent on the adaptor protein MyD88. In addition to MyD88 and Mal, TLR4 also recruits TRIF and TRAM to the receptor. In contrast, TLR3 does not signal through MyD88 but rather recruits only TRIF (12). The TRIF-depen- dent pathway activates the transcription factors NF-B and IRF3 (interferon regulatory factor 3) leading to subsequent induction of genes, such as beta interferon, IP-10 (interferon- inducible protein of 10 kDa), and RANTES (12). However, the best-characterized TLR adaptor protein is MyD88, which is required for the initial rapid activation of mitogen-activated protein kinases (MAPKs) and NF-B. In addition to the TIR domain, MyD88 contains a death do- main (DD), a highly conserved protein interaction domain, which enables it to recruit other DD-containing signaling transducers involved in innate immune signaling called in- terleukin-1 receptor-associated kinases (IRAKs). Upon re- cruitment by MyD88, interleukin-1 receptor-associated ki- nase 4 (IRAK4) phosphorylates IRAK1, triggering IRAK1 auto- and trans-phosphorylation. Hyperphosphorylated IRAK1 dissociates from the TLR4-MyD88 complex and as- sociates with tumor necrosis factor (TNF) receptor-acti- vated factor 6 (TRAF6), which triggers the oligomerization and polyubiquitination of TRAF6. Functionally, TRAF6 is an E3 ubiquitin ligase which in collaboration with the di- meric ubiquitin-conjugating enzyme (E2) complex Ubc13/ Uev1A catalyzes autologous synthesis of lysine 63 (K63)- linked polyubiquitin chains (10). The K63 ubiquitin chains are recognized by the ubiquitin acceptor proteins transform- ing growth factor -activating kinase 1-binding protein 2 and 3 (TAB2 and TAB3) which function as adaptors to assemble a signaling complex which phosphorylates and ac- tivates transforming growth factor -activated kinase 1 * Corresponding author. Mailing address: Department of Medical Biophysics, British Columbia Cancer Research Centre, 675 West 10th Ave., Vancouver, BC V5Z 1L3, Canada. Phone: (604) 675-8033. Fax: (604) 675-8049. E-mail: akarsan@bccrc.ca.  Published ahead of print on 4 September 2007. 7394  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  (TAK1) (21). Activated TAK1 leads to the activation of the IB kinase (IKK) complex, NF-B, and the MAPK kinases that activate p38, extracellular signal-regulated (ERK) and c-Jun NH2-terminal protein kinase (JNK). LPS also acti- vates the phosphatidylinositol 3-kinase (PI3K) pathway downstream of TRAF6 (33). FADD is a DD-containing adaptor protein that is recruited to death receptors following receptor occupancy. FADD binds directly to Fas/CD95 and to the adaptor protein TRADD in response to Fas ligand and TNF, respectively, to activate caspase 8 and initiate apoptosis (16, 28). Interestingly, others have suggested that TLR2 recruits FADD via MyD88 and thereby also induces apoptosis (2). However, the latter study examined only FADD-MyD88 associations in overexpression systems. Whether FADD is involved in TLR4 signaling to activate the MAPK and PI3K pathways by LPS has not been demonstrated. We report here that FADD, in a manner re- quiring the DD, negatively regulates LPS signaling by sup- pressing activation of JNK and PI3K pathways. FADD inter- acts with IRAK1 and MyD88 and appears to reduce the stability of the MyD88-IRAK1 interaction, thereby attenuating the LPS signal. IRAK1 is essential to allow the MyD88-FADD interaction, as MyD88 and FADD do not associate in IRAK1- null cells. FADD also inhibits IRAK1-MyD88-dependent sig- naling in response to activation of other receptors, such as TLR2 and IL-1R. We demonstrate further that enforced ex- pression of FADD plays a role in deregulating endothelial function by limiting endothelial sprouting in response to LPS, but not vascular endothelial growth factor (VEGF). Addition- ally, FADD-deficient cells show significantly enhanced cyto- kine release in response to TLR4 and TLR2 agonists, but not in response to TLR3. Thus, FADD represents a novel negative regulator of IRAK1/MyD88-dependent innate immune re- sponses. MATERIALS AND METHODS Reagents. LPS and anti-Flag M2 antibody were purchased from Sigma (St. Louis, MO). Anti-JNK1, antiubiquitin, and anti-TRAF6 antibodies were from Santa Cruz Biotechnology (Santa Cruz, CA). The phospho-specific antibodies against JNK and Akt and anti-Akt antibody were purchased from Cell Signaling Technology (Beverly, MA). Protein A and G agarose, anti-IRAK1, anti-IRAK4, and anti-FADD monoclonal and polyclonal antibodies were obtained from Up- state (Charlottesville, VA). Anti-MyD88 antibody was purchased from Alexis Biochemicals (Lausanne, Switzerland). TrueBlot anti-rabbit immunoglobulin im- munoprecipitation beads and rabbit immunoglobulin G (IgG) TrueBlot were obtained from eBioscience (San Diego, CA). Recombinant TNF alpha was purchased from R&D Systems. Poly(I)  poly(C) and the synthetic tripalmitoy- lated lipopeptide Pam3CSK4 were purchased from InvivoGen (San Diego, CA). Cell culture. Human microvascular endothelial cells (HMEC) were provided by the Centers for Disease Control and Prevention and were cultured in MCDB131 medium supplemented with 10% calf serum. 293T cells and mouse embryonic fibroblasts (MEF) were cultured in Dulbecco modified Eagle medium supplemented with 10% calf serum. FADD-deficient MEF were obtained from Amgen, Inc. (Thousand Oaks, CA), and IRAK1-deficient and MyD88-deficient MEF have previously been described (1, 32). Recombinant plasmids and transfection. Full-length FADD containing a Flag epitope at the N terminus was cloned into the murine stem cell virus-internal ribosome entry site-yellow fluorescent protein (MSCV-IRES-YFP) (MIY) ret- roviral vector. Transient transfections of the Ampho Phoenix packaging cell line were carried out using Fugene 6. Viral supernatants were used to transduce HMEC, and yellow fluorescent protein-positive cells were selected by flow sort- ing. 293T cells were transfected with Fugene 6 according to the manufacturer’s instructions (Roche Applied Science Laval, QC, Canada). Coimmunoprecipitation, immunoblotting, and kinase assays. HMEC, 293T cells, or MEF were lysed for 30 min in 20 mM HEPES (pH 7.4), 150 mM NaCl, 12.5 mM -glycerol phosphate, 1.5 mM MgCl2, 10 mM NaF, 1 mM sodium orthovanadate, 2 mM EGTA, 0.5% Triton X-100, and protease inhibitor cocktail to examine noncovalent interactions. Antibodies (3 g) were added to cell lysates (8 mg) for 3 h at 4°C and captured by the addition of protein A or G or TrueBlot anti-rabbit immunoglobulin immunoprecipitation beads for an additional 12 h at 4°C. The immune complexes were washed three times with lysis buffer followed by the addition of sodium dodecyl sulfate (SDS) sample buffer. The bound proteins were separated by SDS-polyacrylamide gel electrophoresis, transferred to nitrocellulose membranes, and analyzed by immunoblotting. HMEC treated without or with LPS were lysed and immunoprecipitated under denaturing con- ditions (50 mM Tris, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% SDS, 20 mM N-ethylmaleimide and protease inhibitors) to examine cova- lent polyubiquitination of TRAF6. Cell lysates (3 mg) were immunoprecipitated with antibody against TRAF6 (5 l) for 3 h followed by an additional 12 h of incubation in the presence of TrueBlot anti-rabbit immunoglobulin immunopre- cipitation beads. Immunoprecipitates were washed three times with lysis buffer followed by the addition of SDS sample buffer. Proteins were separated on SDS-polyacrylamide gels and subjected to immunoblotting with antiubiquitin and TRAF6 antibodies. For JNK activity, cells were exposed to LPS (100 ng/ml) at the times indicated. At each time point, cells were lysed in buffer containing protease and phosphatase inhibitors, and lysates were obtained by centrifugation. The assay is dependent upon the isolation of JNK with glutathione S-transferase (GST)–c-Jun followed by an in vitro kinase assay using GST–c-Jun as the sub- strate in the presence of -32P-labeled ATP or nonradioactive ATP. The kinase reaction mixtures were then separated on SDS-polyacrylamide gels and sub- jected to autoradiography (in the former case). For the latter case, the separated proteins were transferred to nitrocellulose membranes and analyzed by immu- noblotting with a phospho-specific c-Jun (Ser 73) antibody. ELISA. MEF were seeded in 24-well plates at a density of 200,000 cells per well and cultured for 24 h. Following treatment, the supernatants were recovered and centrifuged (850  g, 10 min). The supernatants were analyzed using com- mercially available kits for murine IL-6 (eBioscience) and IP-10 (R&D Systems). Endothelial sprouting assay. Endothelial sprouting was assessed as previously described (25). Briefly, microcarrier beads coated with gelatin (Cytodex 3; Sigma) were seeded with HMEC lines and embedded in fibrin gels in 96-well plates. Fibrin gels were supplemented with VEGF (15 ng/ml) or LPS (100 ng/ml). The overlying medium contained either MCDB131 medium supplemented with 2% fetal bovine serum alone or was further supplemented with VEGF (15 ng/ml) or LPS (100 ng/ml). After 2 days of incubation with daily medium changes, the number of capillary-like tubes formed was quantitated by counting the number of tube-like structures longer than 150 m. RESULTS FADD inhibits activation of MAPKs and Akt by LPS. We first investigated the role of FADD on LPS-induced activation of JNK. Human microvascular endothelial cells were stably transduced with either full-length FADD (HMEC-FADD) or the empty vector MIY (HMEC-MIY), and expression of FADD was confirmed by immunoblotting (Fig. 1E). Cells were stimulated with LPS, and JNK activity was measured by mon- itoring the phosphorylation of c-Jun. In HMEC transduced with the empty vector, c-Jun phosphorylation was apparent at 30 min after LPS stimulation, whereas FADD-overexpressing cells exhibited a 30-min lag in c-Jun phosphorylation (Fig. 1A). Additionally, at all time points, the magnitude of c-Jun phos- phorylation in FADD-overexpressing cells was considerably lower than in the control cells. LPS and TNF signaling path- ways share some similarities, such as the activation of MAPKs, including JNK. Moreover, FADD is a critical signaling adaptor molecule which bridges the TNF receptor to the extrinsic apoptotic pathway. We thus tested whether overexpression of FADD also intersected the TNF pathway to JNK activation. As shown in Fig. 1A, TNF treatment led to a similar pattern of c-Jun phosphorylation between FADD-overexpressing and VOL. 27, 2007 FADD NEGATIVELY REGULATES LPS SIGNALING 7395  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  vector-transduced cells. Therefore, FADD-mediated attenua- tion of JNK activation is specific to the LPS pathway. The effect of FADD on JNK activity could arise either from the inhibition of JNK phosphorylating activity directly or through inhibition of JNK activation by upstream effectors. In order to determine whether FADD was inhibiting JNK directly or indirectly via a different molecule, we determined the phos- phorylation status of JNK. Figure 1B shows the phosphoryla- tion of JNK as determined by a JNK phospho-specific anti- body. Overexpression of FADD diminished LPS-induced phosphorylation of JNK, suggesting that FADD does not in- terfere with JNK activity directly but rather targets upstream molecules involved in JNK activation by LPS. In addition to JNK, LPS simultaneously activates the PI3K pathway. LPS-stimulated phosphorylation of Akt at Ser 473, a downstream effect of the activation of PI3K, was impaired in cells overexpressing FADD (Fig. 1C). In contrast to LPS, TNF- mediated activation of JNK was not inhibited in cells overex- pressing FADD (Fig. 1D). LPS-induced activation of ERK1/2 MAPKs was also suppressed by enforced expression of FADD FIG. 1. Enforced expression of FADD inhibits JNK and Akt activation by LPS but not TNF. (A) Endothelial cells expressing full-length FADD (HMEC-FADD) or empty vector (HMEC-MIY) were left untreated or treated with TNF (10 ng/ml) or LPS (100 ng/ml) for the indicated times (in minutes). Cell lysates were subjected to an in vitro JNK protein kinase assay using a GST–c-Jun fusion protein as the substrate. (B) Lysates from unstimulated endothelial cells or endothelial cells stimulated with LPS (100 ng/ml) were subjected to immunoblot analysis with antibodies against phosphorylated JNK (pJNK) and total JNK. Results were quantified using densitometry and are expressed in the bar graph as the change in stimulation (of both p46 and p54 JNK isoforms) above the basal level. Data represent the means plus standard errors of the means (SEMs) (error bars) for three independent experiments. Values that were significantly different (P  0.05) are indicated (*). (C) Lysates from the blots in panel B were immunoblotted with antibodies against phosphorylated Akt (pAkt) (Ser 473) and total Akt. (D) Lysates from unstimulated endothelial cells or endothelial cells stimulated with TNF (10 ng/ml) were immunoblotted for phosphorylated JNK (pJNK) and total JNK. JNK (p46 and p54) activation was quantified by densitometric scanning and is presented in the bar graph as the ratio of phosphorylated to unphosphorylated JNK. Data represent the means plus SEMs for six independent experiments. (E) Lysates from HMEC-MIY and HMEC-FADD were immunoblotted (IB) with anti-FADD antibody to monitor the expression of FADD. Molecular mass (in kilodaltons) is indicated. to the left of the blot. (F) HMEC cells were pretreated for 20 min with vehicle (dimethyl sulfoxide [DMSO]) or 50 M Z-IETD-fmk and then stimulated with LPS for the indicated times (in minutes). Activation of JNK was monitored by immunoblotting for phospho-JNK (pJNK) and total JNK. 7396 ZHANDE ET AL. MOL. CELL. BIOL.  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  (data not shown). We also observed inhibition of LPS-medi- ated activation of NF-B by FADD, consistent with a previous report (data not shown) (3). FADD and caspase 8/10 are essential for the induction of apoptosis through death recep- tors. To determine whether caspase 8/10 were also involved in suppression of LPS-mediated JNK activation, we used a caspase 8/10-specific inhibitor, Z-IETD-fmk. Inhibition of caspase 8/10 by Z-IETD-fmk (50 M) did not affect activation of JNK by LPS (Fig. 1F), although Z-IETD-fmk was able to inhibit TNF-induced apoptosis in the same cells (data not shown). Together these results indicate that the site at which FADD impinges on the LPS pathway lies upstream of MAPK, NF-B, and Akt activation and is independent of caspase 8/10 activation. The death domain of FADD-DD is sufficient and necessary to inhibit LPS signaling. The FADD death domain (FADD- DD) has previously been reported to inhibit LPS-induced apoptosis (8). Moreover, JNK has been shown to be involved in endothelial cell apoptosis in response to LPS (17). There- fore, we examined the effect of FADD-DD on LPS-induced activation of JNK in HMEC. HMEC were stably transduced with AU1-tagged FADD-DD or the empty vector, and the expression of FADD-DD was verified by immunoblotting (Fig. 2D). Similar to full-length FADD, expression of FADD-DD was sufficient to inhibit LPS stimulation of JNK activity (Fig. 2A). The expression of FADD-DD also impaired the phosphor- ylation of JNK by its activators as determined by immunoblot- ting with a JNK phospho-specific antibody (Fig. 2B). As with FADD, FADD-DD also inhibited Akt (Fig. 2B) and ERK1/2 activation (data not shown). To determine whether the DD of FADD-DD was necessary for the inhibitory function of FADD, we introduced a point mutation in FADD to replace the codon for valine 121 (V121) with that for asparagine (N) (FADDmt), which disrupts the interaction of FADD with Fas (7). As shown in Fig. 2C, FADDmt was not able to inhibit LPS-induced activation of JNK and Akt. Thus, the integrity of the DD of FADD is neces- sary for its modulation of LPS signals. Figure 2D shows expres- sion of Flag-tagged FADDmt as determined by immunoblotting. FADD-deficient cells show hyperactivation of MAPK and Akt by LPS, but not TNF. To further explore the biological relevance of FADD inhibition of LPS signaling, we examined LPS-stimulated JNK activity in murine embryonic fibroblasts from gene-targeted mice lacking FADD. We speculated that if indeed FADD serves as a negative regulator of LPS-induced cell signaling, then the absence of FADD should lead to hy- peractivation of the specific kinases under the negative regu- lation of FADD. Consistent with this hypothesis, LPS induced greater JNK activity in FADD-null MEF than in control MEF expressing wild-type FADD (Fig. 3A and B). This effect was not caused by a variation in the level of JNK protein expression (Fig. 3B). Additionally, lack of FADD markedly enhanced the activation of Akt in response to LPS (Fig. 3C). As expected, JNK activation by TNF was unaffected in FADD-deficient cells (Fig. 3D). FADD acts upstream of TRAF6. TRAF6 is a key interme- diary molecule in the LPS signaling pathway that serves as a convergence point for coordinate activation of NF-B, JNK, and Akt (14, 33). As mentioned above, TRAF6 is ubiquitinated in a K63-linked manner upon stimulation which is required for FIG. 2. The death domain of FADD is sufficient and necessary to inhibit LPS signaling. Endothelial cells expressing the death domain of FADD (HMEC-FADD-DD) or the empty vector (HMEC-LNCX) were either left untreated or treated with LPS (100 ng/ml) for the indicated times (in minutes). (A and B) Cell lysates were subjected to JNK in vitro protein kinase assay using GST–c-Jun as the substrate (A) or immunoblot analysis with antibodies against phosphorylated and total JNK and Akt (B). pJNK, phosphorylated JNK; pAkt, phos- phorylated Akt. (C) Endothelial cells expressing the empty vector (MIY) or mutant FADD (FADDmt) with the DD point mutation (V121N) were left untreated or exposed to LPS (100 ng/ml). Cell lysates were immunoblotted for phosphorylated and total JNK and Akt. (D). Lysates were immunoblotted (IB) with anti-AU1 or anti-Flag antibodies to monitor the expression of FADD-DD and FADDmt, respectively. Molecular mass (in kilodaltons) is shown to the left of the blots. VOL. 27, 2007 FADD NEGATIVELY REGULATES LPS SIGNALING 7397  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  activation (10). We thus investigated the role of FADD on TRAF6 ubiquitination. We treated HMEC-MIY and HMEC- FADD with LPS for various times. TRAF6 was then immuno- precipitated followed by immunoblotting with a ubiquitin-spe- cific antibody. Within 5 min of LPS treatment, a ladder of polyubiquitinated TRAF6 appeared in vector control cells (HMEC-MIY). This ladder was still evident after 30 min of LPS treatment (Fig. 4A). In contrast, HMEC-FADD exhibited transient and weaker TRAF6 ubiquitination that reached a peak at 20 min and had decayed by 30 min of LPS treatment (Fig. 4A). This result was not due to a variation in the protein level of TRAF6, since the amounts of immunoprecipitated TRAF6 were comparable in the different samples. Together, these findings indicate that FADD negatively regulates LPS signaling upstream of TRAF6 ubiquitination. FADD interacts with IRAK1 and MyD88. Given that the action of FADD lies upstream of TRAF6, a mechanistic model involving direct interactions between FADD, MyD88, and IRAKs can be postulated, since all of these proteins contain a DD. We used coimmunoprecipitation assays to probe for these associations. We first tested whether FADD interacts with MyD88. 293T cells were transiently transfected with Flag- tagged FADD and Myc-tagged MyD88 and immunoprecipi- tated with an anti-Myc antibody. Coimmunoprecipitated Flag- tagged FADD was detected by immunoblotting with an anti-Flag antibody. Under these conditions, FADD interacted with MyD88, confirming the results of a previous study (2) (Fig. 4B). We then addressed whether FADD could also in- teract with IRAK1. 293T cells were transiently transfected with Flag-tagged FADD and Flag-tagged IRAK1. Cell lysates were immunoprecipitated with an antibody specific for IRAK1, and coimmunoprecipitated FADD was detected with an anti- FADD antibody. As shown in Fig. 4C, FADD associated with both the transfected Flag-tagged IRAK1 and endogenous IRAK1. In reverse immunoprecipitations with an antibody against FADD, transfected FADD associated with transfected IRAK1 (Fig. 4D). Furthermore, transfected FADD also inter- acted with endogenous IRAK1 (Fig. 4D). Control immunopre- cipitations with rabbit polyclonal IgG did not immunoprecipi- tate FADD or IRAK1. We also examined the interaction of FADD with another member of the IRAK family, IRAK4, which also participates in LPS signaling. In contrast to IRAK1, there was no association between FADD and IRAK4 (data not shown). To confirm that the DD of FADD was the crucial motif in the IRAK1-FADD interaction, we transfected 293T cells with the empty vector, various epitope-tagged FADD constructs or IRAK1, and immunoprecipitated IRAK1. Immunoprecipita- tion of endogenous IRAK1 pulled down endogenous FADD, as well as overexpressed FADD and overexpressed FADD- DD, but not FADD-death effector domain (DED) (Fig. 4E). IRAK1 immunoprecipitation also pulled down overexpressed IRAK1 (Fig. 4E); however, control immunoprecipitations with rabbit polyclonal IgG did not immunoprecipitate IRAK1 or FADD. Thus, the FADD-IRAK1 interaction requires the DD, but not the DED motif, of FADD. FADD interferes with IRAK1-MyD88 interaction. We dem- onstrated above that FADD interacts with IRAK1 and MyD88. Next we wanted to investigate the consequences of these associ- ations with respect to LPS signaling. We performed time course experiments to examine IRAK1-MyD88 and IRAK1-TRAF6 in- teractions in the presence of endogenous or overexpressed FADD. HMEC transduced with the empty vector (MIY) or full- length FADD were either left untreated or stimulated with LPS for different times, and cell lysates were immunoprecipitated with the anti-IRAK1 antibody followed by immunoblot analyses with FIG. 3. FADD-deficient cells show hyperactivation of JNK and Akt in response to LPS but not TNF. Wild-type (FADD-WT) and FADD- deficient (FADD-KO [KO stands for knockout]) MEF were treated with LPS (100 ng/ml) for the times indicated. Cell lysates were ana- lyzed by a JNK in vitro protein kinase assay using GST–c-Jun as the substrate (A) or by immunoblotting for phospho- and total JNK (B) and phospho- and total Akt (C). pJNK, phosphorylated JNK; pAkt, phosphorylated Akt. The bar graph in panel A shows the change in phosphorylation of c-Jun relative to untreated samples. Data rep- resent the means plus standard errors of the means (error bars) for three independent experiments. Values that were significantly different (P  0.05) are indicated (*). (D) Lysates from MEF stimulated with TNF (10 ng/ml) were immunoblotted for phospho- and total JNK. Data are representative of three independent experiments. 7398 ZHANDE ET AL. MOL. CELL. BIOL.  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  antibodies against MyD88, TRAF6, FADD, and IRAK1. As ex- pected, endogenous IRAK1 interacted with MyD88 immediately after LPS stimulation (Fig. 5A). The association was evident by 1 min and was sustained for up to 10 min of LPS stimulation in vector control cells. The kinetics of IRAK1-MyD88 interaction was similar in both control and FADD-overexpressing cells; how- ever, the association was considerably weaker and transient in the FADD-overexpressing cells (Fig. 5A). As expected, endogenous IRAK1 also associated with TRAF6. In the absence of stimulation, TRAF6 interacted weakly with IRAK1 in control cells (Fig. 5A). However, after stimulation, there was an increase in the amount of TRAF6 that associated with IRAK1 as expected. In contrast, FADD- overexpressing cells exhibited stronger IRAK1-TRAF6 associ- ation prior to LPS stimulation with no evident changes in IRAK1-TRAF6 association following LPS stimulation (Fig. 5A). There was little association of endogenous FADD with IRAK1 in unstimulated cells. However, this association in- creased significantly upon LPS stimulation (Fig. 5A). Overex- pressed FADD clearly interacted with IRAK1 before stimula- tion, and this association increased further with stimulation. In reciprocal immunoprecipitations, FADD interacted with both MyD88 and IRAK1 (Fig. 5B). The interaction between FADD and MyD88 peaked at 5 min following LPS stimulation in HMEC-MIY. In contrast, in HMEC-FADD, FADD inter- acted with MyD88 prior to LPS stimulation, and LPS induced a further increase in the FADD-MyD88 interaction. To confirm and extend our findings, we examined the effects of FADD deficiency on IRAK1-MyD88 and IRAK1-TRAF6 associations. MEF expressing wild-type FADD or MEF defi- FIG. 4. FADD acts upstream of TRAF6 and associates with MyD88 and IRAK1. (A) Endothelial cells transduced with the empty vector (HMEC-MIY) or FADD (HMEC-FADD) were either left untreated or stimulated with LPS (100 ng/ml) for the indicated times (in minutes). Whole-cell lysates were prepared and immunoprecipitated with anti-TRAF6 antibody under denaturing conditions followed by immunoblotting with antiubiquitin and anti-TRAF6 antibodies. Lanes A are a negative control which contained the immunoprecipitating antibody with lysis buffer. Lanes L are a positive control and contain 30 g of unstimulated protein lysate. Molecular mass (in kilodaltons) is shown to the left of the blots. IP, immunoprecipitation; WB, Western blotting or immunoblotting; (Ub)n TRAF6, polyubiquitinated TRAF6. Data are representative of three independent experiments. (B) (Top) Coimmunoprecipitation analysis of lysates of 293T cells cotransfected with the indicated combinations of epitope-tagged expression plasmids (Myc-tagged MyD88 [Myc-MyD88] and Flag-tagged FADD [Flag-FADD]) and immunoprecipitated (IP) with an anti-Myc antibody. (Bottom) Immunoblot (IB) of whole-cell lysates, using anti-Flag or anti-Myc antibodies. NS, nonspecific band recognized by anti-Flag antibody. (C) (Top) Coimmunoprecipitation analysis of lysates of 293T cells cotransfected with the indicated combinations of epitope-tagged expression plasmids (Flag-tagged IRAK1 [Flag-IRAK1] and Flag-FADD) and immunoprecipitated with an anti-IRAK1 antibody. (Bottom) Immunoblot of whole-cell lysates using anti-Flag antibody. NS, nonspecific band recognized by anti-Flag antibody. (D) (Top) Coim- munoprecipitation analysis of lysates of 293T cells cotransfected with the indicated combinations of epitope-tagged expression plasmids (Flag- IRAK1 and AU1-tagged FADD [AU1-FADD]) and immunoprecipitated with an anti-FADD antibody or control IgG. (Bottom) Whole-cell lysates were immunoblotted with anti-FADD or anti-IRAK1 antibodies. (E) (Top) Coimmunoprecipitation analysis of lysates of 293T cells transfected with empty vector (pcDNA3) or the indicated epitope-tagged expression plasmid (AU1-FADD, Flag-FADD-DED, AU1-FADD-DD, or Flag- IRAK1) and immunoprecipitated with an anti-IRAK1 antibody or control IgG. (Bottom) Whole-cell lysates were immunoblotted with anti-FADD or anti-Flag antibodies. Molecular mass (in kilodaltons) is shown to the left of the blots. VOL. 27, 2007 FADD NEGATIVELY REGULATES LPS SIGNALING 7399  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  cient in FADD were stimulated with LPS or left untreated. Protein lysates were immunoprecipitated with an antibody against IRAK1 followed by immunoblotting with antibodies against MyD88, TRAF6, FADD, and IRAK1. MyD88 inter- acted with IRAK1 immediately after stimulation. In wild-type MEF, this IRAK1-MyD88 association peaked at 1 min and became weaker thereafter (Fig. 5C). In contrast, FADD-defi- cient MEF exhibited a robust IRAK1-MyD88 association which lasted up to 10 min, consistent with a role for FADD in inhibiting IRAK1-MyD88 interactions. Formation of IRAK1- TRAF6 complexes was also dependent on LPS in both cells expressing wild-type FADD and cells deficient in FADD, but TRAF6 ubiquitination was stronger in the FADD-deficient cells. Taken together, the temporal course of the IRAK1- FADD-MyD88 interactions above suggests that LPS stimula- tion recruits an IRAK1-FADD complex to MyD88, which re- duces the stability of the IRAK1-MyD88 interaction and attenuates TRAF6 activation. IRAK1 is required for FADD-MyD88 interaction. To define further the sequence of events during IRAK1-FADD-MyD88 interactions following LPS stimulation, we performed coimmu- noprecipitations using cells deficient in either MyD88 or IRAK1. We first tested whether MyD88 was required for FADD-IRAK1 interaction by examining the association of FADD and IRAK1 in MyD88-deficient cells. As shown in Fig. 6A, FADD interacted with IRAK1 in MyD88-deficient cells, which implies that the FADD-IRAK1 interaction does not depend on the presence of MyD88. In contrast, the FADD- MyD88 interaction was impaired in cells lacking IRAK1 (Fig. 6B), suggesting that IRAK1 plays a critical role in shuttling FADD to MyD88 to activate the LPS signaling cascade in a controlled and limited fashion. FADD attenuates signaling only through MyD88/IRAK1- dependent receptors. Our model would predict that FADD modulates pathways which depend on IRAK1 and MyD88 for FIG. 5. FADD interferes with IRAK1-MyD88 interaction. (A) En- dothelial cells expressing the empty vector (HMEC-MIY) or FADD (HMEC-FADD) were either unstimulated or stimulated with LPS (100 ng/ml) for the indicated times (in minutes). Cell lysates were immunoprecipitated (IP) with anti-IRAK1 antibody and then immu- noblotted with antibodies against MyD88, TRAF6, FADD, and IRAK1. (B) Lysates from endothelial cells were prepared as described above for panel A and subjected to immunoprecipitation (IP) with anti-FADD antibody, followed by immunoblotting for IRAK1, MyD88, and FADD. (C) MEF expressing wild-type FADD (MEF- FADD-WT) or MEF deficient in FADD (MEF-FADD-KO) were stimulated with LPS (1 g/ml) for the indicated times (in minutes). Cell lysates were subjected to anti-IRAK1 immunoprecipitation, fol- lowed by immunoblotting for MyD88, TRAF6, ubiquitin, FADD, and IRAK1. Molecular mass (in kilodaltons) is shown to the left of the blots. Ubn-TRAF6, polyubiquitinated TRAF6. FIG. 6. IRAK1 is required for FADD-MyD88 interaction. (A) MEF expressing wild-type MyD88 (MEF-MydD88-WT) or MEF lacking MyD88 (MEF-MydD88-KO) were stimulated with LPS (1 g/ ml) as indicated, and cell lysates were immunoprecipitated with anti- FADD antibody. Coimmunoprecipitating IRAK1 and MyD88 were detected by immunoblotting with anti-IRAK1 and anti-MyD88 anti- bodies, respectively. (B) Lysates from IRAK1-deficient MEF (MEF- IRAK-1-KO) or MEF expressing wild-type IRAK1 (MEF-IRAK-1- WT) controls were prepared and subjected to immunoprecipitation with anti-FADD antibody. Coimmunoprecipitating MyD88 and IRAK1 were detected using anti-MyD88 and anti-IRAK1 antibodies, respectively. IP, immunoprecipitation, NS, nonspecific band recog- nized by antibody. 7400 ZHANDE ET AL. MOL. CELL. BIOL.  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  signaling. We thus examined the ability of FADD to modulate signaling by other MyD88/IRAK1-dependent receptors, such as TLR2 and IL-1R, and the MyD88/IRAK1-independent re- ceptor, TLR3. Activation of JNK and Akt by Pam3CSK4, a TLR2/TLR1 ligand, was enhanced in FADD-deficient cells compared to the wild-type cells (Fig. 7A). Similarly, IL-1 stimulation led to the hyperactivation of JNK and Akt in FADD-deficient cells consistent with activation of a MyD88/ IRAK1-dependent pathway (Fig. 7B). Activation of JNK or Akt by poly(I)  poly(C), a TLR3 ligand, was weak in HMEC and MEF. In contrast to TLR2, TLR4, and IL-1R, this weak TLR3-mediated JNK and Akt activation was not enhanced in FADD-deficient cells (Fig. 7C). Taken together, these data confirm a role for FADD as a negative regulator of MyD88/ IRAK1-dependent signaling. FADD inhibits LPS-induced endothelial sprouting. We have previously shown that LPS induces endothelial sprouting in vitro and angiogenesis in vivo by directly activating endothelial cells (31). Activation of both JNK and NF-B are essential for LPS-initiated endothelial sprouting (31). Given that enforced expression of FADD in endothelial cells inhibits both JNK and NF-B activation, we expected that a functional outcome of FADD overexpression would be to block endothelial sprout- ing. We and others have used a microcarrier-based endothelial tube-forming assay that has shown remarkable correlation be- tween in vitro and in vivo findings (23, 25, 29, 31). We thus tested whether FADD overexpression would block endothelial sprouting. As seen in Fig. 8A and B, endothelial cell sprouting in response to LPS, but not serum or VEGF, was completely blocked in the presence of FADD. We also performed RNA interference using lentivirus-me- diated transfer of two short hairpin RNAs targeting distinct FADD sequences into HMEC and performed endothelial sprouting assays. Despite not being able to completely knock out FADD, we found that targeted knockdown of FADD by RNA interference slightly increased endothelial sprouting (data not shown). The increased sprouting was best seen when both short hairpin RNA constructs were used to target FADD in the same population (data not shown). As the effect of targeting FADD was only slight, these findings suggest that even small amounts of FADD may be sufficient to attenuate LPS signaling. Thus, FADD likely plays a physiological role in modulating angiogenesis in response to TLR4 signals. FADD differentially modulates cytokine induction in re- sponse to activation of TLR4, TLR2, and TLR3. We next analyzed IL-6 and IP-10 responses of MEF expressing wild- type FADD and FADD-deficient MEF following LPS, poly(I)  poly(C), and Pam3CSK4 stimulation. LPS induced an 18-fold increase of IL-6 in the medium of wild-type MEF over that of untreated MEF cultures. As would be predicted, FADD-deficient MEF displayed a robust 48-fold induction of IL-6 production in response to LPS (2.7-fold induction over wild-type MEF) (Fig. 8C). Similarly, Pam3CSK4 stimulation led to enhanced secretion of IL-6 in FADD-deficient cells compared with wild-type MEF (twofold induction over wild- type MEF) (Fig. 8C). In contrast, poly(I)  poly(C) led to an eightfold stimulation of IL-6 production in wild-type MEF, whereas FADD-deficient MEF showed almost no responsive- ness to the same stimulation (Fig. 8C). This finding confirms previous data showing a requirement for FADD in TLR3- mediated signaling (18). To confirm that the enhanced IL-6 production in LPS- and Pam3CSK4-stimulated FADD-defi- cient MEF was due solely to the absence of FADD, FADD was transduced into FADD-deficient MEF, and IL-6 concentration was measured in the medium. Reconstitution of FADD in FADD-deficient MEF reversed the enhanced IL-6 production following stimulation by LPS and Pam3CSK4 (Fig. 8D). Poly(I)  poly(C) activates the IRAK1/MyD88-independent TLR3 pathway using alternative adaptor proteins, such as TRIF. LPS is able to activate both IRAK1/MyD88-dependent and -independent pathways. Our findings thus far would pre- dict that the IRAK1/MyD88-independent pathway should not be enhanced in FADD-deficient cells. To confirm that the IRAK1/MyD88-independent, TRIF/IRF3-dependent pathway was not enhanced in FADD-deficient MEF, IP-10 production FIG. 7. FADD attenuates signaling only through IRAK1/MyD88- dependent receptors. MEF expressing wild-type FADD (FADD-WT) or deficient in FADD (FADD-KO) were stimulated with Pam3CSK4 (500 ng/ml) (A), IL-1 (50 ng/ml) (B), and poly(I)  poly(C) (Poly I:C) (100 g/ml) (C) for the indicated times (in minutes), and lysates were immunoblotted for phospho- and total JNK and Akt. Results are representative of three independent experiments. pJNK, phospho- JNK; pAkt, phospho-Akt. VOL. 27, 2007 FADD NEGATIVELY REGULATES LPS SIGNALING 7401  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  was examined (22). Poly(I)  poly(C) led to robust IP-10 pro- duction in wild-type MEF, but IP-10 production in FADD- deficient MEF was significantly impaired. Although a similar trend was observed between wild-type and FADD-deficient MEF following LPS stimulation, the reduced IP-10 response was not statistically significant (P  0.47) (Fig. 8E). Neverthe- less, the lack of enhanced IP-10 production by LPS in FADD- deficient MEF proves the differential effect of the role of FADD in IRAK1/MyD88-dependent and -independent events. Together, our data indicate that FADD negatively regulates cytokine production by attenuating the IRAK1/MyD88-de- pendent pathway, but not the IRAK1/MyD88-independent pathway. DISCUSSION TLRs function as primary sensors of microbial products. TLR activation is necessary to provide host defense against invading pathogens. However, an excessive inflammatory re- sponse to LPS, for example, can be harmful and in some cases leads to fatal septic shock. Consequently, cytokine production must be tightly controlled by mechanisms which temper the TLR response. Negative regulation of the TLR response oc- curs at many levels and can be achieved by a combination of different mechanisms, which include degradation of positive signaling components, such as IRAK1, TIRAP, and TLR4 (9, 26, 35). Recently, several diverse intracellular proteins (such as RP105, ST2, IRAK-M, MyD88s, and A20) have been identi- fied as negative regulators of TLR signaling (4–6, 11, 24). Many of these are inducible by LPS and therefore embed the pathway with a negative-feedback loop for desensitization. However, there exists a considerable lag period for induction of some of these inhibitors. The negative regulators Dok1 and Dok2 in contrast are constitutively expressed, and this allows for the control of the initial burst of the inflammatory re- sponse. However, Dok1 and Dok2 target only activation of ERKs by LPS (31a). In this study we demonstrate that FADD interacts with IRAK1 and MyD88 to inhibit MyD88-dependent (LPS, Pam3CSK4, and IL-1), but not MyD88-independent [poly(I)  poly(C) and TNF], signaling pathways. Our data support a model where LPS stimulation recruits a FADD-IRAK complex to MyD88, where- upon increasing accumulation of FADD-bound IRAK1 to MyD88 destabilizes the ternary complex, resulting in dissociation of IRAK1 from MyD88. Indeed, overexpression of FADD sig- nificantly reduces detectable MyD88-IRAK1 interactions (Fig. 5A), whereas complete loss of FADD enhances and prolongs the MyD88-IRAK1 interaction (Fig. 5C). Importantly, the IRAK1- FADD interaction is required for MyD88-FADD interaction to occur, suggesting that IRAK1 shuttles FADD to MyD88 at the receptor complex. Although IRAK4 also contains a DD, we were unable to detect an association between FADD and IRAK4 (data not shown). It is conceivable that the binding specificity is con- ferred by other determinants which are present in IRAK1 but are absent in IRAK4. Although previous studies have demonstrated the ability of FADD to inhibit LPS-induced NF-B activation (3), the mechanism of this activity has until now remained unknown. Our findings demonstrate that FADD acts at a very early point in the MyD88-dependent TLR cascade and thus serves as an imme- diate mechanism to temper signaling from the activated receptor. FIG. 8. FADD suppresses LPS-induced endothelial sprouting and IL-6 production, but not IP-10 production. Microcarrier beads seeded with endothelial cells were embedded in fibrin gels and exposed to various stimuli. (A) Micrographs of endothelial cell-coated microcarrier beads stimulated with 5% fetal bovine serum (FBS) (negative control) (i) or LPS plus 5% FBS (100 ng/ml) (ii). Arrows indicate endothelial sprouts formed after LPS stimulation. (B) Quantitation of the number of sprouts formed following stimulation by 5% FBS, VEGF (15 ng/ml), or LPS (100 ng/ml) using HMEC overexpressing FADD (MIY-FADD) or vector control (MIY). Each condition was evaluated in triplicate, and the data are the means plus standard errors of the means (error bars) from three inde- pendent experiments. Values that were significantly different (P  0.001) from the values for vector-transduced cells stimulated with LPS are indi- cated (*). (C) MEF expressing wild-type FADD (FADD-WT) and FADD-deficient (KO) MEF were treated for 6 h with medium alone (control) or stimulated with LPS (100 ng/ml), poly(I)  poly(C) (PIC) (100 g/ml), or Pam3CSK4 (500 ng/ml), and cell culture supernatants were analyzed for IL-6 by an enzyme-linked immunosorbent assay (ELISA). Data are means plus standard errors of the means (SEMs) (error bars) from four independent experiments. Values that were significantly differ- ent (P  0.05) from the values for MEF expressing FADD-WT stimu- lated with LPS, poly(I)  poly(C), or Pam3CSK4 are indicated (*). (D) FADD-KO MEF reconstituted with the empty vector (MIY) or full-length FADD (MIY-FADD) were treated for 6 h with LPS or Pam3CSK4. Cell culture supernatants were analyzed for IL-6. Data are means plus SEMs (error bars) from five independent experiments. Values that were significantly different (P  0.05) from the values for vector- transduced cells stimulated with LPS or Pam3CSK4 are indicated (*). (E) MEF expressing FADD-WT and FADD-KO were exposed to LPS or poly(I)  poly(C) (PIC) for 6 h. Supernatants were analyzed for IP-10 by ELISA. Results are means plus SEMs (error bars) from two independent experiments performed in triplicate. Values that were significantly differ- ent (P  0.05) are indicated (*). (F) Lysates from cells in panel D were immunoblotted (IB) with anti-FADD antibody to confirm reconstitution of FADD in FADD-KO MEF. Molecular mass (in kilodaltons) is indi- cated to the left of the blot. 7402 ZHANDE ET AL. MOL. CELL. BIOL.  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  We also show that enforced expression of FADD in endo- thelial cells suppresses LPS-induced activation of multiple sig- naling pathways. This inhibition also occurs in cells expressing only the DD of FADD, indicating that the DD alone is suffi- cient to confer this negative role. These findings are consistent with previous data showing that the FADD DD can inhibit LPS-induced apoptosis, which is mediated by JNK activation (8, 17). In contrast, a point mutation in the DD (V121N) abrogates the effects of overexpressed FADD on LPS signal- ing, indicating that the integrity of the DD is critical for the inhibitory function of FADD. In the absence of FADD, LPS leads to hyperactivation of JNK, Akt, and ERK, further sup- porting the role of FADD as a negative regulator. Interest- ingly, although we were able to knock down FADD protein to less than 20% of normal levels by RNA interference (data not shown), we were not able to detect an effect on downstream MAPK signaling, suggesting that FADD is present in vast excess with respect to its function in modulating IRAK1/ MyD88-dependent pathways. Together, these results suggest that under normal conditions, FADD provides an early phase safety check of the innate immune response by conferring a proper magnitude of cell activation, rather than a complete abrogation of MyD88-dependent TLR signaling. Alternatively, it may be necessary for innate immune cells to reduce the expression of FADD to allow for a more vigorous response against invading pathogens. Regulation of the innate immune response by FADD likely plays a significant physiological role and may in part explain the major defects in T- and B-cell development in FADD-null mice that appear to be independent of death receptor path- ways (19, 36, 37). FADD-deficient embryos also show signifi- cant heart defects, are hemorrhagic, and die on embryonic day 10.5 (36). In the context of the vasculature, LPS-induced acti- vation of the endothelium promotes angiogenesis (31). We have previously shown that LPS-induced angiogenic pathways lie downstream of TRAF6 and that JNK and NF-B activity are essential for LPS-induced endothelial sprouting (31). Our finding that FADD overexpression inhibits endothelial sprout- ing suggests that FADD likely modulates the angiogenic re- sponse in the innate immune pathway. FADD-deficient cells also displayed enhanced cytokine pro- duction following activation of MyD88-dependent (TLR4 and TLR2) but not MyD88-independent (TLR3 and TLR4) path- ways. The enhanced IL-6 production correlated with hyperac- tivation of MAPK and NF-B signaling pathways which are critical in both transcriptional and posttranscriptional control of IL-6 production. Reconstitution of FADD reversed the en- hanced IL-6 production, providing further functional confir- mation of the role of FADD in regulating MyD88-dependent pathways. Although most TLRs use the adaptor protein MyD88 lead- ing to the activation of IRAK, TLR4 and TLR3 use an alter- nate adaptor protein, TRIF. In the case of TLR4, both MyD88- and TRIF-dependent pathways are engaged, whereas TLR3 functions only through TRIF (12). This latter pathway proceeds without IRAK participation and induces expression of genes, such as IP-10 (18). TLR3 stimulation with poly(I)  poly(C) led to robust IP-10 production in wild-type cells but was impaired in FADD-deficient cells, thus indicating a positive role for FADD in TLR3-regulated IP-10 gene ex- pression. This finding was expected, since a role for FADD in TLR3-mediated responses, such as B-cell proliferation, as well as in apoptosis, has previously been reported (18, 20). In con- trast to the IL-6 response, LPS-induced IP-10 production was not affected in the absence of FADD, thereby confirming that FADD diminishes only IRAK1/MyD88-dependent signaling. Given that IP-10 can also be regulated by NF-B, which is hyperactivated—through the IRAK1/MyD88-dependent path- way—in the absence of FADD, this may explain the lack of suppression of IP-10 induction in response to LPS stimulation. Taken together, our data indicate that whereas FADD is re- quired to propagate the TRIF-dependent response, FADD acts to attenuate IRAK1/MyD88-dependent signaling. In conclusion, our findings demonstrate that FADD is a novel negative regulator of LPS signaling through its interac- tion with IRAK1 and inhibition of IRAK1-MyD88 interac- tions. Our data also suggest that IRAK1 acts as a shuttle to bring FADD in contact with activated MyD88, which serves as a switch to attenuate the LPS signal. Given the ubiquitous expression of FADD in immune cells, our findings implicate FADD in playing a critical role in fine-tuning the innate im- mune response and may explain some of the death receptor- independent functions postulated for FADD (19, 36, 37). ACKNOWLEDGMENTS This work was supported by grants to A.K. from the Canadian Institutes of Health Research and the Heart and Stroke Foundation of British Columbia and the Yukon. S.M.D. is the recipient of a student- ship award from the Canadian Institutes for Health Research and the Michael Smith Foundation for Health Research. A.K. is a Senior Scholar of the Michael Smith Foundation for Health Research. REFERENCES 1. Adachi, O., T. Kawai, K. Takeda, M. Matsumoto, H. Tsutsui, M. Sakagami, K. Nakanishi, and S. Akira. 1998. Targeted disruption of the MyD88 gene results in loss of IL-1- and IL-18-mediated function. Immunity 9:143–150. 2. Aliprantis, A. O., R. B. Yang, D. S. Weiss, P. Godowski, and A. Zychlinsky. 2000. The apoptotic signaling pathway activated by Toll-like receptor-2. EMBO J. 19:3325–3336. 3. Bannerman, D. D., J. C. Tupper, J. D. Kelly, R. K. Winn, and J. M. Harlan. 2002. The Fas-associated death domain protein suppresses activation of NF-kappa B by LPS and IL-1 beta. J. Clin. Investig. 109:419–425. 4. Boone, D. L., E. E. Turer, E. G. Lee, R. C. Ahmad, M. T. Wheeler, C. Tsui, P. Hurley, M. Chien, S. Chai, O. Hitotsumatsu, E. McNally, C. Pickart, and A. Ma. 2004. The ubiquitin-modifying enzyme A20 is required for termina- tion of Toll-like receptor responses. Nat. Immunol. 5:1052–1060. 5. Brint, E. K., D. Xu, H. Liu, A. Dunne, A. N. McKenzie, L. A. O’Neill, and F. Y. Liew. 2004. ST2 is an inhibitor of interleukin 1 receptor and Toll-like receptor 4 signaling and maintains endotoxin tolerance. Nat. Immunol. 5:373–379. 6. Burns, K., S. Janssens, B. Brissoni, N. Olivos, R. Beyaert, and J. Tschopp. 2003. Inhibition of interleukin 1 receptor/Toll-like receptor signaling through the alternatively spliced, short form of MyD88 is due to its failure to recruit IRAK-4. J. Exp. Med. 197:263–268. 7. Chinnaiyan, A. M., K. O’Rourke, M. Tewari, and V. M. Dixit. 1995. FADD, a novel death domain-containing protein, interacts with the death domain of Fas and initiates apoptosis. Cell 81:505–512. 8. Choi, K. B., F. Wong, J. M. Harlan, P. M. Chaudhary, L. Hood, and A. Karsan. 1998. Lipopolysaccharide mediates endothelial apoptosis by a FADD-dependent pathway. J. Biol. Chem. 273:20185–20188. 9. Chuang, T. H., and R. J. Ulevitch. 2004. Triad3A, an E3 ubiquitin-protein ligase regulating Toll-like receptors. Nat. Immunol. 5:495–502. 10. Deng, L., C. Wang, E. Spencer, L. Yang, A. Braun, J. You, C. Slaughter, C. Pickart, and Z. J. Chen. 2000. Activation of the IkappaB kinase complex by TRAF6 requires a dimeric ubiquitin-conjugating enzyme complex and a unique polyubiquitin chain. Cell 103:351–361. 11. Divanovic, S., A. Trompette, S. F. Atabani, R. Madan, D. T. Golenbock, A. Visintin, R. W. Finberg, A. Tarakhovsky, S. N. Vogel, Y. Belkaid, E. A. Kurt-Jones, and C. L. Karp. 2005. Negative regulation of Toll-like receptor 4 signaling by the Toll-like receptor homolog RP105. Nat. Immunol. 6:571– 578. VOL. 27, 2007 FADD NEGATIVELY REGULATES LPS SIGNALING 7403  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  12. Doyle, S., S. Vaidya, R. O’Connell, H. Dadgostar, P. Dempsey, T. Wu, G. Rao, R. Sun, M. Haberland, R. Modlin, and G. Cheng. 2002. IRF3 mediates a TLR3/TLR4-specific antiviral gene program. Immunity 17:251–263. 13. Fitzgerald, K. A., E. M. Palsson-McDermott, A. G. Bowie, C. A. Jefferies, A. S. Mansell, G. Brady, E. Brint, A. Dunne, P. Gray, M. T. Harte, D. McMurray, D. E. Smith, J. E. Sims, T. A. Bird, and L. A. O’Neill. 2001. Mal (MyD88-adapter-like) is required for Toll-like receptor-4 signal transduc- tion. Nature 413:78–83. 14. Gohda, J., T. Matsumura, and J. Inoue. 2004. TNFR-associated factor (TRAF) 6 is essential for MyD88-dependent pathway but not Toll/IL-1 receptor domain-containing adaptor-inducing IFN-beta (TRIF)-dependent pathway in TLR signaling. J. Immunol. 173:2913–2917. 15. Horng, T., G. M. Barton, and R. Medzhitov. 2001. TIRAP: an adapter molecule in the Toll signaling pathway. Nat. Immunol. 2:835–841. 16. Hsu, H., H. B. Shu, M. G. Pan, and D. V. Goeddel. 1996. TRADD-TRAF2 and TRADD-FADD interactions define two distinct TNF receptor 1 signal transduction pathways. Cell 84:299–308. 17. Hull, C., G. McLean, F. Wong, P. J. Duriez, and A. Karsan. 2002. Lipopoly- saccharide signals an endothelial apoptosis pathway through TNF receptor- associated factor 6-mediated activation of c-Jun NH2-terminal kinase. J. Im- munol. 169:2611–2618. 18. Imtiyaz, H. Z., S. Rosenberg, Y. Zhang, Z. S. Rahman, Y. J. Hou, T. Manser, and J. Zhang. 2006. The Fas-associated death domain protein is required in apoptosis and TLR-induced proliferative responses in B cells. J. Immunol. 176:6852–6861. 19. Kabra, N. H., C. Kang, L. C. Hsing, J. Zhang, and A. Winoto. 2001. T cell-specific FADD-deficient mice: FADD is required for early T cell devel- opment. Proc. Natl. Acad. Sci. USA 98:6307–6312. 20. Kaiser, W. J., and M. K. Offermann. 2005. Apoptosis induced by the Toll- like receptor adaptor TRIF is dependent on its receptor interacting protein homotypic interaction motif. J. Immunol. 174:4942–4952. 21. Kanayama, A., R. B. Seth, L. Sun, C. K. Ea, M. Hong, A. Shaito, Y. H. Chiu, L. Deng, and Z. J. Chen. 2004. TAB2 and TAB3 activate the NF-kappaB pathway through binding to polyubiquitin chains. Mol. Cell 15:535–548. 22. Kawai, T., O. Takeuchi, T. Fujita, J. Inoue, P. F. Muhlradt, S. Sato, K. Hoshino, and S. Akira. 2001. Lipopolysaccharide stimulates the MyD88- independent pathway and results in activation of IFN-regulatory factor 3 and the expression of a subset of lipopolysaccharide-inducible genes. J. Immunol. 167:5887–5894. 23. Kim, I., H. G. Kim, S.-O. Moon, S. W. Chae, J.-N. So, K. N. Koh, B. C. Ahn, and G. Y. Koh. 2000. Angiopoietin-1 induces endothelial cell sprouting through the activation of focal adhesion kinase and plasmin secretion. Circ. Res. 86:952–959. 24. Kobayashi, K., L. D. Hernandez, J. E. Galan, C. A. Janeway, Jr., R. Medzhitov, and R. A. Flavell. 2002. IRAK-M is a negative regulator of Toll-like receptor signaling. Cell 110:191–202. 25. Leong, K. G., X. Hu, L. Li, M. Noseda, B. Larrivee, C. Hull, L. Hood, F. Wong, and A. Karsan. 2002. Activated Notch4 inhibits angiogenesis: role of 1-integrin activation. Mol. Cell. Biol. 22:2830–2841. 26. Mansell, A., R. Smith, S. L. Doyle, P. Gray, J. E. Fenner, P. J. Crack, S. E. Nicholson, D. J. Hilton, L. A. O’Neill, and P. J. Hertzog. 2006. Suppressor of cytokine signaling 1 negatively regulates Toll-like receptor signaling by me- diating Mal degradation. Nat. Immunol. 7:148–155. 27. Medzhitov, R., P. Preston-Hurlburt, E. Kopp, A. Stadlen, C. Chen, S. Ghosh, and C. A. Janeway, Jr. 1998. MyD88 is an adaptor protein in the hToll/IL-1 receptor family signaling pathways. Mol. Cell 2:253–258. 28. Nagata, S. 1999. FAS ligand-induced apoptosis. Annu. Rev. Genet. 33: 29–55. 29. Nakatsu, M. N., R. C. A. Sainson, J. N. Aoto, K. L. Taylor, M. Aitkenhead, S. Perez-del-Pulgar, P. M. Carpenter, and C. C. W. Hughes. 2003. Angio- genic sprouting and capillary lumen formation modeled by human umbilical vein endothelial cells (HUVEC) in fibrin gels: the role of fibroblasts and Angiopoietin-1. Microvasc. Res. 66:102–112. 30. Oshiumi, H., M. Sasai, K. Shida, T. Fujita, M. Matsumoto, and T. Seya. 2003. TIR-containing adapter molecule (TICAM)-2, a bridging adapter re- cruiting to Toll-like receptor 4 TICAM-1 that induces interferon-beta. J. Biol. Chem. 278:49751–49762. 31. Pollet, I., C. J. Opina, C. Zimmerman, K. G. Leong, F. Wong, and A. Karsan. 2003. Bacterial lipopolysaccharide directly induces angiogenesis through TRAF6-mediated activation of NF-kappaB and c-Jun N-terminal kinase. Blood 102:1740–1742. 31a.Shinohara, H., A. Inoue, N. Toyama-Sorimachi, Y. Nagai, T. Yasuda, H. Suzuki, R. Horai, Y. Iwakura, T. Yamamoto, H. Karasuyama, K. Miyake, and Y. Yamanishi. 2005. Dok-1 and Dok-2 are negative regulators of lipo- polysaccharide-induced signaling. J. Exp. Med. 7:333–339. 32. Thomas, J. A., J. L. Allen, M. Tsen, T. Dubnicoff, J. Danao, X. C. Liao, Z. Cao, and S. A. Wasserman. 1999. Impaired cytokine signaling in mice lacking the IL-1 receptor-associated kinase. J. Immunol. 163:978–984. 33. Wong, F., C. Hull, R. Zhande, J. Law, and A. Karsan. 2004. Lipopolysac- charide initiates a TRAF6-mediated endothelial survival signal. Blood 103: 4520–4526. 34. Yamamoto, M., S. Sato, H. Hemmi, S. Uematsu, K. Hoshino, T. Kaisho, O. Takeuchi, K. Takeda, and S. Akira. 2003. TRAM is specifically involved in the Toll-like receptor 4-mediated MyD88-independent signaling pathway. Nat. Immunol. 4:1144–1150. 35. Yamin, T. T., and D. K. Miller. 1997. The interleukin-1 receptor-associated kinase is degraded by proteasomes following its phosphorylation. J. Biol. Chem. 272:21540–21547. 36. Yeh, W. C., J. L. Pompa, M. E. McCurrach, H. B. Shu, A. J. Elia, A. Shahinian, M. Ng, A. Wakeham, W. Khoo, K. Mitchell, W. S. El-Deiry, S. W. Lowe, D. V. Goeddel, and T. W. Mak. 1998. FADD: essential for embryo development and signaling from some, but not all, inducers of apoptosis. Science 279:1954–1958. 37. Zhang, J., D. Cado, A. Chen, N. H. Kabra, and A. Winoto. 1998. Fas- mediated apoptosis and activation-induced T-cell proliferation are defective in mice lacking FADD/Mort1. Nature 392:296–300. 7404 ZHANDE ET AL. MOL. CELL. BIOL.  at UNIV O F BRITISH CO LUM BIA on Decem ber 11, 2009 m cb.asm .org D ow nloaded from  169              APPENDIX II                               170    APPENDIX II          The following are the animal care approval certificates required during this thesis.  ANIMAL CARE CERTIFICATE  The Animal Care Committee has examined and approved the use of animals for the above experimental project. This certificate is valid for one year from the above start or approval date (whichever is later) provided there is no change in the experimental procedures.  Annual review is required by the CCAC and some granting agencies.    THE UNIVERSITY OF BRITISH COLUMBIA Application Number: A06-0038 Investigator or Course Director: Aly Karsan Department: Pathology & Laboratory Medicine Funding Sources:   Animals:   Mice SASH1-flox/flox x VE-Cre 60 Mice SASH1+/- 43 Mice SASH1-flox/+ x VE-Cre 36 Mice Rosa x VE-Cre 12 Mice Rosa x Tek-Cre 4 Mice SASH1+/- 48 Mice Myosin10+/- 4 Mice Myosin10+/- 2 Mice SASH1+/- 25 Mice C57Bl/6J 6 Start Date:  March 15, 2006 Approval Date: May 11, 2009 Funding Agency:  Canadian Institutes of Health Research (CIHR) Funding Title:  Lipopolysaccharide signaling in the vasculature  Funding Agency:  Canadian Institutes of Health Research (CIHR) Funding Title:  Lipopolysaccharide signaling in endothelial cells  Unfunded title:  N/A Page 1 of 2 11/9/2009https://rise.ubc.ca/rise/Doc/0/JURLO5NL7ERKPB9I1DQC1B9Q7E/fromString.html A copy of this certificate must be displayed in your animal facility.  Office of Research Services and Administration 102, 6190 Agronomy Road, Vancouver, BC V6T 1Z3 Phone: 604-827-5111 Fax: 604-822-5093 Page 2 of 2 11/9/2009https://rise.ubc.ca/rise/Doc/0/JURLO5NL7ERKPB9I1DQC1B9Q7E/fromString.html   THE UNIVERSITY OF BRITISH COLUMBIA  ANIMAL CARE CERTIFICATE BREEDING PROGRAMS  Application Number: A09-0275 Investigator or Course Director: Aly Karsan Department: Pathology & Laboratory Medicine Animals:  Mice VE-Cre 92 Mice VE-Cre x ROSA 66 Mice SASH-1+/- 305 Mice ROSA 96 Mice TekCre x ROSA 64 Mice TekCre 70 Mice SASH-1-flox/flox x VECre 132 Mice SASH-1-flox/flox 150 Mice Myosin10+/- 150  Approval Date: September 27, 2009 Funding Sources:  Funding Agency:   Canadian Institutes of Health Research (CIHR) Funding Title:   Lipopolysaccharide signaling in endothelial cells   Unfunded title:    N/A  The Animal Care Committee has examined and approved the use of animals for the above breeding program. This certificate is valid for one year from the above approval date provided there is no change in the experimental procedures.  Annual review is required by the CCAC and some granting agencies.  A copy of this certificate must be displayed in your animal facility.  Office of Research Services and Administration 102, 6190 Agronomy Road, Vancouver, BC V6T 1Z3 Phone: 604-827-5111 Fax: 604-822-5093  

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
https://iiif.library.ubc.ca/presentation/dsp.24.1-0069549/manifest

Comment

Related Items