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Participation of dendritic cells in neuroinflammation : factors regulating adhesion to human cerebral… Arjmandi Rafsanjani, Azadeh 2008

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PARTICIPATION OF DENDRITIC CELLS IN NEUROINFLAMMATION: FACTORS REGULATING ADHESION TO HUMAN CEREBRAL ENDOTHELIUM  by  AZADEH ARJMANDI RAFSANJANI BSc. (Psychology), The University of British Columbia, 2004  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES (Experimental Pathology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  August 2008  © Azadeh Arjmandi Rafsanjani, 2008  ABSTRACT  Dendritic cells (DCs) form a key component of the immune response, as they are involved in the innate and adaptive immunity and in the process of tolerance. Under normal conditions, DCs are absent from the Central Nervous System (CNS), as the blood brain barrier (BBB) restricts their entry.  However, DCs have recently been implicated in the pathogenesis of several CNS  diseases. The molecular mechanisms that mediate DC trafficking across the BBB are poorly understood. The objectives of this study were to examine the role of endothelial cell adhesion molecules (eCAMs) and their ligands in the process of DC adhesion to the BBB endothelium, and to investigate the participation of DCs in human CNS diseases. To study DC adhesion, DCs were generated in vitro by culturing human blood monocytes in the presence of GM-CSF and IL4, and DC maturation was induced by adding inflammatory cytokines (TNF-cL, IL-1f3, IL-6) and . Immature and mature DCs displayed differences in their expression of surface molecules, 2 PGE including eCAM ligands, by flow cytometry.  Adhesion to the cerebral endothelium was  investigated using an in vitro model of the BBB consisting of primary cultures of human brain microvessel endothelial cells (HBMEC). Immature or mature DCs were incubated with resting or TNF-ct-activated HBMEC for up to one hour. Only a few DCs adhered to resting HBMEC, but adhesion was upregulated upon activating HBMEC (p<O.Ol). Moreover, immature DCs adhered to activated HBMEC to a greater extent compared to mature DCs (p<O.OOl). Blocking experiments indicated that the adhesion of both immature and mature DCs to HBMEC was dependent upon ICAM-1-CD18 or ICAM-2-CD18, ICAM-2-DC-SIGN, and PECAM-l PECAM-l interactions. In addition, VCAM-1-VLA-4 interactions mediated the adhesion of immature but not mature DCs to activated HBMEC.  Using immunohistochemistry for DC  markers, we also examined the presence of DCs in human inflammatory, infectious, and neurodegenerative diseases, stroke and tumours.  The results indicate accumulation of DC  SIGN—, fascin—, and MHC class Il—expressing DCs in the CNS under most pathological conditions. These findings provide further insight into the mechanisms of neuroinflammation, and highlight the role of DCs and the BBB endothelium in this process.  11  TABLE OF CONTENTS  Abstract  ii  Table of Contents  iii  List of Tables  vii  List of Figures  viii  List of Abbreviations  x  Acknowledgements  xiii  CHAPTER!: INTRODUCTION 1.1.  1.2.  1  The Inflammatory Response  1  1.1.1. The Process of Leukocyte Trafficking to Tissues  2  1.1.2. Endothelial Cell Adhesion Molecules and Their Ligands  3  1.1.2.1.  Selectins  4  1.1.2.2.  Integrins  4  1.1.2.3.  Immunoglobulins  5  1.1.2.4.  DC-SIGN: A Novel Cell Adhesion Molecule  7  The Central Nervous System (CNS)  8  1.2.1. Inflammation and CNS Pathology  8  1.2.2. The Blood Brain Barrier: Structural and Functional Properties  9  1.2.3. Cells Participating in CNS Inflammation  11  1.2.3.1.  Astrocytes  11  1.2.3.2.  Microglia  12  1.2.3.3.  Monocytes and Other Myeloid Cells  12  1.2.3.4.  Lymphocytes  14  1.2.4. Inflammatory Cytokines in the CNS 1.2.4.1.  TumourNecrosis Factor (TNF)-c  1.2.5. CAMs in CNS Inflammation  15 15 16  111  1.3.  1.4.  1.5.  1.6.  Human Brain Microvessel Endothelial Cells (HBMEC)  17  1.3.1. Endothelial Cell Heterogeneity  17  1.3.2. Role of Endothelial Cells in CNS Inflammation  18  1.3.3. In vitro Model of the Human Blood Brain Barrier  18  Antigen Presentation in the CNS  19  1.4.1. Ag Presentation and CNS Pathology  20  1.4.2. Ag Presenting Cells of the CNS  21  1.4.2.1.  Microglia  21  1.4.2.2.  Perivascular Macrophages  21  1.4.2.3.  Endothelial Cells  22  1.4.2.4.  Dendritic Cells  22  Dendritic Cells (DCs)  23  1.5.1. Role of DCs in Immunity and Tolerance  23  1.5.2. DC Heterogeneity  24  1.5.3. In Vitro Generation of DCs from Monocytes  26  1.5.4. Cytokines Involved in DC Differentiation and Maturation  27  1.5.4.1.  GM-CSF  27  1.5.4.2.  IL-4  28  1.5.4.3.  TNF-cL  29  1.5.4.4.  IL-113  30  1.5.4.5.  IL-6  30  1.5.4.6.  2 PGE  32  1.5.5. Role of DCs in CNS Pathology  32  1.5.6. The Great Debate: Origin of CNS DCs  34  1.5.7. DC Adhesion to and Migration across Endothelial Barriers  35  Objectives and Specific Aims  36  1.6.1. Objectives and Hypotheses  36  1.6.2. Specific Aims  36  iv  CHAPTER 2: MATERIALS AND METHODS  38  2.1.  Endothelial Cell Cultures  38  2.1.1. Isolation of Human Brain Microvessel Endothelial Cells (HBMEC)  38  2.1.2. Culture Conditions  38  Dendritic Cell Generation  39  2.2.1. Monocyte Isolation  39  2.2.2. In-vitro Generation of Immature and Mature DCs  40  FACS Analysis  40  2.3.1. Cell Surface Staining  40  2.3.2. Data Collection and Analysis  41  Antibodies  41  2.4.1. Flow Cytometry Antibodies  41  2.4.2. Antibodies for Adhesion Assays and Blocking Studies  42  2.4.3. Antibodies for In-Situ Study  43  2.5.  Adhesion Assay and Immunocytochemistry  43  2.6.  Enzyme-Linked Immunosorbent Assay (ELISA)  44  2.7.  Blocking Studies and Immunoperoxidase Staining  45  2.8.  Dendritic Cells in CNS Pathology  46  2.8.1. Patients  46  2.8.2. Immunohistochemistry  46  Statistical Analysis  48  2.2.  2.3.  2.4.  2.9.  CHAPTER 3: RESULTS  50  3.1. Human Brain Microvessel Endothelial Cells (HBMEC)  50  3.2. Surface Phenotype of in vitro-generated DCs  50  3.2.1. Characterization of Immature and Mature DCs  50  3.2.2. Expression of eCAM Ligands by Immature and Mature DCs  51  3.3. Adhesion of Immature and Mature DCs to HBMEC  52  3.3.1. Adhesion to Resting and Activated HBMEC  52  3.4.2. DC Adhesion Change with Time  52  3.4. Surface Expression of ICAM-2 by HBMEC  V  53  3.5. Regulation of DC adhesion to HBMEC by eCAMs and their ligands  53  3.5.1. DC Adhesion to Resting HBMEC  53  3.5.2. DC Adhesion to Activated HBMEC  54  3.6. Dendritic Cell Participation in the CNS Immune Response  55  3.6.1. DC-SIGN-Positive DCs  57  3.6.2. Fascin-Positive Cells  58  3.6.3. CD4O-Positive Cells  58  3.6.4. MHC Class Il-Positive Cells  59  3.6.5. Immature vs. Mature DC Participation in CNS Pathology  59  CHAPTER 4: DISCUSSION  60  4.1. HBMECs as a Model of the BBB  60  4.2. Characterization of Immature and Mature DCs  61  4.3. DC Adhesion to HBMEC  64  4.4. Regulation of DC Adhesion to HBMEC by eCAMs and their Ligands  66  4.5. Participation of DCs in CNS Pathology  69  CHAPTERS: CONCLUSIONS  75  5.1. Summary and Significance  75  5.2. Future Directions  77  REFERENCES  78  vi  LIST OF TABLES  Table 1  Patient Data  49  Table 2  DC Participation in CNS Pathology  56  vii  LIST OF FIGURES  Figure 1  Overview of leukocyte trafficking across ECs  Figure 2  Summary of eCAM-ligand interactions in DC adhesion to HBMEC  Figure 3  Primary cultures of Human Brain Microvessel Endothelial Cells  8 69 103  Figure 4 (a & b) Characterization of human monocyte-derived immature and mature DCs  104  Figure 4c  Expression of eCAM ligands by immature and mature DCs  105  Figure 5a  Immature and mature DC adhesion to resting or TNF-ct-activated HBMEC (Cytokine activation time: 24 h)  Figure Sb  106  Immature and mature DC adhesion to resting or TNF-ct-activated HBMEC (Cytokine activation time: 5 h)  107  Figure Sc  Immature and Mature DC adhesion to HBMEC increases with time  108  Figure 6  Relative surface expression of ICAM-2 by resting and TNF-a.-activated HBMEC as measured by ELISA  Figure 7  DC adhesion to resting HBMEC in the presence of blocking Abs against eCAMs  Figure 8  110  DC adhesion to resting HBMEC in the presence of blocking Abs against eCAM ligands  Figure 9  111  Adhesion of immature DCs to TNF-ct-activated HBMEC in the presence of blocking Abs against eCAMs  Figure 10  114  Adhesion of mature DCs to TNF-a-activated HBMEC in the presence of blocking Abs against eCAM ligands  Figure 13  115  Adhesion of immature DCs to TNF-a-activated HBMEC in the presence of blocking Abs against eCAMs and their ligands  Figure 14  113  Adhesion of mature DCs to TNF-a-activated HBMEC in the presence of blocking Abs against eCAMs  Figure 12  112  Adhesion of immature DCs to TNF-cL-activated HBMEC in the presence of blocking Abs against eCAM ligands  Figure 11  109  116  Adhesion of mature DCs to TNF-a-activated HBMEC in the presence of blocking Abs against eCAMs and their ligands  viii  117  Figure 1 5(a-e)  DC-SIGN expression in situ in normal CNS and in CNS inflammatory diseases  118  Figure 15(f-k)  DC-SIGN expression in situ in infectious CNS pathologies  119  Figure 15(l-q)  DC-SIGN expression in situ in the CNS of patients with tuberculosis, sarcoidosis, neurodegenerative diseases, and tumours  120  Figure 16  DC-SIGN expression in situ in normal and pathological CNS  121  Figure 1 7(a-e)  Fascin expression in situ in normal CNS and in CNS inflammatory diseases  122  Figure 17(f-k)  Fascin expression in situ in infectious CNS pathologies  123  Figure 17(l-q)  Fascin expression in situ in the CNS of patients with tuberculosis, sarcoidosis, neurodegenerative diseases, and tumours  124  Figure 18  Fascin expression in situ in normal and pathological CNS  125  Figure 1 9(a-e)  CD4O expression in situ in normal CNS and in CNS inflammatory diseases  126  Figure 19(f-k)  CD4O expression in situ in infectious CNS pathologies  127  Figure 1 9(l-q)  CD4O expression in situ in the CNS of patients with tuberculosis, sarcoidosis, neurodegenerative diseases, and tumours  128  Figure 20  CD4O expression in situ in normal and pathological CNS  129  Figure 21(a-e)  MHC class II expression in situ in normal CNS and in CNS inflammatory diseases  130  Figure 21 (f-k)  MHC class II expression in situ in infectious CNS pathologies  131  Figure 21 (l-q)  CD4O expression in situ in the CNS of patients with tuberculosis, sarcoidosis, neurodegenerative diseases, and tumours  Figure 22  CD4O expression in situ in normal CNS and in CNS inflammatory diseases  Figure 23a  133  Immature DC participation in normal and pathological CNS tissue in situ  Figure 23b  132  134  Immature vs. mature DC participation in normal and pathological CNS tissue in situ  134  ix  LIST OF ABBREVIATIONS  Ab  antibody  AD  Alzheimer’s disease  AEC  3-amino, 9 ethyl-carbazole  Ag(s)  antigen(s)  ALS  amyotrophic lateral sclerosis  ANOVA  analysis of variance  APC  antigen presenting cell  BBB  blood-brain barrier  CAA  cerebral amyloid angiopathy  CAM(s)  cell adhesion molecule(s)  CNS  central nervous system  CSF  cerebrospinal fluid  CTL  cytotoxic T cell  DC(s)  dendritic cell(s)  DC-SIGN  dendritic cell-specific ICAM-grabbing non-integrin  EAE  experimental autoimmune encephalomyelitis  eCAM(s)  endothelial cell adhesion molecule(s)  EC(s)  endothelial cell(s)  ECM  extracellular matrix  EDTA  Ethylenediaminetetraacetic acid  ELISA  enzyme-linked immunosorbent assay  FACS  fluorescence-activated cell sorting  x  GBM  Glioblastoma multiforme  G1yCAM-1  glycosylation-dependent cell adhesion molecule-i  GM-CSF  granulocyte-macrophage colony stimulating factor  HAMJTSP  HTLV-1 associated myelopathy/tropical spastic paraparesis  HBMEC  human brain microvessel endothelial cell(s)  HIV- 1  human immunodeficiency virus-i  HLA  human leukocyte antigen  HRP  horseradish peroxidase  HSV  Herpes simplex virus  HUVEC  human umbilical vein endothelial cell(s)  ICAM-1  intercellular adhesion molecule-i  ICAM-2  intercellular adhesion molecule-2  IFN-y  interferon gamma  Ig  immunoglobulin  IL-i f3  interleukin- if3  IL-4  interleukin-4  IL-6  interleukin-6  JAM(s)  junctional adhesion molecule(s)  LCA  leukocyte common antigen  LFA-1  lymphocyte function-associated antigen-i  LN(s)  lymph node(s)  LPS  Iipopolysaccharide  Mac-i  macrophage- 1 antigen  xi  MFI  mean fluorescence intensity  MHC  major histocompatibility complex  MIP- 1 a  Macrophage inflammatory protein-i alpha  MS  multiple sclerosis  NK cells  natural killer cells  PBS  phosphate buffered saline  PD  Parkinson’s disease  PE  Phycoerythrin  2 PGE  prostaglandin E2  PECAM- 1  platelet/endothelial cell adhesion molecule-i  PML  Progressive Multifocal Leukoencephalopathy  PMN(s)  polymorphonuclear leukocyte(s)  PSGL-1  P-selectin glycoprotein ligand-1  SIV  simian immunodeficiency virus  sLex  sialyl-Lewis x  TB  tuberculosis  TCR  T cell receptor  TGF-13  transforming growth factor-13  TJ(s)  tight junction(s)  TNF-a  tumour necrosis factor-a  VCAM-i  vascular cell adhesion molecule-i  VLA-4  very late activation antigen-4  VZV  Varicella zoster virus  xli  ACKNOWLEDGEMENTS  My sincere thanks go to my supervisor, Dr. Katerina Zis, for her guidance, mentorship and solid support, and also for her tireless and inspiring love of science and her genuine concern for the progress of students. I would also like to thank the members of my supervisory committee, Dr. Pauline Johnson, Dr. Wayne Moore, Dr. Susan Porter, and Dr. Douglas Waterfield, for their insightful suggestions and invaluable advice. Also, many heartfelt thanks go to the members of the Neuropathology Research Laboratory for their endless help and support: Mr. Ken Liu, Ms. Rukmini Prameya, Ms. Hong Li, Dr. Jaya Taireja, Ms. Lu Yao, Dr. Elaine Humphrey, and past lab members: Dr. Kaveh Koochesfahani, Mr. Reza Shahidi and Ms. Farah Bahrami. I also thank Ms. Vivian Wu for her assistance with flow cytometry, and Ms. Esther Leung for her encouraging words. I would also like to thank the Department of Pathology and Laboratory Medicine, and in particular Dr. David Walker and Dr. Marcel Bally for having an active interest in fostering student learning. As well, my thanks go to Ms. Penny Woo for her timely management of student affairs. I would also like to extend my gratitude to the Multiple Sclerosis Society of Canada and The Michael Smith Foundation for Health Research for providing research studentships. Finally, I would like to thank my family and friends, near and far, for their love and patience. Special thanks go to my parents, for all their selfless support, and Pouyan, Sheida and Gelareh, for all the laughter and joy.  xiii  CHAPTER 1: INTRODUCTION 1.1.  THE INFLAMMATORY RESPONSE  The immune system is the body’s defense system against threats to the health and survival of the organism, be it an attack by micro-organisms (such as bacteria and viruses) or tissue damage due to trauma and injury. The immune system is divided into the two branches of “innate” and “adaptive” immunity. In each branch, the host organism utilizes immune cells (leukocytes) as well as proteins secreted by leukocytes or other cells in order to exert its effect. Innate immunity, which in vertebrates is the first line of defense against pathogens, consists of a complex cascade of events initiated by the cells and molecules at the site of infection or injury, with the goal of containing and suppressing the damage. This process is known as the acute inflammatory response. It is now believed that certain molecular structures commonly associated with pathogens (known as pathogen-associated molecular patterns, such as bacterial lipopolysaccharide (LPS) or flagellin, viral double-stranded RNA, and unmethylated CpG motifs) are recognized by the receptors on the cells of the innate immune system and initiate the acute inflammatory cascade, which is a non-specific response. The inflammatory events that follow include the secretion of cytokines and other inflammatory mediators, vasodilation, and an increase in the expression of adhesion molecules on the surface of blood vessel endothelial cells (ECs), as well as on leukocytes, which collectively lead to increase in the permeability of the local blood vessels and the extravasation of polymorphonuclear leukocytes (PMNs, also known as acute inflammatory cells) and plasma proteins into the site of injury or infection. Although in many cases the acute inflammatory process is capable of controlling pathogens, it can also lead to severe tissue damage, partially due to its non-specific nature and  1  the release of cytotoxic molecules (e.g. reactive oxygen and nitrogen species). Thus, more persistent pathogens are typically confronted and eliminated through an adaptive immune response, which entails chronic inflammation. This is a much more specific response, which depends upon the presentation of antigens (Ags) by antigen-presenting cells (such as dendritic cells and macrophages), the recognition of these antigens by cells of the adaptive immunity (B and T lymphocytes), prolonged vascular permeability, and infiltration of tissue by chronic inflammatory cells (lymphocytes, macrophages, and plasma cells). Thus, chronic inflammation responds to persistent or frequently-encountered pathogens through the activation of humoral (antibody-mediated) and/or cell (B or T lymphocyte)-mediated immunity. Furthermore, through the formation of “immunological memory”, adaptive immunity enables the host to launch a quicker and more efficient response during subsequent encounters of the same pathogen. Hence, the inflammatory response, which plays a key role in both the innate and adaptive immunity, is often a major contributor to the protection and survival of the organism (Janeway et al., 2001). 1.1.1. The Process of Leukocyte Trafficking to Tissues  One of the hallmarks of the inflammatory response is the process of leukocyte trafficking to tissues, a process which has been studied extensively in the course of the past several years. This process has been divided into four sequential steps: rolling, activation, firm adhesion, and transmigration (Fig. 1). Rolling involves reversible interactions between molecules on the surface of leukocytes, such as P-selectin glycoprotein ligand- 1 (PSGL- I), with their receptors on ECs  —  namely E- and P-selectin. The upregulation of selectins on the surface of ECs is one of  the initial events of an inflammatory cascade. The reversible interactions between selectins and their ligands enable the circulating leukocytes to slow down and roll along the vessel wall. Eventually, a leukocyte may encounter a stimulus which leads to its activation. This  2  stimulus is typically a chemokine which has been secreted within the tissue and has crossed into the vessel lumen, thus encountering the rolling leukocyte. In addition, ECs are induced by cytokines to synthesize chemokines and present them on their luminal surface to circulating leukocytes. Chemokines, which are a family of small chemotactic cytokines (typically 8-10 kilodaltons), are known to direct the trafficking of various leukocytes within the body. Furthermore, binding of chemokines to specific receptors on leukocytes induces the activation of leukocytes and de novo expression, upregulation, or mere clustering of a number of adhesion molecules on the surface of the leukocytes. These include molecules such as integrins and immunoglobulins. Under inflammatory conditions, blood vessel ECs also begin the synthesis or upregulation of adhesion molecules such as intercellular adhesion molecule-i (ICAM- 1) and vascular adhesion molecule-i (VCAM-1). Interactions between these adhesion molecules and their ligands on the surface of leukocytes lead to the firm adhesion of the leukocytes to the ECs and the arrest of leukocyte rolling. Subsequently, the leukocyte extends pseudopodia, crosses the EC layer, and transmigrates into the tissue. The migration route is either through the junctions connecting two adjacent endothelial cells (para-cellular migration), or through the plasma membrane and cytoplasm of a single endothelial cell (trans-cellular migration). Once in the tissue, the leukocyte typically travels down a chemokine concentration gradient in order to arrive at the site of infection or injury and exert its effector function (Janeway, 2001; and Smith, 2008). 1.1.2. Endothelial Cell Adhesion Molecules and Their Ligands The role of cell adhesion molecules (CAMs) in inflammation has been the subject of extensive investigation. Several CAMs are expressed on endothelial cells, which interact with their corresponding ligands on leukocytes under inflammatory conditions. The CAMs are generally divided into the three categories of selectins, integrins, and immunoglobulins.  3  i.l.2.1.Selectins Selectins are a group of C-type lectins that support leukocyte adhesion through the recognition of carbohydrate moieties on glycoproteins and glycolipids. The group consists of three members: L-selectin (expressed on the surface of circulating leukocytes), and P-and E selectins (expressed by EC5). P-selectin is constitutively expressed by ECs and platelets, and is stored in cytoplasmic granules (Weibel-Palade bodies in ECs and ct granules in platelets) and is mobilized to the surface within minutes of encountering an acute inflammatory stimulus. E selectin is expressed on the surface of ECs following stimulation with pro-inflammatory cytokines such as tumour necrosis factor-ct (TNF-ct), interleukin- 113 (IL-i f3) and other inflammatory mediators such as bacterial LPS. Selectins reversibly interact with carbohydrates or mucins under shear stress. Isomers of the sialyl-Lewis x (sLeX) moiety on the surface of glycoproteins such as PSGL-i are recognized by E- and P-selectin; alternatively, L-selectin interacts with mucins, such as CD34 and glycosylation-dependent cell adhesion molecule-i (G1yCAM- 1). Through these reversible interactions, selectins can mediate the rolling and initial tethering of leukocytes flowing in circulation at high speeds (Janeway et al., 2001; Smith, 2008). 1.1 .2.2.Integrins Integrins are a family of transmembrane heterodimers linked by non-covalent interactions. In vertebrates, 18 different c subunits combine with 8 f3 subunits to form 24 integrins involved in cell/cell interactions and cell/extracellular matrix (ECM) interactions. Whereas  13’  (also known as CD29) subunits are found in most cell types, the  f32  (CD 18) and  137  integrins occur exclusively in leukocytes. Combinations of c’. subunits L, M, X, and 4, with j3 subunits 1, 2, and 7, lead to the formation of five key inte grins involved in leukocyte/EC interactions: lymphocyte function-associated antigen-l (LFA-l; a.k.a. 2 ctij3 ) , macrophage-1 4  antigen (Mac-i; a.k.a.  XM 2 3 ),  p150,95  (ccx32)  and the  cL4  integrins: very late antigen-4 (VLA-4;  a.k.a. ) 1 f 4 cL 3 and VLA-7 7 (cr43 ) . LFA-1 (a.k.a. CD1 1aJCD18), VLA-4 (CD49d/CD29), and VLA 7 (LPAM- 1) are expressed on T cells and monocytes, whereas Mac-i (CD 11 b/CD 18) and p150,95 (CD ii c/CD 18) are expressed by cells of the monocytic lineage (including monocytes, macrophages and dendritic cells), PMNs, and natural killer (NK) cells (Smith, 2008). The function of integrins requires the activation of leukocytes, which in turn serves to increase integrin affinity for its ligands on ECs or the ECM. Although the mechanism of this process is not fully understood, it is known that integrin activation mediates the firm adhesion of leukocytes to ECs in the process of leukocyte extravasation into tissues. Integrins have been shown to play major roles in many pathological processes. For example, a role for LFA-l has been described in ischemic injury, arthritis, asthma, graft rejection, and cancer metastasis (reviewed by Mazzone & Ricevuti, 1995). Likewise, VLA-4 has been shown to mediate the pathophysiology of multiple sclerosis, inflammatory bowel disease, diabetes, and pulmonary allergic inflammation (Engelhardt & Kappos, 2008; Ghosh, 2003; Lobb et al., 1996; Michie et al., 1998). 1.1 .2.3.Immunoglobulins Members of the immunoglobulin (Ig) superfamily are expressed on ECs, leukocytes, and other cells, and they play a major role in EC/leukocyte interactions, and the firm adhesion and transmigration of leukocytes into tissues. ICAM-1 (also known  as  CD54) was the first to be  identified as the endothelial receptor for LFA- 1. In fact, ICAM- 1, -2, -3, -4, and -5, all bind LFA-1 with high affinity (Fawcett et al., 1992; Smith, 2008; Staunton et a!., 1989). ICAM-1 expression on ECs undergoes dramatic upregulation following the encounter of inflammatory stimuli, such as TNF-c, interferon-y (IFN-y), IL-i 3, and LPS. ICAM-2 is constitutively  5  expressed, suggesting a role in leukocyte trafficking to non-inflamed tissues. ICAM-3, which is expressed by most leukocytes, plays a role in leukocyte-leukocyte interactions. ICAM-4 and ICAM-5 are expressed in erythrocytes and neurons respectively (reviewed by Smith, 2008). VCAM-1 (CD1O6) is another member of the Ig superfamily, with 6 to 7 Ig domains, which is expressed in several cell types, such as bone marrow stromal cells, spleen stromal cells, thymic epithelial cells, peripheral lymph node (LN) and mesenteric LN high endothelial venules, and some dendritic cells in the spleen. It is induced on the surface of ECs following treatment with cytokines, such as IL-i 3, IL-4, TNF-a, and IFN-y. VCAM-l interacts primarily with VLA 4 on the surface of leukocytes, but is also known to bind some  f32  integrins (e.g.  ctx132  and  aD1 2 3 )  (Needham et al., 1994; Kilger et al., 1995; Soriano & Piva, 2008; Smith, 2008; Wu, 2007). Platelet/endothelial cell adhesion molecule (PECAM-i, CD31) is a 130 kD Ig in platelets, neutrophils, monocytes, NK cells, some T cells, and in EC junctions (between two adjacent ECs) (Mamdouh et al., 2003; Newman, 1997). PECAM-l is expressed at high levels in the kidney, lung, and trachea, and at lower levels in the heart, liver, and brain (Wang et al., 2003). Numerous studies have indicated a role for ICAM-i and VCAM-i in a variety of pathological processes, such as autoimmune diseases (e.g. multiple sclerosis, inflammatory bowel disease, and rheumatoid arthritis), infectious diseases (e.g. HIV- 1 and rhinovirus infections) and transplant rejection (reviewed by Yusuf-Makagiansar et al., 2002). For example, a therapeutic role for ICAM- 1 blocking has been suggested in rheumatoid arthritis, atherosclerosis, and liver and kidney transplants (Flavin et al., 1991; Haug et al., 1993). In addition, anti-ICAM- 1 therapy has resulted in reductions in myocardial infarct size (loculano et al., 1994; Yamazaki et al., 1993), and anti-VCAM-1 therapy has led to an amelioration of bronchial inflammation in experimental animal models (Lobb et al., 1996).  6  Similarly, PECAM- 1 has been implicated in the pathogenesis of several inflammatory disorders, including atherosclerosis, ischemic injury, rheumatoid arthritis, and septic shock (Graesser et al., 2002; Ishikaw et al., 2002; Maas et al., 2005). For instance, animal studies of lung disease and atherosclerosis have established an association between PECAM-1 deficiency and reduced inflammatory responses (reviewed by Woodfin et al., 2007).  1.1.2.4. DC-SIGN: A Novel Adhesion Molecule The dendritic cell-specific ICAM-3-grabbing non-integrin (DC-SIGN, also known as CD209) which is a member of the C-type lectin family, was discovered as a non-integrin ligand for ICAM-3 on a class of antigen presenting leukocytes known as dendritic cells (DCs) (Geijtenbeek et al., 2000a). This molecule, which was first described in the context of DC/T cell interactions and the initiation of a primary I cell immune response, has since been found to be involved in other immune processes, such as binding to the human immunodeficiency virus-I (HIV-1) and the immunoglobulin ICAM-2. The high affinity of DC-SIGN for ICAM-2 on the surface of ECs has defined the role of DC-SIGN as an adhesion molecule (Geijtenbeek et al., 2000b; Bleijs et al., 2001). Shortly after its discovery, DC-SIGN was shown to be involved in the trafficking of dendritic cells, and its participation in the inflammatory response is a subject of ongoing investigations (Geijtenbeek et al. 2000b). It has since been found that this molecule is involved in the pathophysiology of viral, fungal, and mycobacterial infections, as well as tumour recognition and autoimmune processes, and it plays a significant role in neutrophil-mediated immune responses (Aarnoudse et al., 2006; Buzas et al., 2006; Geijtenbeek et al. 2000c; Torrelles et aL, 2008; van Gisbergen et al., 2005; Willment & Brown, 2008).  7  Figure 1. Overview of Leukocyte Trafficking across ECs Step  Step 2: Activation  1:  Step 3: Firm  Step 4: Transmia ratio  Integrins: (e.g. LFA-i, Mac-i. VLA-4’)  Immunoglobulins: (e.g. CAM-i, ICAM-2, PECAM-i. VCAM-1  Immunoglobulins: (e.a. CAMs. PECAM-i)  Selectins (e.g. P-selectin,  E-seIectin -——*  EC  —*  ECM Components  .  0  Junctional Proteins  . Cytokines +0 and chemokines 0  1.2  •  .  .  0. 0 •0 .0  •  .  THE CENTRAL NERVOUS SYSTEM (CNS)  1.2.1. Inflammation and CNS Pathology The development of immune responses in the CNS is fundamentally different from other organs due to the presence of the blood-brain barrier (BBB) and specialized CNS cells, which, under normal conditions restrict the movement of leukocytes to the brain (Banks, 2006; Galea et al., 2007). In fact, the CNS has traditionally been considered “immunologically privileged”, since under  normal conditions leukocytes are rarely encountered, expression of the major histocompatibility complex (MHC) class II molecules (associated with professional antigen-presenting cells) is at low levels, and the production of cytokines and expression of endothelial cell adhesion molecules (eCAIVIs) is low or absent (Arvin et al., 1996; Hauser et al., 1983; Perry, 1998). Furthermore, early in-vivo studies have indicated that tumours and tissue grafts evaded immune recognition in the CNS (Medawar, 1948). Similar observations with certain bacteria and viruses served to perpetuate the  8  notion of “immune privilege” (Matyszak & Perry, 1998; Stevenson, 1997). In the past several years, however, a number of studies have challenged this concept. Indeed, it has been demonstrated that transplant rejection does occur in the CNS, similar to extracerebral organs (reviewed by Poltorak et al., 1997). Moreover, in response to infection, ischemia, trauma, and in autoimmune and degenerative CNS diseases, leukocytes readily migrate across the cerebral vasculature and the brain becomes the site of intense inflammation (Danton & Dietrich, 2003; Galea et al., 2007; Hafler et al., 2005; Kim, 2006; Nguyen et al., 2002; Wang et al, 2006; Zipp & Aktas, 2006; Zlokovic, 2008).  Hence, inflammation has increasingly been implicated as one of the  hallmarks of pathology in the CNS (Galea et al., 2007; Soriano & Piva, 2008; Zipp & Aktas, 2006). The molecular events that mediate inflammatory responses in the CNS are currently not well understood.  However, since cerebral ECs are the first resident cells of the CNS to encounter  circulating leukocytes, molecular changes of the BBB endothelium have been increasingly implicated in the pathogenesis of neuroinflammation (Quan, 2006).  1.2.2. The Blood-Brain Barrier (BBB): Structural and Functional Properties The BBB is a special property of the CNS microvasculature, which plays a crucial role in regulating cellular and molecular traffic into and out of the CNS and thus maintaining CNS homeostasis. In 1885, Ehrlich observed that the brain was not stained following a systemic injection of dyes. Several decades later, Reese and Karnovsky’s ultrastructural studies on mice demonstrated that the cerebral ECs lining the lumen of the CNS microvasculature formed the anatomical basis of the BBB (Reese & Karnovsky, 1967). The cerebral ECs, which are the interface between systemic circulation and the CNS parenchyma, serve to maintain the delicate chemical balance of the CNS environment and assist in the regulated transport of essential molecules into and out of the CNS. ECs are surrounded by  9  various other components of the neurovascular unit, including the basal lamina (composed of the basement membrane and ECM components such as laminin and collagen), pericytes (relatively undifferentiated mesenchymal cells implicated in the regulation of capillary blood flow and EC differentiation), and the end-feet of astrocytes (common CNS glial cells with many important functions) (Persidsky et al., 2006). Although these structures have all been implicated in the maintenance of BBB integrity, the barrier function is a direct result of the special structural properties of the CNS ECs, namely their specialized junctional complexes, insignificant vesicular transport, and specific transport systems. The junctional complexes of the BBB consist of tight junctions (TJs). These complexes serve not only to restrict the paracellular diffusion of hematogenous cellular and molecular components, but also to maintain the chemical segregation of the apical and basal microenvironments. The TJs consist of an assembly of transmembrane and cytoplasmic proteins, arranged into an intricate network of multiple, parallel, and interconnected barriers.  The  transmembrane component, which forms the physical barrier, is composed of the claudin family, occiudin, and junctional adhesion molecules (JAMs). This component is linked to the actin cytoskeleton by cytoplasmic accessory proteins, such as the zonula occludens-l, -2, and -3, AF6, 7H6, and cingulin. Several signaling pathways are involved in the regulation of TJ activity, including calcium-dependent pathways, serine, threonine, and tyrosine phosphorylation, as well as G-protein-mediated mechanisms. The maintenance of the BBB junctional complexes has led to an “epithelial-like” transendothelial electrical resistance of 1500  —  2000 c2.cm , which is in 2  sharp contrast to the resistance of 22-52 icm 2 observed in human umbilical vein endothelial cells (HUVEC) (Crone & Olesen, 1982; Jinga et a!., 2000; Persidsky et al, 2006). In addition to the physical barrier, the CNS ECs are equipped with specialized transport  10  systems, including carrier-mediated influx and efflux mechanisms, and receptor- and adsorptivemediated transcytosis.  Some of these mechanisms allow the passage of essential molecules  (such as glucose, amino acids, ions, nucleosides, and certain proteins such as insulin, transferrin, histone, and avidin) into the brain. Other systems (such as ABC transporters) serve to prevent a wide range of compounds (such as drugs and neurotoxins) from entering the CNS environment (reviewed by Begley & Brightman, 2003; and Zhang & Stanimirovic, 2005).  1.2.3. Cells Participating in CNS Inflammation 1.2.3.1. Astrocytes Astrocytes are the most common glial cells in the CNS and they are involved in a variety of vital functions, such as neuronal development and migration, neurotransmitter metabolism, maintenance of the pH and ion balance of the CNS, and regulation of the CNS vascular tone and neuronal synapses (Kim et al., 2006; Ullian et al., 2001). Astrocytic endfeet envelop more than 99% of the cerebral ECs (Hawkins & Davis, 2005). Thus, astrocytes have been implicated in the development and maintenance of the BBB (Janzer & Raff, 1997; Kacem et al., 1998). The role of astrocytes in CNS inflammation is not fully understood. They have recently emerged as immune regulators due to their expression of immune receptors (such as Toll-like receptors) and their ability to secrete cytokines and chemokines (e.g. IL-6, TNF-a, CCL2, CCL3, and CXCL1O) upon activation (Carpentier, 2005; Farina et al., 2007). There is also some evidence that they can activate T cells in the presence of adaptive immune cytokines (Carpentier, 2005). Although astrocytes can be induced to express MHC class II molecules necessary for antigen presentation, the present lack of consensus regarding their expression of co-stimulatory molecules (e.g. CD8O and CD86) warrants further studies regarding their antigen-presenting capacities (Dong & Benveniste, 2001; Pagenstecher et al., 2000; Piehl & Lidman, 2001).  11  1.2.3.2. Microglia Microglia are specialized cells of myeloid lineage with hematopoietic origins, which express the common myeloid Ag CD1 lb and low levels of the leukocyte common Ag CD45. Microglia have been established as the resident phagocytic and immunocompetent cells of the CNS. In response to inflammatory stimuli they become activated, which leads to enhanced phagocytosis, the production of numerous cytokines and chemokines, and the expression of Fcy and complement receptors (Aloisi, 2001; Kim & de Vellis, 2005). Several studies have established a role for microglia in CNS pathological processes. For instance, microglial pro-inflammatory cytokines have been linked to a variety of CNS diseases such as viral infections and neurodegenerative diseases (Dickson et a!., 1991; Gonzalez-Scarano & Baltuch, 1999; McGeer et al., 1993; Thomas, 1992). Other studies suggest that activated microglia are responsible for the production of neurotrophic substances necessary for neuronal survival (Elkabes et al., 1998; Miwa et al., 1997; Nakajima & Kohsaka, 1993). Microglia have also been implicated in guiding monocyte migration into the CNS (Peridsky et al., 1999). In addition to their role in innate immune responses, microglia can be induced to express MHC class II and the co-stimulatory molecules CD4O, CD8O, CD86, which are necessary for efficient Ag presentation and have been shown to directly stimulate T cell responses (reviewed by Aloisi, 2001). These findings provide support for the involvement of activated microglia in CNS pathophysiology as an Ag presenting cell (APC).  1.2.3.3. Monocytes and Other Myeloid Cells Blood monocytes act as the precursors to several classes of myeloid cells, including macrophages, neonatal microglia, and dendritic cells. They are a heterogeneous population of bone-marrow derived cells distinguished by their surface expression of CD 14.  12  Monocytes  (which are capable of harboring viral infections) have been proposed to play a role in the pathogenesis of HIV and simian immunodeficiency virus (SIV) encephalitis (Kim et al., 2003). CNS macrophages, which derive from monocytes, are either observed in the meninges and the choroid plexus, or within the perivascular space, under the basement membrane surrounding the ECs (Hickey & Kimura, 1988; Hickey et a!., 1992; Williams & Hickey, 2002). Perivascular macrophages, which are also sometimes known as perivascular microglia, are phenotypically distinguished from parenchymal microglia by their high-level expression of the leukocyte common Ag (LCA; a.k.a. CD45) in humans and rodents (Sedgwick et a!., 1991; and Ulvestad et al., 1994). Functions such as the activation of CNS microglia and the production of chemokines have frequently been associated with perivascular macrophages (Polfliet et a!., 2002). These cells have also been suggested to play a role in directing T cell migration and Ag presentation, due to their strategic location and their expression of MHC class II and co stimulatory molecules (Aloisi, et a!., 2000; Ante! & Prat, 2000; Tran et al. 1998).  The  macrophage populations of the meninges and the choroid plexus have also been found to display MHC II and phagocytic activity. Similar to microglia, CNS macrophages have been associated with the pathogenesis of several CNS diseases, including autoimmune, infectious, and degenerative conditions (Chavarria & Alcocer-Varela, 2004; Kim et al., 2003; Zlokovic, 2008). A pivotal role for macrophages has been described in the processes of demyelination and axonal damage associated with multiple sclerosis (MS) and its animal model, experimental autoimmune encephalomyelitis (EAE).  These processes are likely carried out by the innate  immune functions of the CNS macrophages, such as the secretion of proinflammatory cytokines, free radicals, excitatory neurotransmitters, and matrix metalloproteases (reviewed by Hendriks et a!., 2005).  It has even been found that the depletion of macrophages leads to a complete  13  disappearance of clinical symptoms in EAE (Huitinga et a!., 1990), which further highlights the important role of this cell type in CNS inflammation. Myeloid dendritic cells are another group of cells related to the monocyte/macrophage lineage. These cells have also been implicated in CNS inflammation (Karman et al., 2004a), and will be discussed in detail in section 1.5.  1.2.3.4. Lymphocytes The participation of lymphocytes, particularly that of T cells, in CNS inflammation has been extensively studied over the years. T lymphocytes are derived from hematopoietic stem cells and differentiate into CD4 (helper) or CD8 (cytotoxic) subtypes in the thymus, based on their affinity for MHC class II or MHC class I molecules, respectively. Once naïve T cells recognize Ags presented on an APC’s MHC molecules, the process of cell-mediated immunity is initiated. Following initial priming, T cells acquire an immunological “memory”, which enables them to launch a more rapid Ag-specific response upon subsequent encounters with the same Ag. Many studies have indicated that the CNS is under constant immune surveillance by T cells. Activated T cells are able to enter the CNS irrespective of their Ag specificity (Wekerle, 1987), but their retention in the CNS occurs only upon Ag recognition (Hickey et al., 1991). Numerous studies have hence suggested a role for T lymphocyte-mediated immune responses in CNS inflammatory conditions, including MS and its animal models, viral infections (including HIV encephalitis and viral meningitis), neurodegenerative disorders (including Alzheimer’s disease (AD) and Parkinson’s Disease (PD)), ischemic injury, and tumour pathogenesis (Arumugam et al., 2005; Czlonkowska et a!., 2002; Lahrtz et a!., 1998; Rafalowska, 1998; Schneider-Schaulies, 2001; Walker et al., 2003; Weiner & Selkoe, 2002). The involvement of B cells and plasma cells in infectious conditions such as viral  14  encephalitis and sleeping sickness has long been established (Griffin et al, 1992, Hooper et al., 1998; Pentreath et al., 1994). In recent years, the discovery of anti-myelin antibodies (Abs) in MS patients and EAE animals has also generated renewed interest in investigating the involvement of these cells in CNS autoimmunity (Ziemssen & Ziemssen, 2005).  1.2.4. Inflammatory Cytokines in the CNS Several cytokines have been implicated in CNS inflammation due to their ability to mediate immune responses. Activated infiltrating leukocytes as well as the resident cells of the CNS both express and respond to a number of cytokines, including TNF-cL, IFN-y, transforming growth factor-13 (TGF-13), and several interleukins, under inflammatory conditions.  There is  even evidence for the activation of CNS cells by peripheral cytokines despite the presence of the BBB (Arvin et al., 1996; Gibson et al., 2004; Wilson et a!., 2004).  1.2.4.1. Tumor Necrosis Factor (TNF)-a TNF-ct. is synthesized as a transmembrane protein with 157 amino acid residues that is cleaved by proteolysis and exists in soluble form as a homotrimer (Sriram & O’Callaghan, 2007).  Its discovery goes back to the early observation of occasional tumour regression  following acute bacterial infections (Coley, 1893).  This cytokine is mainly produced by  monocytes and macrophages, but T cells, natural killer (NK) cells, smooth muscle cells, and epidermal cells are also capable of its production. TNF-a exerts its functions through binding as a trimer to its high-affinity receptors TNF receptor-i (TNFR- 1) and TNFR-2, and activating a number of proteins downstream, such as transcription factors (e.g. nuclear factor-icB and activator protein-i), protein kinases (e.g. extracelluiar signal-regulated kinases (ERKs)), phospholipases (e.g. PLA2 and PLC), and caspases (Darnay & Aggarwal, 1999; Vilcek & Lee,  15  1991). Thus, this potent inflammatory cytokine can play prominent roles in a wide array of biological processes, including the activation of inflammatory cells, induction of acute-phase protein secretion, vascular permeability, production of oxygen and nitrogen radicals, and regulation of cell proliferation, necrosis, and apoptosis (Idriss & Naismith, 2000; Tracey & Cerami, 1993). There is a wealth of evidence surrounding the physiological and therapeutic roles of TNF-cL, such as its involvement in embryonic development (Wride & Sanders, 1995), sleep regulation (Krueger et al., 1998), and resistance to infections and tumours (Aggarwal & Vilcek, 1991; Vilcek & Lee, 1991). On the other hand, TNF-a has emerged as a causative agent of morbidity and mortality in infectious diseases (Fiers, 1991). There has also been increasing evidence for the involvement of TNF-a in CNS pathology. The production of this cytokine in the CNS may also be carried out by astrocytes and microglia. A neurotoxic role for TNF-a has been documented in a multitude of CNS diseases, including MS and EAE, neurodegenerative diseases (e.g. AD and PD), stroke and its animal models, traumatic injury, and infectious diseases (e.g. bacterial meningitis and HIV infection). Furthermore, some studies have found TNF-a to play a protective role in experimental models of demyelination, AD, and excitotoxicity (Arvin et al, 1996; Sriram & O’Callaghan, 2007). Therefore, TNF-cL has been established as an active participant in CNS inflammation.  1.2.5. CAMs in CNS Inflammation A major role for CAMs in CNS inflammation has been defined by various groups. Previous in-vitro work in our laboratory has shown that the trafficking of T cells and PMNs across brain ECs that are treated with inflammatory cytokines is affected significantly by endothelial CAMs (eCAM5) (Wong et al, 1999, 2007).  Furthermore, in-vivo studies have  indicated that T cell adhesion to brain ECs is regulated by ICAM-1/LFA-1 and VCAM-1IVLA-4 16  interactions, whereas the transmigration process only utilizes the ICAM- 1 /LFA- 1-dependent pathway (Engelhardt, 2006; Laschinger et al., 2002; Pryce et al., 1997; Tsukada et al., 1993). Other in-vivo studies have shown that blocking ICAM-1 and 132 integrins leads to a significant reduction in leukocyte infiltration, brain edema, and infarct volume in animal models of stroke (Soriano et a!., 1996, 1999). Similarly, in animals with cerebral malaria and bacterial meningitis, 132 integrin blockage led to a decrease in leukocyte infiltration, brain edema, and mortality (Grau et al., 1991; Tuomanen et a!., 1989). A clear role for anti-VLA-4 Abs has also been shown in preventing the adhesion of lymphocytes and monocytes to brain ECs and in the reversal of clinical symptoms in EAE animals (Cannella & Raine, 1995). Furthermore, an anti VLA-4 Ab (Natalizumab) is currently being used as a relatively effective treatment for patients with relapsing-remitting MS (reviewed by Engelhardt & Kappos, 2008). In addition, ICAM-1, VCAM-1, and PECAM-1 levels have all been shown to be elevated in MS patients. PECAM-1 has also been implicated in the pathogenesis of cerebral ischemia, AD, and HIV-1 encephalitis (reviewed by Kalinowska & Losy, 2006). These few examples serve to highlight the important role of eCAMs and their ligands in the process of CNS inflammation.  1.3.  HUMAN BRAIN MICROVESSEL ENDOTHELIAL CELLS (HBMEC)  1.3.1. Endothelial Cell Heterogeneity The ECs lining the lumen of all blood vessels play a role in several physiological and pathological conditions, including homeostasis, coagulation, angiogenesis, and the regulation of leukocyte trafficking.  ECs from different vascular beds and different species vary in their  morphological and functional properties (Zetter, 1988). The two broad categories of ECs are large vessel and microvessel ECs. Large vessel ECs are responsible for physiological functions such as the maintenance of vascular tone. Based on their morphology, microvascular ECs are  17  further divided into three categories: fenestrated, discontinuous, and continuous ECs. Tissues and organs undergoing high rates of molecular and cellular exchange, such as the gastrointestinal tract, kidney glomerulus, and choroid plexus) are lined by discontinuous and fenestrated ECs. On the other hand, the delicate homeostasis of the CNS is maintained by the continuous ECs of the BBB which are bound by tight junctional complexes (Risau, 1995; Risau, 1998).  1.3.2. Role of Endothelial Cells in CNS Inflammation  The role of cerebral ECs in CNS inflammation is illustrated through the capacity of this cell type to express and respond to a variety of cytokines, chemokines, and adhesion molecules under inflammatory conditions.  Furthermore, alterations in the barrier properties of the brain ECs  constitute one of the hallmarks of neuroinflammation.  The mechanisms responsible for the  increased BBB permeability have not been fully elucidated, although it is likely a combination of increased junctional permeability and migration of inflammatory cells. Besides establishing the important role of eCAMs (such as ICAM-1, VCAM-1, PECAM-1, and E-selectin) in the processes of leukocyte trafficking to the CNS (Wong et al., 1999, 2007), our laboratory has also shown that brain ECs synthesize, secrete, and bind several  I chemokines  (such as CCL3, CCL4, and CCL5), thus further influencing the adhesion and migration of CD4 T cells across the BBB (Quandt & Dorovini-Zis, 2004; Shukaliak & Dorovini-Zis, 2000).  1.3.3. In-vitro Model of the Human Blood Brain Barrier  An in-vitro model of the human BBB has previously been developed in our laboratory. In this model, primary cultures of human brain microvessel ECs isolated from autopsy brains or temporal lobectomy specimens form confluent monolayers that retain important properties of the BBB in vivo, namely a paucity of pinocytotic vesicles, a high transendothelial electrical  18  resistance, and the presence of tight junctional complexes that restrict the paracellular passage of macromolecules such as horseradish peroxidase (Dorovini-Zis et al., 1991). The purity of this EC culture is established by the expression of von Willebrand Factor (Factor VIII), the binding of Ulex europaeus agglutinin, and the uptake of acetylated low density lipoprotein. This model has been replicated several times and has been utilized repeatedly for the study of BBB under physiological and inflammatory conditions, and the process of leukocyte trafficking across the BBB (Huynh & Dorovini-Zis, 1993; Wong & Dorovini-Zis, 1992, 1995, 1996a, 1996b).  1.4.  ANTIGEN PRESENTATION IN THE CNS Antigen (Ag) presentation is the critical process through which T cell immune responses  are generated, be it the protective response against infections and tumours or the destructive immune response in autoimmunity. Once an Ag is taken up by an APC, it is processed and loaded on the surface of MHC molecules. MHC class I molecules, which are expressed on the surface of most nucleated cells, are classically described as specialized for the presentation of Ags synthesized in the cytosol (such as viral proteins) and stimulate CD8 T cells. On the other hand, MHC class II are specialized for the presentation of exogenous peptides from intracellular vesicles.  Therefore, MHC class II can present Ags derived from pathogens that have been  phagocytosed by macrophages, B cells, or DCs.  This molecule is thus associated with  “professional” antigen presentation, and binds to CD4 cells. Frequently, exogenous peptides are also presented through the MHC class-I-mediated pathway, a phenomenon referred to as cross-presentation (Groothuis & Neefjes, 2005; Kasturi & Pulendran, 2008). Apart from interacting with their cognate Ags presented on an MHC molecule, T cells also require “co-stimulatory signals” in order to be activated, the lack of which has been associated with T cell apoptosis and immune tolerance. Proper Ag presentation involves the  19  interactions of certain CAMs (e.g. LFA-1 (CD1 la), LFA-3 (CD58), ICAM-1 (CD54)) and co stimulatory molecules (CD4O, CD8O, and CD86) on the surface of the APC with their receptors on the T cell, including CD28 and CTLA-4 (Slavik et al., 1999; van Kooten and Banchereau, 2000). This leads to the activation of signal transduction pathways which in turn lead to the activation, proliferation, and differentiation of the T cell, as well as cytokine production. Although Ag presentation can take place within tissues or inside secondary lymphoid organs, recent evidence suggests that the presence of local APCs is required for an optimal T cell response to cutanous Ags in vivo (Itano et al., 2003).  1.4.1. Ag Presentation and CNS Pathology  Whether the nature of CNS Ag presentation differs from that of peripheral organs has been the subject of much interest and investigation over the course of the past several years. The presence of resident CNS cells that perform immune functions such as astrocytes and microglia, as well as the cellular restrictions imposed by the BBB create a unique situation for CNS Ag presentation (Hart & Fabry, 1995). Previous studies in the field have shed light on some immunological properties of the CNS, such as the crossing of the BBB by leukocytes (Bechmann, 2005; and Hickey, 2001), the draining of CNS Ags into cervical lymph nodes via the cribriform plate and perineural sheath of cranial nerves (Cserr, & Knopf, 1992), and the existence of similar migration patterns for APCs (Hatterer et al, 2006; and Karman et al., 2004b). Therefore, although it is believed that T cell activation and expansion occurs within lymphoid organs and not in the CNS, the CNS may still be the site where neuroinflammation is initiated (Becher et al., 2006; Karman et al., 2004b). Ag presentation by the different cells of the neuro-immune compartment seems to be a critical event in several CNS pathological conditions such as MS and infectious diseases (Antel  20  & Prat, 2000; DOrries, 2001; Traugott, 1987) and even neurodegenerative diseases and tumours (Badie & Schartner, 2001; Minagar et al., 2002). Also, evidence from animal studies suggests that Ag presentation occurs prior to the onset of clinical symptoms (Ponomarev et al., 2005).  1.4.2. Ag Presenting Cells of the CNS  1.4.2.1. Microglia Numerous lines of evidence have established a role for microglia in CNS Ag presentation. Microglia can be induced to express MHC Class II and several adhesion and co stimulatory molecules (such as  f32  integrins, CD4O, CD8O, CD86, CD54, and CD58) and have  been shown to actively participate in AD, PD, MS, HIV infection and stroke pathology (Aloisi, 2001; Katz-Levy et a!., 1999; Kim & de Vellis, 2005; McGeer et al., 1988; Stoll & Jander, 1999).  Studies in EAE show an increase in microglial expression of the non-classical CD1  family of lipid antigen-presenting molecules, suggesting that microglia may participate in the presentation of a variety of antigens during CNS inflammation (Cipriani et al., 2003). The ability of microglial cells to directly stimulate CD4 T cell (both Thl and Th2) responses has also been suggested by several groups (reviewed by Aloisi, 2001).  These findings provide  support for the involvement of activated microglia in CNS pathophysiology as an APC.  1 .4.2.2.Perivascular Macrophages Macrophages residing in the perivascular space are likely the first group of APCs which is encountered by activated T cells entering the CNS, and they have thus been implicated in CNS Ag presentation (Perry, 1998). Indeed, perivascular macrophages expresses MHC class II even under non-pathological conditions, and they show upregulation of MHC class II expression following CNS injury (Ante! & Prat, 2000; Streit et al., 1989). Furthermore, there is constitutive  21  expression of the co-stimulatory molecule B7.2 (CD86) on this cell population; and this is complemented by an inducible expression of B7. 1 (CD8O) under inflammatory conditions, such as MS (Williams et al., 1993). It has also been suggested that perivascular macrophages migrate to secondary lymphoid organs in order to present Ags subsequent to phagocytosis  —  a finding  that is yet to be confirmed (Broadwell et al., 1994; Matyszak & Perry, 1995; Perry, 1998).  1 .4.2.3.Endothelial Cells Findings from extra-cerebral ECs have indicated that human ECs are capable of presenting Ags to T cells and stimulating their proliferation and differentiation (Biedermann & Pober, 1998 and 1999). Furthermore, expression of MHC molecules by vascular ECs suggests that these cells may play a role in presenting Ags to the T cells in peripheral circulation (i.e. memory T cells) (Hayry et al., 1980; Natali et al, 1981; Pober, 1999). Several studies have investigated the possibility of Ag presentation by cerebral ECs through examining the expression of MHC class I, MHC class II, and co-stimulatory molecules under inflammatory conditions (Hoftberger et al., 2004; Huynh & Dorovini-Zis, 1993; Oman & Dorovini-Zis, 2001). Previous work from our laboratory has shown that treatment of brain ECs with IFN-y leads to their de novo expression of MHC class II (Huynh & Dorovini-Zis, 1993). Furthermore, cerebral ECs have been shown to display de novo or upregulated expression of the co-stimulatory molecules CD8O, CD86, CD4O, and LFA-3 in the presence of IFN-y, and to stimulate the proliferation of CD4 T cells in vitro (Oman et al., 1999; 2001; 2003).  1.4.2.4. Dendritic Cells Dendritic cells (DC5) are the most potent APCs in the immune system, because of their ability to endocytose and process antigens, express high levels of MHC class II and co  22  stimulatory molecules, and their ability to migrate to secondary lymphoid organs to efficiently activate naïve T cells and induce their proliferation and differentiation (Aloisi, 2001). DCs have been shown to be capable of stimulating various subsets of the CD4+ T helper cell family, inducing Thi, Th2, and Th17 type responses as well as regulatory T cell and cytotoxic T cell responses, thus affecting a wide range of pathological processes from infectious and allergic inflammation, to tumours and autoimmune responses in the CNS and other tissues (Banchereau & Steinman, 1998; Baumgart & Carding, 2007; Cheung et al., 2008; Heufler et al., 1996; Mantovani et al., 2008; Skallova et a!., 2008; Steinbrink et al., 1997).  1.5.  DENDRITIC CELLS (DCs)  1.5.1. Role of DCs in Immunity and Tolerance Dendritic cells (DC5) are migratory bone marrow-derived cells which play a key role in the innate and adaptive immune responses as well as in the induction of immune tolerance (Chavarria and Alcocer-Varela, 2004; Reis e Sousa, 2006; Suter et al., 2003). The classical description of DCs categorizes them into two developmental stages.  Immature DCs or their  precursors behave as phagocytes in peripheral tissues and they have a low capacity for Ag presentation and T cell stimulation. Upon encountering pathogenic or tissue damage-associated signals such as cytokines and bacterial products (e.g. LPS) DCs become activated, release a variety of inflammatory mediators, and begin to migrate towards secondary lymphoid organs. This process is typically accompanied by “maturation”, a number of morphological and phenotypic changes that are important to DCs’ function as APCs.  The hallmarks of maturation include an increased  expression of class II MHC, co-stimulatory molecules (CD4O, B7.1, B7.2, CD83), and the chemokine receptor CCR7, and a decreased expression of DC-SIGN and several chemokine receptors (such as CCRI, CCR2, and CCR5) (Guermonprez et al., 2002; Hsieh et al., 2001;  23  Plumb et al., 2003; Sallusto et a!., 1998; Serafini et al., 2006; Sozzani et al., 1998). Other important characteristics of mature DCs are the expression of the chemokine receptor CCR7 and the actin-bundling protein fascin (Dieu et a!., 1998; Zhang et al., 2008). CCR7 is the receptor for chemokines CCL 19 and CCL2 1 (which are expressed in secondary lymphoid organs), and thus it orchestrates the migration of DCs to the secondary lymphoid organs. Fascin, on the other hand, is an intracellular globular molecule. It is a 55 kDa actin-bundling protein, which functions to organize F actin filaments into orderly bundles. It also appears at cell protrusions, and spikes on the leading edges of motile cells. Fascin also plays a role in the adhesion of the cell to extracellular matrix components, and therefore it may play a role in the migration of mature DCs (Kureishy et a!., 2002). Once in the secondary lymphoid organs, DCs efficiently activate Ag-specific T cells and prime naïve T cells. Adaptive immunity is thus induced as activated lymphocytes migrate to the site of infection or injury to launch an Ag-specific immune response (Chavarria and Alcocer-Varela, 2004; Del Prete et al., 2006; Guermonprez et a!., 2002; Inaba et al., 2000). It is important to keep in mind that maturation, far from referring to a discrete step in DC development, encompasses a wide range of changes spanning a spectrum of morphological and phenotypic characteristics (Braun et al., 2006; Reis e Sousa, 2006) A major role for DCs has also been described in the process of immune tolerance. DCs are capable of dampening T cell responses through a variety of mechanisms, such as clona! deletion, and the induction of T cell anergy and regulatory T cells (Liu, et al. 2002; and Yamazaki, et a!., 2003).  1.5.2. DC Heterogeneity DCs comprise a heterogeneous population with differences in origin, phenotype and function. There are also differences in DC populations between species. Mouse DCs have been extensively studied over the years, and despite their differences with human DCs, they have been  24  utilized to investigate many aspects of DC biology (reviewed by Wilson & O’Neill, 2003). The human DC populations are divided into the two broad categories of myeloid and plasmacytoid DCs.  Both are the descendents of CD34 hematopoietic stem cells, and are  distinguished by their expression of the C-type lectin DC-SIGN (also known as CD209) (Geijtenbeek et al., 2000a; McMahon et al., 2006). Myeloid DCs are either differentiated from blood monocytes within tissues, or are derived directly from the CD34 precursors and circulate in peripheral blood (sometimes referred to as pre-DC) (Shortman & Naik, 2007). The myeloid DC population encompasses several other subsets, such as Langerhans cells (epidermal DCs), dermal DCs, and Kupfer cells in the liver.  These subsets have been shown to be activated  following encounters with inflammatory stimuli, and they subsequently function in inflammatory processes, via the production of inflammatory cytokines, and activation of T cells. Phenotypically, myeloid DCs are mostly CD4LinCD1 1c+CD123m and  CD45ROCD2.  (reviewed by McMahon et al., 2006; and Shortman & Naik, 2007). Plasmacytoid DCs, on the other hand, are distinguished by their lower expression of CD1 ic and high surface expression of CD123 (IL-3 receptor). They are a circulating leukocyte population and are the most important source of type I interferons (IFN-cL and  )  in the body.  Earlier experiments had found that plasmacytoid DCs were capable of inducing Th2 responses, whereas Thi responses were linked solely to myeloid DCs (Rissoan et al., 1999). However, a role in inflammatory Thi and tolerogenic processes has been increasingly attributed to human plasmacytoid DCs (Arpinati et al., 2003; Cella et al., 2000; Kawamura et al., 2006; Krug et al., 2001).  Although it was once believed that plasmacytoid DCs are exclusively derived from  lymphoid progenitors, there is now evidence suggesting that both the common lymphoid  25  progenitor and the common myeloid progenitor have the potential to give rise to either DC category (reviewed by Takeuchi & Furue, 2007; and Wu & Liu, 2007).  1.5.3. In-Vitro Generation of DCs from Monocytes The small number of DCs in the human peripheral blood has made the in-vitro generation of DCs from peripheral blood monocytes a common experimental practice for a number of years. The in-vitro-generated DCs have been shown to be a good model for studying the properties and behaviour of myeloid DCs under inflammatory conditions (Shortman & Naik, 2007). The in-vitro generation technique involves culturing isolated peripheral blood monocytes in the presence of cytokines or bacterial LPS for a period of time which is typically between 2 and 7 days. Over the course of the years, various methods and various combinations of reagents have been used for the generation of DCs in vitro. This has primarily been carried out by culturing freshly-isolated monocytes in the presence of granulocyte-macrophage colony stimulating factor (GM-CSF), often in combination with other molecules, such as TNF-a, IL-4, IL-b, and TGF-13. To induce the maturation of DCs, bacterial LPS has frequently been used. Various cytokine cocktails have also been used for this purpose, including a combination of TNF-c, IL-i I, and IL-6 (which are the components of monocyte-conditioned media), and T cell-conditioned media (Kato et al., 2001; Reddy et al., 1997)  .  Prostaglandin E2 (PGE ) is 2  another molecule which has been found to exert some affect on the maturation process of DCs, and thus it has been utilized in some studies in addition to other reagents in order to induce the maturation of monocyte-derived DCs (Feuerstein et al., 2000).  26  1.5.4. Cytokines Involved in DC Differentiation and Maturation 1.5.4.1. GM-CSF Granulocyte-macrophage colony stimulating factor (GM-CSF) is a 14.5-35 kDa monomeric glycoprotein, which, as its name suggests, was originally discovered through its involvement in the differentiation of both granulocytes and macrophages from mouse hematopoietic stem cells following LPS injection (Burgess et al., 1977). It has since been shown to affect the survival and activities of mature myeloid cells, such as granulocytes, macrophages, and eosinophils (Handman & Burgess, 1979; Hamilton et al., 1980; Simon et al., 1997). GM-CSF was classified as a pro-inflammatory cytokine shortly after its discovery (Hamilton et al., 1980). Although it is not essential for the formation of myeloid cells under steady-state conditions (Vremec et a!., 1997), GM-CSF production by various cell types (such as macrophages, mast cells, T cells, fibroblasts, and EC5) undergoes dramatic upregulation in response to inflammatory stimuli (Cousins et al., 1994; Nimer & Uchida, 1995). This molecule has been found to play an important role in a wide array of pathologies, including rheumatoid arthritis, and autoimmune renal and pulmonary diseases which highlights its role as an inflammatory cytokine (Hamilton & Anderson, 2004). In the normal CNS, astrocytes are chiefly responsible for the production of GM-CSF (Malipiero, 1990). However, peripherally-produced GM-CSF is also capable of entering the CNS by crossing the BBB (McLay et a!., 1997).  In response to inflammation, GM-CSF is  produced in large amounts by activated brain ECs and activated T cells, and delays the apoptotic program of recruited neutrophils, thus prolonging their inflammatory activities (Coxon et al., 1999; Shi et al., 2006). Furthermore, GM-CSF has been shown to activate and prime microglia for Ag presentation (Re et a!., 2002).  Elevated GM-CSF levels are also observed in the  27  cerebrospinal fluids of patients with MS, stroke, AD, and vascular dementia (Carrieri et al., 1998; Tarkowski et al., 1997; 2001). In addition, GM-CSF null mice have been shown to be resistant to EAE and unable to sustain leukocyte trafficking to the CNS (McQualter et al., 2001). Taken together, these findings strongly suggest a role for GM-CSF in CNS inflammation. In the generation of functional DCs from monocytes, GM-CSF (in combination with IL4) has long been shown to be an effective agent (Sallusto & Lanzavecchia, 1994). Furthermore, it has been demonstrated that human bone-marrow-derived CD34 cells can transform into DCs in the presence of various cytokines, including a combination of GM-CSF and TNF-a (Caux et al., 1996). Since GM-CSF-generated CD11b DCs are able to potently stimulate inflammatory Thi as well as Th17 responses, they are considered to be a good model for myeloid DCs under inflammatory conditions in vivo (Bailey et al., 2007; Boonstra et al., 2003).  1.5.4.2. IL-4  Interleukin-4 (IL-4) is a 20 kDa cytokine secreted by subsets of Th2 helper T cells and mast cells.  It is composed of four antiparallel ct-helices and two long end-to-end ioops  connected by a short 13-sheet against the helices (Mueller et al., 2002). IL-4 contains 6 cystein residues forming the disulphide bonds necessary for its biological activity (Sredni-Kenigsbuch, 2002).  This molecule fulfills diverse functions, such as promoting the proliferation and  differentiation of activated lymphocytes and the differentiation of PMNs and monocytes, enhancing the Ag presenting capacity of B cells, the chemoattration of fibroblasts, and the inhibition of several pro-inflammatory cytokines, such as IFN-y and IL- 12 (Van Meir, 1995). In the CNS, IL-4 expression is upregulated following injury as well as infectious and neurodegenerative conditions (Woodroofe & Cuzner, 1993). It is believed to be produced by infiltrating T cells, and exerts its anti-inflammatory functions by controlling glial cell  28  proliferation, inhibiting Ag presentation through downregulating MHC class II, and blocking the production of nitric oxide (Iwasaki et al., 1993). In contrast to the effects of GM-CSF, EAE prone mice lacking IL-4 have been found to display more severe disease compared to their wildtype counterparts (Bettelli et al., 1998; Falcone et al., 1998). There is an interesting interplay between IL-4 and GM-CSF production in vivo. It has been shown that IL-4 inhibits the expression of GM-CSF (Akashi et al., 1991; Jansen et al., 1989), and one study has suggested an increase in IL-4 levels following GM-CSF gene transfer in the mouse lung (Stampfli et al., 1998). Although the relevance of these interactions in the process of DC differentiation in vivo has not been established, it is known that in the context of human DC generation from monocytes in vitro, IL-4 acts as an inhibitor of macrophage colony formation in addition to its role in inducing DC growth and differentiation (Romani et al., 1994). In the presence of GM-CSF alone, DC precursors in blood almost entirely develop into the macrophage family of cells (Jonuleit et al., 1996).  1.5.4.3. TNF-a  In addition to its many roles in physiological and pathological conditions (some of which were described in section 1.2.4.1.), TNF-a has been shown to play a key role in the differentiation of human CD34 precursors (from both cord blood and adult bone marrow) into myeloid DCs (Caux et al., 1992). In fact, together with GM-CSF and IL-4, TNF-u is the most widely-used cytokine for the generation of DCs in vitro (Zou & Tam, 2002). It is also possible to differentiate monocytes directly into mature CD83 DCs by culturing them in GM-CSF, IL-4 and TNF-cL for 10-12 days (Zhou and Tedder, 1996). Various concentrations of TNF-a can also induce the terminal maturation of immature human DCs in vitro, but only in combination with other soluble mediators (Feuerstein et al., 2000; Kato et al., 2001; Reddy et al., 1997).  29  Furthermore, it is believed that TNF-ct is involved in the in-vivo maturation of DCs and inducing their Ag-presenting and migratory behaviours (Jonuleit et al., 1996).  1.5.4.4. IL-113 IL-i 13 is a 1 7-kDa pro-inflammatory cytokine, with a multitude of innate and adaptive immune functions involving various target cells, including leukocytes (Dinarello, 1996).  It  signals through its high-affinity receptor known as type 1 IL-i receptor (IL-1R1), and is capable of inducing several genes via the activation of transcription factors such as NF-icB, APi, and EBPf3 (Iwasaki et a!., 1992; Martin & Wesche, 2002; O’Neill & Greene, 1998). In the CNS, IL-i 13 and its receptor have been identified in various cell types including astrocytes, microglia, perivascular cells, cerebral ECs, and even neurons (Konsman et al., 2007; Rothwell, 1991). This cytokine has also been implicated in a variety of CNS diseases, such as AD, MS, ischemia, infections, seizure, head injury, and fever (Bartfai et a!., 2007; Dickson et al., 1993; Dominguez-Punaro, 2007; McClain et al., 1987; Rothwell & Relton, 1993; Tsukada et al., 1991; reviewed by Konsman et al., 2007). Similar to TNF-ct, IL- 113 acts on cerebral ECs to upregulate the expression of eCAMs (Baumann & Gauldie, 1994). IL- 113’s role in DC maturation was first described in a murine cell line (Yamada & Katz, 1999). It has also been frequently used as a component of various cytokine mixtures in order to induce the maturation of human monocyte-derived immature DCs, often in combination with TNF-a and IL-6 (Feuerstein et al., 2000; Reddy et a!., 1997).  1.5.4.5. IL-6 Interleukin-6 (IL-6) has been characterized as a cytokine with both proinflammatory and anti-inflammatory properties (Akira et al., 1990). It is a 26 kDa molecule which regulates genes  30  involved in cellular proliferation, differentiation, survival, and apoptosis. IL-6 is secreted by many cells, including lymphocytes, monocytes, fibroblasts, ECs, mesenchymal cells and some tumor cells (Blanco et al., 2008). This cytokine is involved in hematopoiesis, as well as in many innate and adaptive immune functions, such as promoting the differentiation of B cells, helper and cytotoxic T cell subsets, macrophages, and megacaryocytes (Kishimoto et al., 1995). It signals through the gpl 30 receptor, which in turn activates the JAKISTAT or MAPK pathways (reviewed by Heinrich et a!., 2003). Similar to IL-i 3, IL-6 is produced in the CNS by astrocytes, microglia, neurons and ECs (Fontana et al., 1989; Rott et a!., 1993). Its production may be induced by a variety of stimuli, including the presence of micro-organisms, traumatic injury, and other cytokines (such as TNF-ct and IL-113). It affects the differentiation of neurons and the proliferation of astrocytes, and has been described as both a neurotrophic and neurodegenerative agent.  Its anti-inflammatory  effects include the suppression of pro-inflammatory cytokine and free radical production by CNS cells, but elevated levels of IL-6 have also been reported in several CNS disorders, such as MS and EAE, viral and bacterial infections, as well as AD (reviewed by Sredni-Kenigsbuch, 2002). IL-6 has been observed to promote the differentiation of CD34 precursors into functional DCs, together with GM-CSF (Bernhard et al., 2000). In contrast, it has also been found to inhibit the differentiation of monocytes into DCs, an effect which was abrogated in the presence of TNF-a, IL-i 13, CD4OL, and LPS (Chomarat et a!., 2000). In combination with TNF ct, IL-i 13, and IFN-cL, various concentrations of IL-6 (from 6ng/m! to 1 gIml) have been shown to lead to the maturation of human monocyte-derived DCs (Reddy et a!., 1997). Variants of this combination have also been used as an effective “maturation stimulus” in other studies of DCs (Jarnjak-Jankovic et al., 2007; Jonuleit et a!., 1997; Thurner et a!., 1999).  31  1.5.4.6. POE 2 Prostaglandin E2 (POE ) is a member of the prostaglandin family, which, together with 2 thromboxanes and leukotrienes, is a class of lipid mediators of inflammation derived from arachidonic acid. The synthesis of prostaglandins from arachidonic acid is dependent on the activity of enzymes known as cycloxygenases (Levi et al., 1998). POE 2 signaling takes place via binding to a family of G-protein-coupled receptors (including EP1, EP2, EP3, and EP4) distributed widely throughout the body (reviewed by Sugimoto & Narumiya, 2007). Although POE 2 can readily cross the BBB (Eguchi et al., 1988), its level in the normal CNS remains low.  However, under pathological conditions such as MS, ischemia, HIV  associated dementia, and trauma, the CNS becomes a source of POE 2 (Farooqui & Horrocks, 1991; Fretland, 1992; Griffin et al., 1994; Suganami et al., 2003). Various functions have been attributed to POE 2 in the brain, ranging from CNS damage to neuroprotection (Cazevieille et al., 1994; Chen & Bazan, 2005; Engblom, D., et al., 2002; Théry et a!., 1994). Several reports have demonstrated the involvement of POE 2 in the process of DC maturation (Feuerstein et al., 2000; Soruri & Zwirner, 2005). It has been found that mature monocyte-derived DCs’ migration capacity in vitro in response to CCR7 ligands is mediated by 2 (Scandella et a!., 2004). Furthermore, a role for POE POE 2 has been described in the in-vivo migration of mouse Langerhans cells (Kabashima et al., 2003). This molecule has also been used in the development of a “maturation cocktail” for the generation of mature monocyte derived DCs for experimental and clinical use (Feuerstein et al., 2000; Thurner et a!., 1999).  1.5.5. Role of DCs in CNS Pathology In the normal CNS, there are only a few DCs in the meninges and the choroid plexus, and none within the brain parenchyma. Since the initial observation of DCs in CNS inflammation by  32  Matyszak & Perry, several studies have pointed to the involvement of DCs in the pathogenesis of CNS inflammatory conditions such as MS and EAE, the neurodegenerative disease amyotrophic lateral sclerosis (ALS), and animal models of infection and stroke (Henkel et al., 2004; Huang et al., 2006; Karman et al., 2004b; Kostulas et a!., 2002; Matyszak & Perry, 1996; Stichel et al., 2006; Weir et al., 2002; also reviewed by McMahon et a!., 2006, and Pashenkov et al., 2003). In infection or autoimmune inflammation, DCs appear at sites of immune response in the CNS. Increased numbers of DCs secreting pro-inflammatory cytokines have been reported in the blood of MS patients (Huang et al, 1999) and in their CNS (Serafini et a!., 2006). In addition, both myeloid and plasmacytoid DCs are present in the cerebrospinal fluid (CSF) of patients with MS and neuroborreliosis (Pashenkov et al., 2001). In animal studies, EAE can be induced by the transfer of DCs pulsed with encephalitogenic peptides into naïve mice (Karman et at, 2004a; Weir et al., 2002). One study has found that mature Ag-pulsed DCs injected into the mouse brain traffic to draining cervical lymph nodes and induce preferential recruitment of Ag-specific T cells to the brain (Karman et a!., 2004b). Furthermore, in acute EAE only perivascular cuffs of DCs are seen, whereas in chronic EAE, DCs are distributed perivascularly as well as diffusely throughout the brain (Serafini et al., 2000). The continued presence of DCs in chronic and relapsing EAE has suggested that DC recruitment and maturation within the CNS may be important for the initiation and progression of autoimmune CNS inflammation (Serafini et al., 2000). It has subsequently been found that DCs alone are indeed sufficient for in-vivo Ag presentation and for mediating inflammation in EAE (Greter et al., 2005).  Therefore, DC recruitment to the CNS seems an important player in the  initiation and progression of CNS inflammation. In addition to these findings, recent evidence has accumulated in support of a tolerogenic role for CNS DCs (Bi!sborough et al., 2003; Martin et al., 2002; Steinman et a!., 2000). It has  33  been suggested that immunogenic vs. tolerogenic properties of DCs may be a function of different culture conditions, distinct DC lineages, or different maturation states of the same lineage. For example, it has been found that TGF-13 treatment of DCs leads to the induction of tolerance and the amelioration of EAE symptoms (Jin et al., 2000). Similarly, DCs matured in the presence of TNF-a alone induce tolerance and resistance to EAE, whereas DCs fully matured by CD4O ligand or LPS lead to autoimmune inflammation in the CNS (Lutz & Schuler, 2002). The enzyme indoleamine deoxygenase expressed by APCs may also mediate tolerogenic effects by inducing tryptophan degradation and T cell apoptosis (Grohmann et a!., 2003; Munn et al., 2002).  Furthermore, DCs may engage in the direct killing of T cells through Fas-FasL  interactions (Matsue et al., 1999), although this has not yet been documented in the CNS.  1.5.6. The Great Debate: Origin of CNS DCs The origin of brain DCs has long been a matter of debate: different reports have suggested either local generation from precursors such as microglia, or direct recruitment of DCs or DC precursors from the periphery (McMahon et al., 2006). In-vitro studies of neonatal microglia have suggested that microglia are a relatively undifferentiated population, whose developmental pathway could be skewed towards a DC- or a macrophage-like phenotype depending on the cytokine milieu (Fischer & Reichmann, 2001; Santambrogio et al., 2001). The relevance of these findings to the adult CNS in vivo is yet to be documented. Another in-vitro study of migration, this time across peripheral ECs, has shown that monocytes that have already migrated across the endothelium have the capacity to migrate back to the apical side of the ECs and differentiate into DCs (Randolph et al., 1998). Although there is currently no consensus regarding the origin of CNS DCs, recent animal studies strongly suggest a peripheral origin as opposed to differentiation from CNS-resident microglia (Deshpande et al., 2007; Greter et al., 2005; McMahon et a!., 2005). The molecular  34  mechanisms of DC recruitment to the CNS, however, have not yet been defined.  1.5.7. DC Adhesion to and Migration across Endothelial Barriers Immature and mature DCs exhibit different cell surface molecules and different migration patterns (Sallusto et al., 1998; Sozzani et a!., 1998). Recent studies on extracerebral ECs indicate that DC trafficking is regulated by unique receptor-ligand interactions, and a number of CAMs and chemokines have emerged as important regulators of this process (de la Rosa et al., 2003; Geijtenbeek et a!., 2000b; Hagnerud et al., 2006; Penna et a!., 2002; Walker et al., 2006). For instance, in an in-vitro study, de la Rosa and colleagues have found PECAM-1 to support the adhesion and migration of DCs across resting and activated HUVEC, and 131 and 132 integrins to mediate adhesion only to resting HUVEC (de la Rosa et al., 2003). Adhesion and migration of DCs across peripheral ECs has also been found to be regulated by ICAM-2 interactions with its high-affinity ligand DC-SIGN (Geijtenbeek et al., 2000b). In the context of DC trafficking to the CNS, it has been found that deficiency in the chemokine receptor CCR2 or its ligand (CCL2, also known as monocyte chemotactic protein-i, or MCP-1) prevented mononuclear cell infiltration and disease pathogenesis in the EAE mouse model (Huang et al., 2001a; Izikson et a!., 2000). DC infiltration of the CNS is also prominent following ischemia, but the role of these cells and the mechanism of their recruitment is unknown (Fischer & Reichmann, 2001). Zozulya et a!. have recently described a role for the chemokine CCL3 (a.k.a. macrophage inflammatory protein, or MIP- 1 a) in the transmigration of mouse DCs across mouse cerebral ECs (Zozulya et al., 2007). Furthermore, an EAE study has suggested that CCL2O and its ligand CCR6 may be involved in the recruitment of DCs to the CNS during autoimmune inflammation (Serafini et al., 2000). The role of eCAMs in the trafficking of human DCs to the CNS, as well  35  as the influence of the phenotypic and functional differences between immature and mature DCs on their differential trafficking across the BBB have not been previously examined.  1.6.  OBJECTIVES AND SPECIFIC AIMS  1.6.1. Objectives and Hypotheses The entry of leukocytes into the CNS is under strict regulations by the BBB. DCs, the most potent Ag presenting cells in the immune system, are not present in the normal CNS, but have recently been implicated in several CNS diseases including MS. The exact role of DCs in CNS pathological processes, the extent of their participation in neuroinflammation, and the mechanisms regulating their trafficking to the CNS are currently not well defined. Immature and mature DCs display differences in phenotype and function in vivo, and they likely utilize distinct mechanisms for CNS trafficking during inflammation. Thus, the overall objective of this study is to test the hypotheses that 1) distinct sets of eCAM/ligand interactions regulate the differential trafficking of immature and mature DCs to the CNS under inflammatory conditions, and 2) DCs are active participants in CNS inflammation.  1.6.2. Specific Aims This study intends to address the following specific aims: 1. To characterize the expression of eCAM ligands in immature and mature human monocyte derived DCs, and to determine the role of eCAMs and their ligands in the adhesion of immature and mature DCs to HBMECs. a. This aim will be addressed through in-vitro experiments, using a well-established invitro model of the human BBB consisting of primary cultures of HBMECs, as well as DCs generated in vitro from human peripheral blood monocytes.  36  b. The expression of eCAM ligands by immature and mature DCs will be analyzed by FACS analysis.  The number of cells expressing each molecule as well as the  intensity of expression on a single cell will be graphed as histograms. The percentage of positive cells will also be recorded as an indicator of expression. The role of eCAMs and their ligands in the adhesion of DCs to HBMECs will be determined by in-vitro adhesion assays and blocking experiments.  For these experiments, the  dependent variable will be the number of DCs adhering to HBMEC cultures following the adhesion assay. These numbers will be determined following staining for light microscopy.  The independent variables will include HBMEC condition  (resting vs. activated), DC subset (immature vs. mature), duration of the adhesion assay (15 mm., 30 mm. or 60 mm.), and blocking condition. 2. To determine the extent of DC participation in CNS pathology in situ. a. Standard indirect immunoperoxidase staining techniques will be used to determine the expression of immature and mature DC markers in several normal and pathological brain sections. b. The independent variable is the condition examined, and the dependent variable is the . Data from all diseases are compared to normal brains. 2 number of DCs per mm  37  CHAPTER 2: MATERIALS AND METHODS 2.1.  ENDOTHELIAL CELL CULTURES  2.1.1  Isolation of Human Brain Microvessel Endothelial Cells (HBMEC) HBMEC were isolated from normal human cerebral cortex obtained at autopsy less than  1 8h post mortem. Several primary cultures were utilized from different autopsy brains, with the approval of the University of British Columbia and Vancouver Hospital ethics committees. Following the removal of the meninges, the cerebral cortex was cut into 1-2 mm cubes and incubated in 0.5% dispase (Life Technologies Inc.) for 3h in a 37 °C shaking water bath. The digested tissue was centrifuged at 1000 x g for 10 minutes.  After the removal of the  supernatants, the pellets were suspended and centrifuged again in 15% Dextran (Sigma, St. Louis, MO) for 10 minutes at 5800 x g.  This was followed by incubation in 0.1%  collagenase/dispase (Roche Diagnostics, Laval, QC) in a 37 °C shaking water bath in order to remove the pericytes and basement membrane components.  The cells were then washed,  suspended in Medium 199 (Ml 99, Gibco/Invitrogen) with 5% horse serum (Cocalico Biologicals Inc., Reamstown, PA), layered over 45% Percoll gradients (Sigma) and centrifuged at 1000 x g for 10 minutes in order to separate the HBMEC from the remaining basement membrane, pericytes, and erythrocytes.  Subsequently, the layer containing HBMEC was aspirated and washed in  Ml 99 with 10% horse serum. The isolated small clumps of HBMEC were then suspended in culture media (see section 2.1.2.) and plated onto fibronectin- or collagen (Sigma)-coated tissue culture-treated flat bottom polystyrene plates (Corning Life Sciences, Corning, NY). 2.1.2  Culture Conditions HBMEC were grown on fibronectin-coated 96 well plates in growth media containing  Ml 99 supplemented with 10% plasma-derived horse serum (Cocalico Biologicals), 100 ig/m1  38  heparin (Sigma), 20 ig/ml endothelial growth supplement (Sigma), 1% antibiotic/antimycotic solution (Invitrogen Canada, Burlington, ON) and 300 jig/mi glutamine (Sigma). Cultures were placed in a 37 °C humidified incubator with 5% CO . The culture media were changed every 2 other day. The cultures reached confluence after 7-10 days. The purity and endothelial nature of the cells was established by positive staining for Factor VIJI-related antigen and Ulex Europeaus I lectin binding, as described previously (Dorovini Zis et al., 1991).  2.2.  DENDRITIC CELL GENERATION  2.2.1. Monocyte Isolation Peripheral blood obtained from healthy volunteers was drawn in heparin-coated tubes, and was immediately treated with Ethylenediaminetetraacetic acid (EDTA, Sigma) upon removal from the tubes. Whole blood was incubated with 50 jil/ml of a human monocyte enrichment cocktail (RosetteSep, StemCell Technologies, Vancouver, BC) for 20 minutes at room temperature, in order to crosslink unwanted mononuclear cells (such as lymphocytes) with erythrocytes. Following the incubation period, the blood was diluted with phosphate buffered saline (PBS) containing 1% fetal bovine serum (GIBCO/Invitrogen) and 0.01% EDTA in a 1:1 ratio. Diluted blood was layered over Ficoll gradients (Histopaque 1077, Sigma) and centrifuged at 400 x g for 30 minutes. All mononuclear cells (except monocytes) were gathered at the bottom of the tubes along with erythrocytes. The opaque white layer containing the negatively selected monocytes was aspirated and washed with PBS. The cells were then centrifuged at 250 x g for 10 minutes. Following the removal of supernatants, the pellet was washed in PBS and centrifuged again for 10 minutes (at 250 x g) in order to further remove impurities, such as platelets and plasma proteins. The pellets then underwent erythrocyte lysis (Human Erythrocyte Lysing Kit, R&D Systems, Minneapolis, MN) to eliminate the remaining erythrocytes.  39  Following another 10-minute centrifugation in the wash buffer provided in the Lysing Kit, the pellets were suspended in RPMI 1640 (Invitrogen) and either stained for FACS analysis (see section 2.3.1) or cultured for the generation of dendritic cells (see section 2.2.2.). Viability was 99% by the trypan blue exclusion test. Purity was >90%, as assessed by CD14 staining. 2.2.2. In-vitro Generation of Immature and Mature DCs In order to obtain immature DCs, freshly isolated monocytes were cultured in complete RPMI media (containing 10% human AB serum, 1% antibiotics, and 1% giutamine) in different wells of a 12-well plate (Corning Life Sciences) in the presence of GM-CSF (Peprotech, Ottawa, ON) and IL-4 (StemCell Technologies), each at a concentration of 100 ng/mi (1000 U/mi). The cells were fed by replacing half of the culture supernatants every other day.  Generation of  mature DCs was carried out by treating immature DCs on day 6 with a cocktail containing TNF a. (Sigma) at 10 ng/ml (1000 U/mi), IL-i f3 (Inter Medico, Markham, ON) at 1000 U/ml, IL-6 (Peprotech) at 100 ng/mi (1000 U/mi), and PGE 2 at 0.35 tg!ml (Cayman Chemical, Ann Arbor, MI).  This cocktail has previously been shown to induce the maturation process of DCs  (Feuerstein et ai., 2000). Thus, on day 7 there were two distinct populations of DCs in culture (immature and mature) as determined by flow cytometry. Viability was assessed by 7-AminoActinomycin D (7-AAD) exclusion during FACS analysis (BD Biosciences, San Diego, CA).  2.3.  FACS ANALYSIS  2.3.1. Cell Surface Staining Expression of cell surface molecules was determined in monocytes as well as in seven day-old immature and mature DC populations by staining live cells with primary and Phycoerythrin (PE)-conjugated secondary Abs at 4 °C. Cells were suspended in RPMI 1640 and were then treated with the appropriate quantities of primary Abs or isotype-matched controls (see  40  section 2.4.) for 20-45 minutes inside 5 ml polystyrene round-bottom tubes (BD Biosciences) in an icebox.  The fluorochrome-conjugated primary Abs were not incubated at this stage.  Following incubation, 500jil of a wash buffer containing PBS with 2% FCS and 0.01% NaN 3 was added to each tube, and the tubes were centrifuged at 1500 R.P.M. for 10 minutes at 4 °C. The supernatants were discarded by decanting and the tubes were left in an upside-down position for 1 minute. In order to remove all remaining unbound primary Abs, the cells were centrifuged again in another 500pi of the wash buffer for 10 minutes. Cells were incubated with 100 il/ml of the secondary Ab (or with fluorochrome-conjugated primary Abs) for 20 minutes in the dark at 4 °C (7-AAD incubation time was 10 minutes only).  Subsequently, another 10-minute  centrifugation at 1500 R.P.M. was performed in the wash buffer, followed by fixation in 300 pi of 1% paraformaldehyde. The tubes were capped and stored in the dark at 4 °C. 2.3.2. Data Collection and Analysis FACS analysis was carried out using a FACSort cytometer (BD Biosciences). Data were acquired using the CellQuest Pro software (BD Biosciences) and were analyzed by FCS Express (De Novo, Thornhill, ON). Results are shown as histograms, as generated by FCS Express.  2.4.  ANTIBODIES  2.4.1. Flow Cytometry Antibodies The following mouse primary antibodies (Abs) were used for flow cytometry: anti human DC-SIGN (CD209, clone 120507, IgG2b) at 5 jig/mI, anti-human CD1 ic (p150 a chain, clone 3.9, IgGi) at 10 jig/mi, anti-human CD4O (clone M3, IgGi) at 10 jig/ml, and anti-human CCR7 (clone 150503, IgG a) at 10 jig/mi (all from R&D Systems), anti-human VLA-4 (CD49d, 2 clone HP2/1, IgG ) at 2 jig/mi and anti-human CD83 (clone HB15a, IgG2b) at 2 jig/mi 1 (Immunotech, Marseille, France), anti-human PECAM-1 (CD31, clone JC7OA, IgGi) at 20 jil/ml 41  and anti-human MHC class II (HLA-DP, -DQ, and —DR, clone CR3143, IgGi) at 17.5 jig/mi (Dako, Denmark), anti-human CD11a (LFA-1 u chain, clone Hull, IgGi) at 5 jig/mi, antihuman CD1a (clone H1149, IgGi) at 5 jig/mi, anti-human s-Le’ (CD15s, clone CSLEX1, 1gM) at 5 jig/mI, anti-human B7.1 (CD8O, clone L307.4, IgG ) at 5 jig/mi, anti-human B7.2 (CD86, 1  clone FUN-i, IgG ) at 5 jig/mi, and fluorochrome-conjugated anti-human CD45/CD14 (clones 1 2D1 and MgP9 respectively, IgGi and IgG2b respectively) at 20 jiu/ml (all from BD Biosciences), and anti-human 132 integrin (CD 18, clone 1B4, IgG a, Calbiochem/Cedarlane). Isotype-matched 2 controls included PE-conjugated mouse IgG 1 (CLCMG1O4), PE-conjugated mouse IgG a 2 (CLCMG2AO4), and PE-conjugated mouse IgG2b (CLCMG2BO4), all at 5 jig/ml (Cedarlane, Hornby, ON).  PE-conjugated goat F(ab’) 2 anti-mouse IgGi (CLCC35004) was used as a  secondary Ab at 10 jig/mi (Cedarlane). 2.4.2. Antibodies for Adhesion Assays and Blocking Studies Following an adhesion assay, DCs were stained for CD45 (leukocyte common antigen, clones 2B1 1 and PD7/26, Dako) at 3.5 jig/ml. HRP-conjugated goat anti-mouse IgG (115-035003) was used as the secondary Ab at 4 jig/ml (Jackson ImmunoResearch, West Grove, PA). For the functional blocking studies the following primary mouse Abs were employed against various eCAMs or their ligands: anti-human ICAM-l (CD54, clone RR1/1, IgG , 1 BioSource/Invitrogen), anti-human ICAM-2 (CD 102, clone CBRIC2/2, IgG a, Serotec/ 2 Cedariane), anti-human VCAM-1 (CD1O6, clone 1G11, IgG , Immunotech), anti-human 1 PECAM-i (CD3 1, clone hec7, gift from Dr. W.A. Muller), anti-human E-selectin (CD62E, clone C126C1OB7, IgG a, BioSource/Invitrogen), as well as anti-human CDI8, DC-SIGN, VLA-4, and 2 s-Le’ (as described in section 2.4.1 .), all at concentrations of 20 jig/mi. This concentration was supraoptimal as previously determined by ELISA. Isotype-matched controls consisted of mouse  42  anti-human CD1a (clone H1149, IgG , BD Biosciences) at 20 jig/mI, mouse anti-human CD3 1 (clone HIT3a, IgG a, BD Biosciences) at 20 jig/mi, and mouse IgG2b control (MOPC 141, 2 Sigma) at 20 jig/mi. All blocking Ab preparations were free of NaN 3 and other preservatives. 2.4.3. Antibodies for In-Situ Study The following mouse primary Abs were used on paraffin-embedded sections to identify DC participation in CNS pathology in situ: anti-human DC-SIGN (as described in section 2.4.1.) at 5 jig/mi, anti-human fascin (p55, clone 55K-2, IgG , Dako) at 0.67 jig/ml, anti-human CD4O 1 (clone 11E9, IgG2b, Lab Vision Corp., Fremont, CA) at 33 jiu/ml, anti-human MHC class II (HLA-DP, -DQ, and —DR; as described in section 2.4.1) at 0.425 jig/mi. An HRP-conjugated goat anti-mouse IgG ([H+Lj, 115-035-003) was used as the secondary Ab at 1.6 jig/mi (Jackson ImmunoResearch).  2.5.  ADHESION ASSAY AND IMMUNOCYTOCHEMISTRY Confluent HBMEC monolayers were used untreated or following incubation with 100  U/mi of TNF-cL for 24 h to optimally upregulate the expression of ICAM- 1 and VCAM- 1 (Wong et al., 1999). In separate experiments, monolayers were treated with TNF-cL for 5 h to optimally upregulate E-selectin expression (Wong & Dorovini-Zis, 1996b). Following a wash with M199, resting or activated monolayers were incubated with immature or mature DC suspensions (5 x DCs/well) for 15, 30, or 60 minutes at 37 °C. At the end of the incubation period, nonadherent DCs were aspirated, each well was washed gently four times with M199 and once with PBS to remove the remaining non-adherent DCs, and the monolayers with adherent DCs were fixed with 5% paraformaldehyde for 15-20 minutes. Quantification of DC adhesion was performed by staining the adherent DCs for CD45 with the indirect immunoperoxidase technique as previously reported (Dorovini-Zis et al. 1992). 43  Briefly, monolayers with adherent DCs were washed with PBS and endogenous peroxidase activity was blocked with 0.75% H 0 in 100% methanol (50 jil/well in a 96-well plate) for 30 2 minutes. Cells were then incubated with the anti-CD45 Ab for 1 hour, washed twice with PBS (for 3-5 mm.), and treated with the secondary Ab for 1 hour. Subsequent to washing with PBS and 2 ddH O , cultures were incubated with 0.05% 3,3’-diaminobenzidine (DAB, Sigma) for 30 minutes, followed by haematoxylin counter-staining. Resting HBMEC monolayers served as negative controls. Adhesion was quantified by counting the adherent DCs in one central and 4 peripheral fields in each well with an ocular grid (area: 0.25 mm ) using a 20x objective of a 2 Nikon Labophot light microscope (Nikon Canada, Mississauga, ON). The number of adherent DCs per mm 2 was determined by calculating the mean of five counts and multiplying the result by four. All experiments were run in triplicate wells and all counts were carried out blindly.  2.6.  ENZYME-LINKED IMMUNOSORBENT ASSAY (ELISA) To measure the surface expression of ICAM-2 in our in-vitro model of the BBB,  HBMEC grown to confluence in triplicate wells of 96-well plates were used in a resting state or following activation with 100 U/ml of TNF-cL for 12h, 24h, or 48h.  Following cytokine  activation, monolayers were fixed in 0.025% glutaraldehyde and incubated with a mouse primary Ab against human ICAM-2 at three different dilutions (5 ig/ml, 10 jig/mi or 20 ig/m1) for 60 minutes at room temperature. Monolayers were then incubated with HRP-conjugated goat anti mouse IgG for 60 minutes.  0-phenylenediamine (OPD) was added at 2 mg/mI for colour  development and the reaction was terminated with 2M sulphuric acid. measured on a plate reader at 492nm.  Absorbance was  Monolayers incubated with carrier buffer instead of  primary Ab served as negative controls, and monolayers incubated with a primary Ab against human ICAM-1 were used as positive controls.  44  2.7.  BLOCKING STUDIES AND IMMUNOPEROXIDASE STAINING For blocking studies, HBMEC monolayers were incubated with blocking Abs against  eCAMs 30 minutes prior to incubation with DCs. In separate experiments, DCs were treated with blocking Abs against the ligands of these eCAMs for 30 minutes prior to the adhesion assay. In additional experiments, Abs against HBMEC adhesion molecules and their ligands on DCs were employed simultaneously. The adhesion assay was performed as described above. Immunocytochemistry was performed using the Animal Research Kit, Peroxidase, for Mouse Primary Antibodies (Dako), to prevent the binding of secondary Ab to adhesion molecules and their ligands. In short, endogenous peroxidase activity was blocked in each well by applying 1-2 drops of the peroxidase block provided in the kit (0.03% H 0 containing NaN 2 ) 3 for 5-6 minutes. Following 2 washes in PBS, cells were incubated with a biotinylated primary Ab solution for 15-20 minutes.  The solution was made just prior to the experiment and  contained 1% anti-human CD45 (at a final concentration of 3.5 tg/ml), 3.5% Biotinylation Reagent (provided in kit, containing modified biotinylated anti-mouse immunoglobulin in Tris HC1 buffer, stabilizing protein and 0.015 mol/L 3 NaN ) , 4% Blocking Reagent (provided in kit, consisting of normal goat serum in Tris-HC1 buffer, stabilizing protein and 0.015 mol/L NaN ), 3 all dissolved in a diluent (PBS with 1% Bovine Serum Albumin (Sigma)). At the end of the incubation period, cells were washed with PBS for 5 minutes twice before treatment with HRP conjugated Streptavidin (provided in kit; 1-2 drops/well in a 96-well plate). DAB (provided in kit) was utilized as the visualization substrate and was incubated with the cells for approximately 5 minutes.  Cells were then washed in ddH O and counter-stained with haematoxylin. DC 2  adhesion was quantified as described in section 2.5.  45  2.8.  DCs IN CNS PATHOLOGY  2.8.1. Patients This study was performed on archived, formalin-fixed and paraffin-embedded brain or spinal cord tissue from 76 patients with various pathological conditions obtained with ethical approval from the Department of Pathology & Laboratory Medicine, Vancouver General Hospital. These cases (which comprised 145 tissue blocks) included 9 MS cases (acute or chronic), 4 cases of cerebral ischemia (3 infarcts and 1 anoxic encephalopathy), 6 cases of vasculitis, 3 cases of vasculitis with cerebral amyloid angiopathy (CAA), 28 infectious disease cases (10 cases of bacterial meningitis, 7 cases of viral infection, 4 cases of fungal infection, 1 case of toxoplasmosis, 2 cases of malaria, and 4 cases of brain abscess), 2 cases of chronic granulomatous inflammation (1 tuberculosis (TB) and I sarcoidosis), 9 cases of neuro degenerative disease (3 cases of amyotrophic lateral sclerosis (ALS), 3 cases of Alzheimer’s disease (AD), and 3 cases of AD with CAA), 1 case of head injury, 1 case of perivenous encephalomyelitis, 1 case of acute rejection following pulmonary transplantation, 1 case of paraneoplastic syndrome, and 11 cases of primary or metastatic tumours (See Table 1). Sections from 6 patients with no CNS pathology (mean age: 54 years) were used as controls. Sections of tonsils or lymph nodes from other unaffected patients were also stained with the appropriate markers and served as positive controls. 2.8.2. Immunohistochemistry im-thick) were incubated at 37 °C overnight and heated to 60 °C for Paraffin sections (3 1 30 minutes. In order to deparaffinize the tissue, sections underwent three 5-minute washes in xylene, followed by two 5-minute washes in 100% ethanol, and one 5-minute wash with 95% ethanol. The tissue was rehydrated immediately by placing in ddH O. 2  46  For antigen retrieval, ddH O was heated in a steamer until the temperature reached 952 100 °C. For DC-SIGN, Fascin, and MHC class II staining, sections were placed in a container with 10 mM citric acid-phosphate buffer (pH 6.0). For CD4O staining, the sections were placed in 1 mlvi Tris-acetate-EDTA buffer (pH 8.0) instead. The buffer (with the sections) was heated to boiling in a microwave for 1 minute, and the solution was replenished and heated for another minute.  The container was then placed inside the steamer and heated for approximately 30  minutes. It was ensured that the slides were covered in buffer throughout the heating period. The container was then removed from the steamer and allowed to cool at room temperature for 20 mm. Sections subsequently underwent three 2-minute washes in Tris-Tween buffer (pH 7.6). In order to block endogenous peroxidase activity, the sections were incubated with 0.5% 0 in 100% methanol for 30 minutes. Following three 5-minute washes in Tris-Tween buffer, 2 H the slides were incubated with Tris-Tween with 5% normal goat serum (Gibco/Invitrogen) for I hour in order to block non-specific secondary Ab binding. Sections were then incubated with the appropriate primary Abs diluted in Tris-Tween buffer. For DC-SIGN, Fascin, and MHC class II staining, sections were incubated with their respective primary Abs for 90 minutes at room temperature, whereas for CD4O staining, incubation with primary Ab was carried out overnight at 4 °C. For Ab control purposes, some sections were treated with Tris-Tween containing no primary Ab.  After three 5-minute washes in Tris-Tween, sections were incubated with the  secondary Ab for 90 minutes. Sections were then washed three times (5 mm. each) in Tris Tween buffer and then washed in acetate buffer (with Tween, pH 5.2).  Each section was  subsequently incubated with approximately 200 il of the chromogen 3-amino, 9 ethyl-carbazole (AEC) for approximately 10 minutes. AEC solution (pH 5.25) was made by diluting AEC powder (Sigma) in dimethylformamide to a concentration of 8 mg/mi and further diluting this in  47  50 mM acetate buffer to a working concentration of 0.4 mg/mi. Following the AEC incubation and three 5-minute washes with water, sections were stained with Carazy’ s haematoxylin for 3-4 minutes.  The slides were then washed thoroughly with ddH O, incubated with base for 45 2  seconds and washed again with H . Two drops of Crystal Mount (ESBE Scientific, Markham, 0 2 ON) were then added to each slide.  Slides were left to dry at room temperature overnight,  mounted with Entellan and coverslipped. DC participation in CNS pathology was quantified by counting positive-stained cells in ten random fields on each slide with an ocular grid (area: 0.25  2)  using a 20x objective of a  Nikon Labophot light microscope (Nikon Canada). The number of DCs per mm 2 of CNS tissue was determined by calculating the mean of the ten counts and multiplying the result by four.  2.9.  STATISTICAL ANALYSIS All statistical analyses were performed using GraphPad Prism 4 (GraphPad Software, San  Diego, CA).  Analysis of variance (ANOVA) was performed on all data, followed by  Bonferroni’s multiple comparison tests to examine the differences between individual treatments. Comparisons between two groups were carried out by Student’s t-tests. Two-factor ANOVA was performed in studies of immature and mature DC adhesion to resting and activated HBMEC. In blocking studies, one-way ANOVA was employed to compare the adhesion of each DC subtype in different blocking conditions.  Similarly, one-way ANOVA followed by  Bonferroni’s post test was utilized for the analysis of the in-situ data. P values of smaller than 0.05 were considered statistically significant in all experiments. All bar graphs were generated by GraphPad Prism 4 (GraphPad Software).  48  Table 1:  Patient Data  Disease Category  Mean  Subtype  Age*  # of  # of  Cases  Blocks  Multiple Sclerosis (MS)  52  Acute  1  1  Chronic  8  14  Cerebral Ischemia  59  Acute Infarct  2  2  Chronic Infarct  1  6  Anoxic encephalopathy  1  2  Vasculitis without cerebral_amyloid_angiopathy_(CAA)  6  12  Vasculitis with CAA  3  5  Vasculitis  62  Bacterial Meningitis  47  10  17  Brain Abscess  55  4  4  Viral Infections  47  Herpes simplex virus (HSV) Encephalitis  1  2  Varicella zoster virus (VZV) Encephalitis  1  1  HTLV- 1 associated myelopathy/ tropical spastic_paraparesis_(HAM/TSP)  1  2  Progressive Multifocal Leukoencephalopathy_(PML)  1  1  Other Viral Infections  3  3  Aspergillus Meningoencephalitis  3  5  Cryptococcus infection  1  1  Toxoplasmosis  1  4  Malaria  2  2  Tuberculosis (TB) Sarcoidosis  1 1  1  Fungal Infections  Protozoa! Infection  55 45  Chronic Granulomatous Inflammation  30  Trauma  67  1  2  Perivenous Encephalomyelitis Transplant Rejection  45  1  2  60  Lung Transplant  1  1  Neurodegenerative Disease  53 75  Amyotrophic Lateral Sclerosis (ALS)  3  9  Alzheimer’s Disease (AD)  3  6  ADwithCAA  3  5  Glioblastoma multiforme (GBM) Gliomatosis Cerebri  1  3  1  2  Fibrillary Astrocytoma  1  1  Oligodendroglioma  2  2  Metastatic Carcinoma  5 1 1  8  Tumours  55  Lymphoma Paraneoplastic Syndrome  56  1  *The age was not included for 11 cases because they were forensic cases with sealed envelopes.  49  1 1  CHAPTER 3: RESULTS 3.1. HUMAN BRAIN MICROVESSEL ENDOTHELIAL CELLS Primary cultures of HBMEC form confluent monolayers (Fig. 3a) after 7-10 days in culture. Strong perinuclear granular staining for Factor VIII (von Willebrand Factor) (Fig. 3b) and Ulex Europaeus agglutinin (Fig. 3c) confirm the endothelial nature of these cells. Previous studies from our laboratory have demonstrated that this in-vitro model of the BBB retains important morphological and functional attributes of its in-vivo counterpart, including the presence of tight junctions (Fig. 3d) and the paucity of pinocytic vesicles (Dorovini-Zis et al., 1991). The endothelial tight junctions, which contribute to the high trans-endothelial electrical resistance of this in-vitro model, are also responsible for restricting the passage of horseradish peroxidase across the monolayers (Dorovini-Zis et al., 1991).  3.2. SURFACE PHENOTYPE OF IN VITRO-GENERATED DCs 3.2.1. Characterization of Immature and Mature DCs FACS analysis confirmed that peripheral blood monocytes differentiated into DCs following a 7-day treatment with GM-CSF and IL-4 in culture. This is illustrated by the dramatic upregulation of DC-SIGN and the disappearance of the monocyte marker CD14 from the culture (Fig 4a). While only 2.87% of the monocyte population expressed DC-SIGN, over 95% of immature DCs expressed this molecule. DC-SIGN expression slightly decreased upon maturation, as shown by a slightly smaller percentage of DC-SIGN-positive cells and a lower mean fluorescence intensity (MFI). On the other hand, CD 14, which was absent from the 7-day culture of both immature and mature DCs, was highly expressed in freshly-isolated monocytes. Furthermore, DCs expressed CD1 ic at high levels, but this molecule is also expressed on some other cells of myeloid lineage, such as monocytes and macrophages (Mazzone & Ricevuti,  50  1995). This molecule was also detected at high levels in freshly-isolated monocytes (Fig. 4a). Of all co-stimulatory molecules tested (i.e. CD8O, CD86, CD4O, CD83), only CD83 was absent from monocytes. CD8O and CD4O displayed intermediate expression (in approximately 49% of cells), whereas CD86 was detected at high levels in monocytes (approximately 90%). Upon differentiation into DCs, CD86 remained highly expressed. Likewise, CD4O and CD83 were only slightly upregulated. CD8O, on the other hand, was downregulated, and it was only expressed in approximately 5% of the 7-day immature DC population. All co-stimulatory molecules, however, displayed dramatic upregulation upon DC maturation (see Fig. 4a). CCR7, a chemokine receptor involved in mature DC migration to secondary lymphoid organs, exhibited intermediate expression in our mature DCs (in approximately 62% of cells). CD 1 a, which according to some studies is a marker for immature myeloid DCs and Langerhans cells, was found to be minimally expressed in both monocytes and DCs. Finally, the antigenpresenting molecule MHC class II, which was expressed highly in monocytes, was upregulated upon differentiation into DCs, and was further upregulated upon maturation (Fig. 4a, last panel).  3.3.2. Expression of eCAM Ligands by Immature and Mature DCs Immature and mature DCs exhibited different expression profiles for the ligands of eCAMs. As shown in Fig. 4b, both immature and mature DCs expressed DC-SIGN (ligand for ICAM-2), however, immature DCs displayed a slightly higher expression. Both the c’. and 13 chains of LFA- 1 (CD 11 a and CD 18) were expressed highly in immature and mature DCs, with little observable difference between the two cell types. In contrast, both DC subtypes expressed sLex (ligand for P- and E-selectin) minimally. PECAM-1 (ligand for PECAM-1 on ECs) was expressed highly in both immature and mature DCs, with a higher expression in immature DCs. On the other hand, VLA-4 (ligand for VCAM- 1) only displayed intermediate level of expression  51  in both cell types, with a somewhat higher expression in immature DCs (see. Fig. 4b).  3.3. ADHESION OF IMMATURE AND MATURE DCs TO HBMEC 3.3.1. Adhesion to Resting and Activated HBMEC Following a 60-minute adhesion assay, adhesion of both immature and mature DCs to resting HBMEC was very low (1% and less than 1% respectively, Fig. 5a). Although the adhesion of immature DCs to resting HBMEC was slightly higher compared to mature DCs, this difference was not statistically significant (p> 0.05). A 24-h treatment of HBMEC with TNF-c, a potent inflammatory cytokine, significantly upregulated adhesion of both immature DCs (p  <  0.001) and mature DCs (p <0.01), as determined by two-factor ANOVA and Bonferroni post tests (Fig. 5a). Immature DCs displayed a greater increase in adhesion, and significantly greater adhesion to activated HBMEC compared to mature DCs (4% and 1% respectively, p <0.001). A similar pattern in adhesion was observed following a 5-hour activation with TNF-cL. The 5-hour TNF-c treatment was carried out for the purpose of optimally upregulating the expression of E-selectin on HBMEC (Wong & Dorovini-Zis, 1 996b). Similar to the 24-hour activation, a 5-hour treatment with TNF-a also led to a significant upregulation in adhesion of both immature DCs (p  <  0.00 1) and mature DCs (p  <  0.05), as determined by two-factor  ANOVA followed by Bonferroni tests (Fig. 5b). Immature DCs displayed significantly greater adhesion to activated HBMEC compared to mature DCs (3% and 1% respectively, p <0.001).  3.4.2. DC Adhesion Change with Time Adhesion of both immature and mature DCs to resting and activated HBMEC was found to be time dependent (Fig. Sc). Immature DC adhesion to resting HBMEC increased significantly when the duration of the adhesion assay was increased from 15 mm to 30 mm (p  52  <  0.01) and  from 30 mm to 60 mm (p <0.001). Mature DC adhesion to resting HBMEC was significantly upregulated only after the duration of the adhesion assay was increased from 30 mm to 60 mm (p <0.01). Adhesion to activated HBMEC followed the same pattern. Immature DC adhesion was significantly upregulated when adhesion assay duration changed from 15 mm to 30 mm (p<O.05) and from 30 mm to 60 mm (p < 0.00 1). On the other hand, mature DC adhesion was increased significantly only after the adhesion assay duration changed from 30 mm to 60 mm (p <0.01). The difference between immature and mature DC adhesion to activated HBMEC was not significant after a 15-minute adhesion assay (p < 0.05, as determined by ANOVA). This difference only became significant when the adhesion assay lasted for 30 minutes (p <0.01) or 60 minutes (p  0.00 1), as determined by ANOVA and Bonferroni post tests.  3.4. SURFACE EXPRESSION OF ICAM-2 BY HBMEC Surface expression of ICAM-2 was determined by ELISA in confluent HBMEC monolayers before and after treatment with TNF-c for 12, 24, and 48 h. Using different concentrations of the anti-ICAM-2 Ab (5—20 ig/ml), a constitutive expression of ICAM-2 was found in resting HBMEC monolayers (Fig. 6). Treatment of HBMEC with TNF-c (100 U/ml) for 12-48 h led to slight downregulation in ICAM-2 expression, but statistically significant decreases in ICAM-2 expression were only observed at 48 h (Fig. 6). Incubation of HBMEC cultures with carrier buffer resulted in no staining, whereas treatment with anti-ICAM- 1 led to strong absorbance.  3.5. REGULATION OF DC ADHESION TO HBMEC BY eCAMS AND THEIR LIGANDS 3.5.1. DC Adhesion to Resting HBMEC Applying blocking Abs against eCAMs and their ligands on DCs showed that the minimal adhesion of immature and mature DCs to resting HBMEC was not downregulated  53  significantly by treating HBMEC with blocking Abs against ICAM-1, ICAM-2, PECAM-1, VCAM-1, or E-selectin (Figures 7a and 7b). Likewise, treating immature and mature DCs with blocking Abs against the respective ligands of these eCAMs (i.e. CD 18, DC-SIGN, PECAM-1, VLA-4 or sLex) did not lead to a significant downregulation of the binding of either immature or mature DCs to resting HBMEC (Figures 8a and 8b).  3.5.2. DC Adhesion to Activated HBMEC Monoclonal Ab blocking of eCAMs and/or their ligands on DCs had a significant effect on adhesion to activated HBMEC. Immature DC adhesion to activated HBMEC was significantly downregulated upon treating HBMEC with Abs against ICAM-1, ICAM-2, PECAM-1 and VCAM-l (p  <  0.001), but not E-selectin (Figures 9a and 9b). Furthermore,  immature DC adhesion to activated HBMEC was significantly downregulated upon blocking DC ligands CD18 (a.k.a. 132 integrin; p  <  0.05), DC-SIGN (p  <  0.001), PECAM-1 (p <0.01), but not  VLA-4 or s-Le’ (Figures lOa and lob). Treating HBMEC or immature DCs with isotype matched control Abs had no effect on adhesion (Figures 9b and lob). Adhesion of mature DCs to activated HBMEC was also decreased significantly in the presence of blocking Abs against ICAM- 1. Blocking of PECAM- 1, VCAM- 1, or E-selectin had no effect on adhesion (p<0.OOl, Figures 1 Oa and lob). Blocking CD 18, DC-SIGN and PECAM 1 (but not VLA-4 or sLe)c) on mature DCs also led to a significant decrease in adhesion (p< 0.001 , Figures ha & llb). Similar to the observation with immature DCs, treating HBMEC or mature DCs with isotype-matched control Abs had no effect on adhesion (see Figs. lib & 12b). In separate experiments where blocking Abs were applied against both eCAMs and their ligands on DCs, adhesion of immature DCs was significantly downregulated by blocking ICAM 1 and CD18 (p <0.05), ICAM-2 and DC-SIGN (p <0.001), PECAM-1 and PECAM-h (p  54  <  0.001), as well as VCAM-1 and VLA-4 (p <0.001, Figures 13a and 13b). Blocking E-selectin and its ligand sLex did not lead to a significant downregulation in immature DC adhesion to activated HBMEC (Fig. 13). In addition, combining all blocking Abs in the same culture led to a significant decrease in immature DC adhesion, to 33% as compared to adhesion in the absence of blocking Abs (p <0.001, Fig. 13b). Likewise, adhesion of mature DCs to activated HBMEC was significantly downregulated by blocking ICAM-1 and CD 18, ICAM-2 and DC-SIGN, and PECAM-1 and PECAM-1 (p <0.05, Figures 14a and 14b), but not by blocking E-selectin and s Lex. In contrast to what was observed in immature DCs, blocking VCAM- 1 -VLA-4 interactions did not lead to a significant decrease in mature DC adhesion to activated HBMEC (Figures 14a and 14b). Furthermore, combining all blocking Abs led to a significant decrease in the adhesion of mature DCs, to 67% of adhesion when no blocking Abs were applied (p<O.O5; Fig. 14b).  3.6. DENDRITIC CELL PARTICIPATION IN THE CNS IMMUNE RESPONSE A summary of the immunohistochemical results is shown in Table 2. The cases of malaria, perivenous encephalomyelitis, trauma, paraneoplastic syndrome, and transplantation were excluded from statistical analysis due to small sample size (n = 1 or 2). There was no significant difference between the ages of patient samples and normal brains (mean age: 54). Most cases stained positive for DC-SIGN, fascin, and MHC Class II molecules. CD4O staining, however, was only observed in some cases. The positively stained cells were often observed in close association with infiltrating lymphocytes. As positive controls, several lymph node sections were stained for DC-SIGN, fascin, CD4O, and MHC class II molecules. They all displayed ample and diffuse staining.  55  Table 2: DC Participation in CNS Pathology  Disease Category  Subtype  DC-SIGN  Fascin  CD4O  MHC Class_II  MS Cerebral Ischemia  Acute and Chronic Acute and Chronic Infarct Anoxic Encephalopathy  +  -7+ -1+ -7+  -  +  -1+ -1+  +  Vasculitis without CAA Vasculitis with CAA Meningitis Malaria Brain Abscess Viral Infections Fungal Infections Toxoplasmosis TB  +  -7+  -1+ -1+ -7+ -7+ -7+ -7+ -7+ -7-i-  -i--i--i-  ++  +++  Sarcoidosis  +++  ++  +++  +++  -7+  -7+  -7+  -7+  -7-i-7+  -7+ -7+  -  +  -  ++  +  -1+  +  -7+ -7+  -  -7+  ++  +  ++  ++  +  -7+ -7+  +  -7-i-  ++  +  -  +  Vasculitis Infectious Disease  Chronic Granulomatous Inflammation Perivenous Encephalomyelitis Trauma Transplant Rejection Neurodegenerative Disease Tumour (Primary) Tumour (Metastatic)  Lung Transplant ALS AD & AD with CAA Metastatic Carcinoma Lymphoma  + +  + + + +  -7+ +  -  Paraneoplastic Syndrome  -/+  Note: The symbols denote the following expression levels: -:0 -/+: 1-20 cells/mm 2 +: 20-40 cells/mm , 2 ++: 40-60 cells/mm , 2 +++: 60+ cells/mm 2  56  -1+ -7+ -7+ -  + + ++ + -  -1+  +  -/+  ++  -7+  +  -  +  ++  3.6.1. DC-SIGN-Positive DCs In all cases, DCs showed positive cell membrane staining for DC-SIGN. In sections of normal brain a small number of DC-SIGN-positive cells were present in the meninges, the choroid plexus, and in the perivascular space around intraparenchymal blood vessels (see Figure 1 5a). In most brain lesions, the numbers of DCs were moderately to highly increased. Significant upregulation of DC-SIGN cells was observed in MS lesions  (p<O.O5), ischemia  (including cerebral infarcts and anoxic encephalopathy; p<O.OO1), vasculitis (p<O.OO1), CAA ), brain abscesses (p<O.OO1), fungal 5 associated vasculitis (p<O.O5), bacterial meningitis (p<O.O infections (p<O.O 1), toxoplasmosis (p<O.OO 1), chronic granulomatous inflammation of tuberculosis (TB) and sarcoidosis (p<O.OO1), familial ALS (p<O.05), AD (p<O.O5), and tumours (p<O.OO1; see Figures 15(a-q) and 16 for details). In these cases, DC-SIGN cells were most abundant in the perivascular regions of both gray matter and white matter in the lesion areas, in addition to the meninges and choroid plexus. In MS, the infiltration of DC-SIGN cells was observed within the chronic inactive plaques in the white matter (Fig. 1 5b). In bacterial meningitis, DC-SIGN DCs were associated with leukocyte infiltration in the meninges (Fig. 15f). In localized inflammatory processes and tumours, DCs were not only perivascular, but were also seen infiltrating diffusely the lesion area. In these cases, perivascular DC-SIGN cells were also present in the surrounding brain parenchyma and a few in the meninges (Figures 1 5d, 15g, 15(i-k), 15(l-m), 15(p-q)). There were 3 MS cases in which the tissue was obtained by biopsy. These cases were excluded from statistical analysis due to their small size and their unsuitability for selecting ten random fields for quantification. Furthermore, unlike other tumours, the lymphoma case was devoid of DC-SIGN cells and it was therefore excluded from DC-SIGN analysis.  57  3.6.2. Fascin-Positive Cells In all diseases studies, the anti-fascin Ab was positive in a few cells present in the parenchyrnal perivascular spaces, choroid plexus, and meninges, although this Ab stained fewer leukocytes as compared to the anti-DC-SIGN Ab (see Figures 1 7(a-q), and 18 for details). Since fascin is expressed in mature DCs as well as in brain ECs and nerve tissue (Kureishy et a!., 2002; Zhang et al., 2008), the non-DC fascin-positive cells were omitted from the counts based on morphology. Similar to the DC-SIGN staining, only a few fascin cells were present in perivascular spaces, choroid plexus, and leptomeninges of normal brains (see Fig. 17a). The numbers remained low for most CNS diseases. However, a statistically significant upregulation of fascin cells was observed in cases of vasculitis (p<O.Ol), meningitis (p<O.O5), toxoplasmosis  (p<O.O5), chronic granulomatous inflammation (p<O.OOl) and tumours (p<O.OO1; see Fig.  18 for  details). Furthermore, one of the three cases of Aspergillus infection studied here contained numerous fascin cells. Similar to DC-SIGN cells, fascin cells were mostly observed in the meninges and the perivascular areas of the cortex.  3.6.3. CD4O-Positive Cells CD4O-positive cells were not detected in sections of normal brain (Fig. 19a). Furthermore, CD40 cells were identified in some but not all brain lesions. There were no CD40 cells in MS lesions, ischemia, toxoplasmosis, or familial ALS (see Figs. 1 9b, 1 9c, 19k, and 1 9n). A small number of CD40 cells was detected in vasculitis, CAA, meningitis, abscess, viral and fungal infections, AD, and tumours (see Figs. 1 9(d-e), 1 9(f-j), and 1 9(o-q)). Interestingly, the only disease category in which there was a statistically significant expression of CD40 cells was chronic granulornatous inflammation, with similar expression levels of CD4O in tuberculosis and sarcoidosis (p<0.OOl for both; see figures 19(1-rn) and 20 for details).  58  3.6.4. MHC Class Il-Positive Cells Similar to DC-SIGN and fascin, there was a clear MHC class II expression in most sections. There were very few MHC class IT-positive cells in the normal brain, but the numbers were highly upregulated in most pathological cases (See figures 21 (a-q)). Cases of MS  (p<O.O5),  ischemia (p<O.OO1), vasculitis (p<O.OO1), CAA (p<O.OO1), meningitis (p<O.OO ), abscess 1 (p<O.OO1), viral infections (p<O.OO1), fungal infections (p<O.Ol), toxoplasmosis (p<O.OO 1), chronic granulomatous inflammation (p<O.OO1), familial ALS (p<O.OO1), and tumours (p<O.OO1), all displayed significantly greater than normal expression of MHC class II molecules. Except for the cases of AD, AD with CAA, and meningitis, MHC class II positive cells were distributed in the brain parenchyma in addition to perivascular and meningeal sites. In this study, the MHC class Il-positive cells which were morphologically distinguishable from DCs based on light microscopy (i.e. microglial and endothelial cells) were excluded from counts. However, there is another category of MHC class Il-positive cells (such as B lymphocytes and macrophages) which is not easily differentiated from DCs based on light microscopy alone and may have been included in this analysis (Perry, 1998; Traugott, 1987).  3.6.5. Immature vs. Mature DC Participation in CNS Pathology Determining the exact participation of immature and mature DCs was made difficult due to the lack of availability of an anti-CD83 primary Ab suitable for paraffin sections. However, since all sections displayed DC-SIGN and fascin expression, and since CD4O and MHC class II are also expressed in other leukocytes, fascin-staining was taken as the best available marker of mature DCs in CNS pathology. Due to the high expression of DC-SIGN in both immature and mature DCs (see Fig. 4a and 4b), immature DC participation was determined by subtracting the number of fascin-positive cells from DC-SIGN-positive cells (Fig. 23a). Based on this  59  calculation, the only cases which displayed significantly greater-than-normal participation of immature DCs were ischemia, vasculitis, and abscess. A comparison of the relative numbers of immature vs. mature DCs in all cases is shown in Fig. 23b. In chronic granulomatous inflammation, ALS, and tumours, there were significantly more mature DCs compared to immature DCs. On the other hand, in MS, ischemia, vasculitis, brain abscess, fungal infections, and AD, there were significantly greater numbers of immature DCs. In CAA, meningitis, toxoplasmosis, viral infections, as well as in normal brains, there was no statistically significant difference between immature vs. mature DC presence in the CNS.  CHAPTER 4: DISCUSSION 4.1. HBMECs AS A MODEL OF THE BBB The entry of circulating leukocytes into the CNS across the BBB is a hallmark of many neurological disorders. Since the ECs of the cerebral blood vessels are the first class of cells that interact with leukocytes, studying leukocyte-EC interactions is of great significance in understanding the pathophysiology of CNS diseases. In-vitro models of the BBB allow investigators to study brain EC-leukocyte interactions without the presence of the in-vivo confounding variables. Our laboratory has thus developed an in-vitro model of the human BBB consisting of primary cultures of HBMECs, which retain important morphological and functional characteristics of the human BBB in vivo (Dorovini-Zis eta!., 1991). These include expression of Factor VIIIR:Ag (von Willebrand factor), binding of the Ulex europaeus lectin, presence of tight junctional complexes between adjacent ECs restricting the paracellular movement of molecules, paucity of cytoplasmic vesicles and absence of a vesicular transport system. This in  60  vitro system has been used reproducibly to study the responses of cerebral ECs to cytokine activation and their role in leukocyte trafficking across the BBB in CNS inflammation (Huynh & Dorovini-Zis, 1993; Quandt & Dorovini-Zis, 2004; Wong & Dorovini-Zis, 1995, 1996a, 1996b). Recently, several studies have devised and utilized models in which leukocyte-EC interactions are examined under conditions of flow. These studies report differences in the kinetics of leukocyte trafficking between “static” and “dynamic flow” models. However, several of these studies utilize extracerebral large vessel ECs such as aortic ECs (Cucullo et al., 2002; Santaguida et al., 2006), which are different from brain microvascular ECs in phenotype, function, and immunological properties. Furthermore, one of the primary concerns driving the use of flow-based models is the association made between the absence of hemodynamic forces and EC apoptosis (Kaiser et al., 1999). However, this occurs as a result of prolonged cultivation. It is therefore believed that despite the absence of flow, our present model is a suitable in-vitro system for the study of leukocyte trafficking, not only because the cultures are used at an optimal time (after 7-10 days), but also because our HBMEC are derived from the cerebral microvascular bed where the blood flow is slow and there is close contact between circulating leukocytes and ECs. Furthermore, during the inflammatory response, vasodilation leads to relative stagnation of blood flow and a reduction in the speed of leukocyte movement. Thus, circulating leukocytes have ample opportunity to interact with, roll along, and adhere to ECs.  4.2. CHARACTERIZATION OF IMMATURE AND MATURE DCs  Myeloid DCs are considered the most potent antigen presenting cells of the immune system (Jiang et al., 2005, Miller et al., 2007). Thus, their recruitment to and activation in the CNS are likely key events in neuroinflammation, as suggested by studies in MS and its animal model EAE, ALS, and animal models of stroke and infectious disease (Greter et al., 2005; Karman et al., 2004a;  61  McMahon et a!., 2006; Pashenkov et al., 2003, Ponomarev et al., 2005). The mechanisms of DC trafficking to the CNS are presently not well understood. In this study we successfully generated and characterized monocyte-derived DCs, which are widely regarded and used as a good in-vitro model for inflammatory myeloid DCs (reviewed by Shortman & Naik, 2007). Our DCs express high levels of DC-SIGN, CD11c, CD86, and MHC class II molecules, as do their in-vivo counterparts. Upon maturation, these cells undergo dramatic upregulation in the expression of co-stimulatory molecules (i.e. CD8O, CD86, CD4O, and CD83), the homing chemokine receptor CCR7, and the antigen presenting MHC class II molecules. These changes are consistent with previous studies of DCs and correspond to DC function as antigen presenting cells (Feuerstein et al., 2000, Hsieh et al., 2001; McMahon et a!., 2006; Reis e Sousa, 2006). Our immature DCs do not express the lipid antigenpresenting molecule CD1a, although this molecule is commonly observed in immature myeloid DCs, especially in Langerhans cells, the specialized skin DCs (Huang et a!., 2001b; La Rocca et al., 2004). There are also studies that point to the existence of CD1 a-negative immature monocyte-derived DCs (Caux et al., 1997; Gogolak et al., 2007), a population bearing greater resemblance to the DCs generated in this study. Since these differences in CD 1 a expression are likely related to different culture conditions, CD1a may not be considered a universal marker for immature mye!oid DCs (Gogolak et al., 2007). It is also interesting that in their study of MS patients, Serafini et al. could only detect CD1a cells in the CNS tissue of one patient, who had been diagnosed with secondary progressive MS, whereas DC-SIGN DCs were observed in all MS cases (Serafini et a!., 2006). CCR7, a chemokine receptor expressed in mature DCs, exhibited intermediate expression in our mature DCs. Other reports show minimal to intermediate expression of CCR7 in mature human DCs using various maturation protocols (Csomor et al., 2007; Desai et al., 2007; Li et al., 2007; Milano et al., 2007; Sordi et al., 2006). Specifically, one study shows that 65% of mature  62  monocyte-derived DCs express CCR7 following treatment with LPS (Li et al., 2007), which is similar to the percentage of CCR7 cells documented here. CCR7 is the receptor for CCL 19 and CCL2 1 (two chemokines expressed by high endothelial venules in lymphoid organs), and has been found to be important for mature DC migration to secondary lymphoid organs via the high endothelial venules (Dieu et al., 1998; Forster et al., 1999; Martin-Fontecha et al., 2003; Ohi et al., 2004; Saeki et al., 1999; and Willimann et al., 1998). Since these two lymphoid chemokines have been shown to be expressed by brain ECs in EAE and since CCL 19 levels were found to be increased in MS, CCR7 may play a role in DC migration across the BBB (Columba-Cabezas et al., 2003; Krumbholz et al., 2007). However, another study has reported the lack of CCL19 and CCL2 1 from cerebral ECs in MS lesions (Kivisäkk et al., 2004). Thus, the exact role of CCR7 in DC migration in vivo across the BBB in CNS pathological conditions remains to be defined. The surface expression of eCAM ligands in DCs was also examined in this study. Immature DCs displayed a higher expression of the C-type lectin DC-SIGN (ligand for ICAM-2), the immunoglobulin PECAM- 1 (CD3 1, ligand for endothelial PECAM- 1), and the integrin VLA-4 (CD49d, ligand for VCAM-1 and extracellular matrix components) as compared to their mature counterparts. Other adhesion molecules such as the 132 integrin LFA-1 (ligand for ICAM-1 and ICAM-2) and the carbohydrate sLex (CD 1 5s, ligand for E- and P-selectin), were expressed at very similar levels in immature and mature DCs. Whereas both the cc and f3 chains of LFA-l were highly expressed by both DC subtypes, the expression level of sLex was very low. A study on human peripheral blood DCs reported CCR7 and CD1 la expression levels in immature myeloid DCs that were similar to what was found in our experiments (de la Rosa et al., 2003). However, compared to the cells employed in this report, our immature DCs displayed a lower expression of VLA-4 (by approximately 20%) and a higher expression of PECAM-1 (by around  63  40%). It is interesting that DC-SIGN was not detected in immature DCs in the above study, whereas this molecule has been identified at high levels in both immature and mature DCs and has been implicated in the process of DC trafficking (Geijtenbeek et al., 2000a, 2000b; Soilleux et aL, 2002). This may be due to differences in DC generation techniques, for although our immature DCs are related to the circulating peripheral blood DCs, they are derived from monocytes and they represent DCs in inflammatory, not homeostatic conditions (Shortman and Naik, 2007; Wu and Liu, 2007). It is also important to note that there are several ways to generate mature DCs. The present method, which uses a combination of TNF-cL, IL-i , IL-6, and PGE 2 (i.e., components of monocyte conditioned media), has been shown to be more effective than LPS, poly I:C, CD4O ligand, and some other cytokine combinations in yielding large numbers of stable mature DCs (Feuerstein et al., 2000). Furthemore, when compared to TNF-cL alone and TNF-a with PGE , the present cocktail leads to 2 greater yield, more pure mature phenotype, and better T cell stimulation (Thurner et a!., 1999). Since this maturation stimulus leads to the activation of multiple receptors involved in the inflammatory response, it likely reflects DC maturation in a variety of pathological conditions in vivo.  4.3. DC ADHESION TO HBMEC Leukocyte trafficking across endothelial barriers is a multi-step process involving rolling, activation, adhesion and migration (see Chapter 1). Our study focused on the process of DC adhesion to brain ECs under resting and inflammatory conditions, and as such it is the first study that documents in detail some of the molecules involved in this process. It was observed that this process followed a time-dependent course, from minimal adhesion at 15 minutes to higher levels at 60 minutes for both resting and activated HBMEC. However, when ECs were in a resting state, only a small fraction of DCs adhered to the monolayers even following a 60-minute adhesion assay (1—3% of immature DCs and 0.4—2% of mature DCs). A recent study on DC adhesion to resting HUVEC  64  reports much greater adhesion, even with a short, 5-minute adhesion assay: 80% for immature DCs and 65% for mature DCs (Jiang et al., 2005). In our study, activating HBMEC with TNF-ct. for 24 hours led to a significant upregulation in adhesion: a 3.5 fold increase for immature DCs and a 2.7 fold increase for mature DCs, on average. Another study of DC adhesion to activated extra-cerebral ECs indicates a similar upregulation in adhesion to activated ECs (D’Amico et al., 1998). However, a direct comparison may not be appropriate due to fundamental differences between various DC generation methods, differences between cell lines (utilized in that study) and primary cultures (used in our work), and differences between large vessel and microvessel ECs. Previous studies from our laboratory also indicate significant upregulations in the trafficking of other leukocytes across TNF-a activated HBMEC cultures. For instance, the adhesion of PMNs has been found to undergo a 4.4  —  6.8 fold increase following EC activation with TNF-a (Wong et al., 2007) and the adhesion of various T cell subsets has displayed increases of up to 4-fold (Quandt and Dorovini-Zis, 2004). Our results also demonstrate differences between immature and mature DC adhesion to activated HBMEC. Following 60-minute adhesion assays, the adhesion of immature DCs to resting HBMEC was 36%  —  150% greater than that of mature DCs, but this difference did not amount to  statistical significance. On the other hand, the adhesion of immature DCs to activated HBMEC was significantly greater than that of mature DCs (by 104%  —  192%). These findings are consistent with  the characteristics and function of DCs in vivo, as most circulating blood DCs are immature DCs and this is presumably the subtype that traffics to various tissues via crossing the endothelial lining of blood vessels in order to perform its innate immune functions (Wu and Liu, 2007). Our results are also supported by previous studies to some extent. According to one report, immature DC adhesion to resting HUVEC is 23% greater than that of mature DCs (Jiang et al., 2005). One possibility is that this difference between immature and mature DC adhesion to activated  65  HBMEC is due to a greater interaction between immature DCs and those molecules that are upregulated on HBMEC as a result of TNF-cL treatment. Previous work from our laboratory has shown that PECAM- 1 is constitutively expressed in resting as well as activated HBMEC monolayers, whereas ICAM-l, VCAM-l and E-selectin levels undergo significant upregulation following TNF-CL treatment (Wong & Dorovini-Zis, 1992; 1995; 1996a; 1996b). Furthermore, the present study shows a constitutive expression of ICAM-2 in our in-vitro model of the BBB, similar to what had been found in extracerebral EC cultures and in brain microvessels in vivo (de Fougerolles et al., 1991; Navratil et al., 1997). Therefore, the interactions of these eCAMs with their ligands on DCs are important elements in the differential adhesion of immature and mature DCs to primary cultures of HBMEC. The differences found in the present study between immature and mature DCs in the expression of the ligands for eCAMs correspond to this trend in adhesion. Interestingly, however, one study which has also reported a maturation-related decrease in DC adhesion to fibronectin and ICAM-1-coated surfaces has found no difference in the expression of CD1 la, VLA-4, and other integrins between immature and mature DCs, and has explained the adhesion pattern based on morphological changes and cytoskeletal rearrangements that take place upon DC maturation (Bums et al., 2004). Since these cytoskeletal changes are an integral part of DC maturation, they may play a significant role in the adhesive and migratory properties of DCs.  4.4.REGULATION OF DC ADHESION TO HBMEC BY ECAMS AND THEIR LIGANDS In order to further investigate the roles of various eCAMs and their ligands in the process of DC adhesion to the BBB, individual molecules were blocked using monoclonal blocking Abs. Our results indicate that the low baseline adhesion of immature and mature DCs to resting HBMEC is not dependent upon the interaction of ICAM- 1, ICAM-2, VCAM- 1, PECAM- 1 and E-selectin with their  66  cognate ligands. Similarly, previous studies from this laboratory have established that the adhesion of PMNs to resting HBMEC is unaffected by blocking ICAM-1, VCAM-l, PECAM-l and E selectin (Wong et al. 2007), and the adhesion of T lymphocytes to resting HBMEC is only affected by blocking ICAM-1, and not by blocking VCAM-1, PECAM-1 or E-selectin (Wong et at, 1999). We have also demonstrated that the adhesion of immature and mature DCs to activated HBMEC undergoes significant downregulation by blocking eCAMs and their ligands. Immature DC adhesion to activated HBMEC is downregulated upon blocking ICAM-l (by 40%), ICAM-2 (by 40%), VCAM-1 (by 62%), and PECAM-1 (by 88%) on HBMEC, and upon blocking CD18 (by 37%), DC-SIGN (by 51%) and PECAM-1 (by 23%) on DCs. Furthermore, when blocking both sides of the interaction in the same adhesion assay, immature DC adhesion to activated HBMEC is downregulated upon blocking ICAM-1 and CD1 8 (by 27%), ICAM-2 and DC-SIGN (by 52%), PECAM-1 and PECAM-1 (by 45%) and VCAM-1 and VLA-4 (by 62%). On the other hand, the adhesion of mature DCs to activated HBMEC is significantly downregulated only when blocking ICAM-1 on HBMEC (by 65%), and CD18, DC-SIGN and PECAM-1 on DCs (by 46%, 54%, and 39% respectively). When blocking molecules on both cell types, mature DC adhesion to activated HBMEC is downregulated upon blocking ICAM- 1 and CD 18 (by 31%), ICAM-2 and DC-SIGN (by 36%), PECAM-1 and PECAI\4-1 (by 33%) but not VCAM-1 and VLA-4. When all of the above molecules are blocked in the same adhesion assay, both immature and mature DCs undergo significant downregulation but not complete inhibition in adhesion to activated HBMEC (by 67% and 33% respectively), which implies the involvement of additional molecules in the process. In the case of DC-SIGN blocking, it is possible that the partial blocking of adhesion is due to the binding of ICAM-2 to CD18, although it has been found that ICAM-2 binds CD18 with lower affinity compared to DC-SIGN (Bleijs et al., 2001). It is also interesting that the ICAM-1/LFA-1 and  67  ICAM-2/DC-SIGN interactions have both been found to resist conditions of shear stress, whereas the ICAM-2/LFA-l interaction has been found incapable of doing so (Bleijs et at, 1999; Geijtenbeek et a!., 2000; Sigal et at, 2000). These fmdings, coupled with the constitutive expression of ICAM-2 in ECs, have led investigators to postulate that ICAM-2/DC-SIGN interactions precede ICAM-1ILFA-1 interactions in the process of DC trafficking across endothelial barriers (Bleijs et al., 2001). The partial blocking of DC adhesion to HBMEC observed in this study may also be explained by the potential involvement of other molecules in this process, such as chemokines and other adhesion-related molecules which were not addressed in this study. Indeed, one study has recently reported the involvement of the chemokine MIP- 1 a and matrix metalloproteinases in the process of murine DC transmigration across murine brain ECs (Zozulya et al., 2007). Whether these molecules are also involved in the adhesion step remains to be investigated. The role of eCAMs in the trafficking of other leukocytes across activated HBMEC has also been previously examined in our laboratory. It has been found that the migration of resting T cells across activated cerebral ECs is dependent on ICAM- 1, and to a less extent on VCAM- 1, PECAM-1 and E-selectin (Wong et a!., 1999). Alternatively, blocking ICAM-1 and E-Selectin significantly downregulates PMN adhesion to HBMEC (Wong et al., 2007). It is interesting that the Eselectin!sLex interaction does not seem to play a major role in DC adhesion to cerebral ECs, whereas E-selectin was found to mediate the adhesion of PMNs to HBMEC (Wong et a!., 2007). Previous studies on the adhesion of DCs to HUVEC have also suggested a role for eCAMs and their ligands in the DC adhesion process. One in-vitro study has suggested that PECAM-1 supports the adhesion and migration of peripheral blood myeloid DCs across resting and activated HUVEC, whereas 131 and f32 integrins mediate adhesion only to resting HUVEC (de la Rosa et a!., 2003). Furthermore, while in our study DC-SIGN and CD 18 seem to be important  68  eCAM ligands in the regulation of DC adhesion, one study of mouse DC trafficking across resting brain ECs has excluded a role for their murine homologues while finding their receptor ICAM-2 to be an important component in DC migration (Wethmar et al., 2006). This latter report, however, may not be directly compared to the present study, for there are well-documented differences in origin, phenotype and function between mouse and human DC subsets (Shortman & Naik, 2007). Overall, our findings suggest that under inflammatory conditions, the adhesion of immature DCs to the BBB is dependent upon ICAM-l—CD18 or ICAM-2—CD18, ICAM-2—DC-SIGN, PECAM-l—PECAM-l, and VCAM-l—VLA-4 interactions, whereas the adhesion of mature DCs to activated HBMEC is mediated by ICAM-l—CD18 or ICAM-2—CD18, ICAM-2—DC-SIGN, and PECAN/I- 1—PECAM- 1 interactions (Fig. 2). It is also possible that other Abs recognizing different epitopes from what was employed here are capable of blocking different intermolecular interactions.  Figure 2: Summary of eCAM-Ligand Interactions in DC Adhesion to HBMEC  4.5. PARTICIPATION OF DCs IN CNS PATHOLOGY The participation of DCs in various CNS diseases has not been studied extensively in humans. This study examined the presence of DCs in a wide spectrum of CNS diseases using an  69  indirect immunoperoxidase technique. This is the first documentation of DC participation in several pathological conditions in humans, namely ischemia, various inflammatory and infectious diseases, neurodegenerative diseases, and some primary and metastatic tumours. In this study, DCs with strong surface staining for DC-SIGN, a specific DC marker, were present in most cases examined. Specifically, DC-SIGN DCs were observed in and at the edges of acute and chronic MS plaques, primary CNS vasculitis, vasculitis associated with CAA, bacterial meningitis, brain abscess, fungal encephalitis, toxoplasmosis, chronic granulomatous inflammation (tuberculosis and sarcoidosis), familial ALS, AD, as well as primary and metastatic tumours. DC-SIGN expressing DCs were most numerous in chronic granulomatous inflammation of TB and sarcoidosis. The other marker used for the analysis of DCs in our study was fascin. Fascin is expressed in mature DCs, as well as in ECs, neurons, and glial cells (Zhang et al. 2008). The numbers of perivascular fascin cells were found to be significantly increased in the brains of patients with vasculitis, meningitis, toxoplasmosis, chronic granulomatous inflammation and tumours. Since the fascin cells were selectively counted based on morphology, they represent an estimate of mature DC presence in the CNS. Although in most CNS cases studied here (namely MS, ischemia, abscess, viral encephalitis, flingal encephalitis, ALS, and AD) the upregulation of fascin was not statistically significant, the number of fascin cells was still somewhat greater than that observed in the normal CNS, which may be of some biological importance. Although sections with abscess seemed to contain a greater number of fascin cells compared to meningitis cases, their numbers don’t reach statistical significance, probably due to their small sample size. Overall, the number of fascin cells was found to be smaller than the number of DC-SIGN cells in all cases. This finding was not unexpected, since DC-SIGN is  70  expressed in both immature and mature DCs and is a characteristic marker of most DCs. CD4O, which is considered a maturation marker for DCs was not readily expressed in our sections. Since sections of lymph nodes used as positive control displayed considerable staining for CD4O, the paucity of CD4O staining cannot be attributed to the antibody’s low affinity. Although CD4O cells were present in small numbers in AD, ischemia, vasculitis, meningitis, and tumours, and even in somewhat greater numbers in CAA-associated vasculitis, brain abscess, and viral and fungal encephalitis, their numbers were not statistically significant. In contrast, increased numbers of CD4O cells were identified in the densely cellular chronic granulomatous inflammation of TB and sarcoidosis. The numbers of these CD4O cells were even larger than those of the fascin cells in this particular disease category. As CD4O has been documented in all APC populations, i.e. microglia, macrophages, and B cells, in addition to mature DCs, this finding may reflect the participation of other cell types in TB and sarcoidosis (Alderson et al., 1993; Stamenkovic et al., 1989; Van Kooten and Banchereau, 1997). Our results demonstrate a significant upregulation of MHC class II in almost all pathological conditions in comparison to normal CNS. The only disease category not displaying a significant upregulation in MHC class II expression was AD with and without CAA. However, even in this disease category, the numbers of MHC class 1I cells were somewhat higher than those found in normal brain sections, which may indicate some biological significance. Indeed, activated microglia (which are MHC class II) are believed to be involved in AD pathogenesis (reviewed by Zlokovic et al., 2008). MHC class II molecules are expressed in great numbers in all APCs, and this expression accounts for their strong staining in this experiment. Hence, the high overall intensity of stain for MHC class II in our cases was consistent with our expectations. The greater staining for MHC class II in comparison to CD4O probably reflects the  71  greater expression of MHC class II molecules in APC populations compared to CD4O expression. This is also supported by our flow cytometry data. It should also be added that MHC class II staining was seen not only in the meninges, perivascular areas, and inflammatory lesions, but also throughout the CNS parenchyma in most cases. Many of these class II MHC expressing cells displayed the elongated and ramified morphology of microglia, and were excluded from statistical analysis. The disease categories where MHC class II staining was restricted to the lesions were MS, ischemia, and meningitis. Another interesting observation in this study was the prominence of mature DCs in chronic and granulomatous inflammatory conditions. This may occur as a result of DC function as APCs and potential propagators of chronic inflammation. Furthermore, it is possible that the chronic inflammatory conditions lead to the creation of microenvironments that are rich in factors stimulating DC maturation. In contrast to MHC class 1I cells, DC-SIGN and fascin cells were closely associated with infiltrating lymphocytes in all cases. For cases such as MS, ischemia, vasculitis, ALS, and AD, DCs were observed in the meninges but more importantly in the perivascular areas within or surrounding the inflammatory lesions. Similarly, in meningitis, DCs were seen almost exclusively in the meninges, where inflammation occurs. However, in certain conditions such as brain abscess, fungal encephalitis, toxoplasmosis, chronic granulomatous inflammation and tumours, DC-SIGN and fascin cells had also infiltrated the brain parenchyma. In many of these latter cases, lesion boundaries were blurred by ubiquitous inflammatory infiltrations. Thus, the in-situ study presented here not only constitutes one of the first comprehensive investigations of DC participation in human CNS diseases, but also suggests an important role for DCs in neuroinflammation and provides support for our in-vitro work regarding the ability of  72  immature and mature DCs to traffic to the CNS. A recent study by Serafini et al. has documented the active participation of DCs in different subsets of MS (2006). This study reports the presence of DC-SIGN cells in all examined brain and spinal cord sections. Similar to the present study, DC-SIGN cells were found in perivascular areas in close association with lymphocytes, but not inside the brain parenchyma. In this report, DCs were found in the early active lesions, on the borders of chronic active lesions, and even in chronic inactive lesions, but not in heavily demyelinated areas. Another in-vivo study has shown that DCs are present in the spinal cord tissue of patients with familial and sporadic ALS (Henkel et al. 2004). In this study, CDla immature DCs were detected in familial ALS, whereas CD83 cells were detected in both familial and sporadic ALS. Interestingly, there were no CDl23 plasmacytoid DCs in the spinal cord sections of ALS patients. Most CD 1 a cells were in perivascular areas, whereas in the ventral horn of the spinal cord, a few parenchymal CDla cells were also observed. CD83 cells remained in the perivascular areas of the spinal cord, and displayed weaker immunostaining compared to CDla DCs. This is not in agreement with the findings of the present study, for although we did not stain our tissue blocks with CD1a or CD83, we detected a significantly larger number of mature DCs (stained for fascin) compared to immature DCs (see section 3.1.5). Furthermore, this study reports an elevation in CD4O mRNA expression compared to controls, but it has not examined the expression of CD4O protein, which renders a direct comparison with our results difficult. One study has also found a significant upregulation in DC infiltration of the CNS in glioblastoma multiforme (GBM). The presence of DCs in GBM (which accounts for more than half of all primary brain tumours) was detected by the expression of both CD1c and CD11c (Hussain et al., 2006). This report is in accordance with our findings, although in our study we  73  examined other types of tumours in addition to GBM. We have nevertheless shown a dramatic and significant upregulation in DC-SIGN and MHC class I1 cells, as well as a statistically significant upregulation of fascin in all tumour cases (except for the one lymphoma case, where there were no DC-SIGN cells). Several in-vivo studies have also demonstrated elevated numbers of DCs in the cerebrospinal fluid (CSF) of patients with various CNS diseases. For example, a significantly greater number of DCs has been documented in the CSF of patients with MS, bacterial meningitis, and Lyme disease, as compared to normal controls (Pashenkov et al., 2001; 2002). DC participation in CNS pathology has been investigated by a number of animal studies as well. For example, in an early experiment, Matyszak & Perry have shown significant DC presence in a delayed-type hypersensitivity reaction to BCG (1996). It has also been shown that DCs drive the initiation and progression of EAE in mice (Karman et a!., 2004a; Miller et al., 2007). Furthermore, DCs have been detected in mouse models of parasitic infection (leishmaniasis and toxoplasmosis) as well as prion disease (Abreu-Silva et al., 2003; Fischer et al., 2000; Rosicarelli et al., 2005). In toxoplasmosis, the CNS DC population has been found to primarily consist of myeloid cells, which is consistent with our detection of DC-SIGN cells in human toxoplasmosis (Fischer et al., 2000). Furthermore, there is evidence for DC participation in mouse and rat models of cerebral ischemia (Kostulas et al., 2002; Reichmann et al., 2002). DCs have also been detected in large numbers in the mSOD1 mouse model of ALS (Henkel et al., 2006), and they have recently been reported to be involved in reducing amyloid plaque formation in a mouse model of AD (Butovsky et al., 2007). Taken together, these studies provide support for our in-situ findings, and further emphasize the role of DCs in CNS pathology.  74  CHAPTER 5: CONCLUSIONS 5.1. SUMMARY AND SIGNIFICANCE At present, the exact immunoregulatory role of DCs in neuroinflammation remains poorly defined. Furthermore, the extent of DCs’ participation in CNS pathology and the mechanisms of their entry into the CNS have not been fully elucidated. As a major step in the process of human DC trafficking across the BBB, DC adhesion to HBMEC is of key importance in the recruitment of DCs from the periphery. In order to study the process of DC adhesion to the brain ECs, in-vitro-generated human monocyte-derived DCs and a well-established in-vitro model of the human BBB have been utilized. DCs were successfully generated from peripheral blood monocytes and it was further demonstrated that monocyte-derived immature and mature DCs display the phenotypic characteristics of their in vivo counterparts. Most relevant to this study is the demonstration of differences in the expression of ligands for eCAMs between immature and mature DCs that prove to be pertinent to the differential adhesion of these DC subsets to the brain endothelium. The adhesion experiments clearly demonstrate that a very small number of DCs adhere to resting brain ECs, which correspond to the BBB under non-inflammatory conditions. Adhesion is highly upregulated upon treatment of ECs with TNF-o to induce an inflanunatory phenotype. Binding of DCs to cerebral ECs follows a time-dependent course for both DC subsets. Furthermore, adhesion to activated ECs is significantly greater for immature DCs in comparison to mature DCs. Blocking studies indicate that DC adhesion to activated brain ECs is partly dependent on interactions between eCAMs and their ligands, and this adhesion process is regulated by distinct eCAM-ligand interactions for the two DC subtypes. Specifically, immature DC adhesion is  75  dependent upon ICAM- 1—CD 18 or ICAM-2—CD 18, ICAM-2—DC-SIGN, PECAM- 1 —PECAM- 1, and VCAM-1—VLA-4 interactions, whereas mature DC adhesion is dependent upon ICAM-1—CD18 or ICAIvI-2—CD 18, ICAM-2—DC-SIGN, and PECAM- 1—PECAM- 1 interactions. The second aim of this study was to investigate the participation of DCs in various human CNS diseases in situ. Based on the great capacity of DCs to perform key functions in both the innate and adaptive immunity, our hypothesis was that DCs actively participate in several pathological conditions in the CNS. This was the first attempt to investigate DC participation for many disease categories. Using immunohistochemistry for DC markers, this study demonstrated that DCs were indeed present in a wide range of human CNS pathologies, including inflammatory diseases (such as MS and vasculitis), ischemia (acute and chronic infarcts and anoxic encephalopathy), infectious diseases (including bacterial meningitis, fungal encephalitis, brain abscess, and toxoplasmosis), chronic granulomatous inflammation (TB and sarcoidosis), neurodegenerative disorders (AD and ALS), as well as CNS tumours. Establishing the active participation of DCs in CNS pathology not only points toward a potentially important role for DCs in the pathogenesis of CNS diseases, but also raises interesting questions regarding Ag presentation in the CNS and emphasizes the importance of investigating the molecular mechanisms of DC trafficking to the CNS under inflammatory conditions. Thus, this study demonstrates that DC infiltration of the CNS is a feature of several diverse diseases affecting the brain and the spinal cord. In addition, our findings indicate that eCAMs and their ligands play an important role in the process of immature and mature human myeloid DC recruitment to the CNS under inflammatory conditions and thus further define the role of cerebral microvascular ECs in regulating the process of inflammation in the human CNS.  76  5.2. FUTURE DIRECTIONS This study leads to several potential lines of investigation, which may further address the issue of DC trafficking to the CNS and clarify the role of cerebral ECs in the initiation and maintenance of the immune response. First, in order to better understand DC trafficking across the BBB, it is necessary to study the process of DC migration across the cerebral endothelial barrier. DC migration across resting vs. cytokine-activated HBMECs will serve as a model of DC migration under physiological versus inflammatory conditions in vivo. Using the same blocking Abs employed in this study, it is possible to shed light upon the role of eCAMs and their ligands in the process of DC migration into the CNS. Secondly, since a large body of evidence suggests that chemokines play an important role in the trafficking of leukocytes into various tissues, and since the brain itself is a source of certain chemokines, a logical step is to study the role of various chemokines in the adhesion and migration of DCs to the brain using blocking Abs. It will be interesting to study DC migration in response to various chemokines in a double-chamber chemotaxis system, and characterize the role of various chemokine/receptor interactions by employing blocking Abs against chemokine receptors on DCs. A third step would be to study the route of DC migration into the CNS. This will involve the blocking of tight junctional molecules, such as claudins, occludin, JAMs, CD99, and PECAM-l. The effect of DC migration on monolayer permeability may also be evaluated by measuring the trans-endothelial electrical resistance before and after the addition of DCs to the EC culture. 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Magnification: (a): lOx; (b) and (c): 20x. Scale bar in (d) = O.5jim  103  Fig.  4a  I::  10 102  Immature DCs  49  96  93 A  Monocytes  ‘: L  io’  0  10 102 io  0  10 102 10’  0  96 9  0  10 102 10’  ° 9 k. 0  io  10  10’ 0  5 91  0  10 10’ 10’  0  10 102 10’  63  5  0  0  10 102 10’  7  0  10 10’ iO’  91  0  10 102 10’  13  2  i oioio i i i i 2 o’oioio’io’oioio’io’oioio’io’oioio o’oioio’io’oioio o°oioio o’oioio o’  Mature DCs  1 88  65 98 96  98  50  Counts o  io 102 jo  0  10  1 io  io  0  10 102 io  0  10 102 i0 3  0  10 102 io 0  10 102 io  93 L 0  10 102 1o  62  0 10 102 io  1  0  10 102 iO  0  10 102 i0 3  II  Fluorescence  Molecule’s Name:  CD14  CD209 (DC-SIGN)  CD11c  CD8O (B7.1)  CD86 (B7.2)  CD4O  CD83  CCR7  CD1a  MHCII  Fhures 4a: Surface phenotype of monocytes, immature DCs and mature DCs as determined by FACS analysis. FACS results are representatives of ten independent experiments. The black curves represent the molecules of interest, and the red curves represent isotype controls. The numbers refer to the percentage of cells expressing each surface molecule. Immature DCs were differentiated from monocytes by culturing monocytes for 7 days in GM-CSF and IL-4 (1000 U/mi) and maturation was induced by culturing immature DCs for 24h in a cocktail of TNF-o (1000 U/mi), IL-113 (1000 U/mi), IL-6 (1000 U/mI) and PGE 2 (0.35 jig/mi).  104  Fig. 4b  Molecule’s Name:  CD209 (DC-SIGN)  CD11a CD18 (132 Integrin, (LFA-1 a chain) LFA-1 13 chain)  CD15s (sLex)  CD31 (PECAM-1)  CD49d (VLA4)  lo:j Immature j 99 0  10  102  3 i0  0  10  102  1O  0  10  4 Lj, 102  3 i0  0  10  102  io  0  10  102  io  0  10  102  i0  100  Mature 1 DC:  88  °:  99  98  i1  _ 10 O.rr  0  ‘o  o  67  o’”io’  42  1O  102  Fluorescence  Molecule’s Receptor:  ICAM-2  ICAM-1, ICAM-2  ICAM-1, ICAM-2  E-Selectin, P-Selectin  PECAM-1  VCAM-1, ECM  Figure 4b: Expression of eCAM ligands by immature and mature monocyte-derived DCs. Results are representatives of seven independent experiments. The red curves represent the isotype controls and the black curves represent the molecules of interest. The numbers refer to the percentage of cells expressing each surface molecule.  105  Resting HBMEC  Activated HBMEC  .  ..  Immature DCs  .,:;;  -:•  :  I ‘  ‘:  4’  :.. .i.,  -,  ..  4  •  .  — .  *  ‘,  Mature DCs  •1 . 4.  —  1-  b  .  *.  “4  •  .0  .  —.  .  •*  4  0 ‘5  s_I  ,  S  •  4) ..  ‘  ER  p.<o.oo: E  I  S  <O.O1  H  Resting  -z  c: immature DCs mature DCs  Activated  HBMEC Condition  Fi2ure 5a: Immature and mature DC adhesion to resting or TNF-cc-activated HBMEC. Cytokine activation time is 24 hours. The micrographs show HBMEC as stained by haematoxylin (elongated blue cells in the background) and DCs as stained with an anti-CD45 Ab (brown globules). Scale bars = lOOjim. Bar graph is representative of seven independent experiments, and illustrates the quantification of DC adhesion to HBMEC by light microscopy. Error bars indicate standard errors of the mean. The asterisks (“*“) denote significant differences in adhesion (p <0.05). 106  * Foi  *  80 Cl  P<O.OO 1  E E I-  a)  immature DCs mature DCs  60.  C) 40  a) .  20.  41:  Resting  Activated  HBMEC Condition  Fiure 5b: Immature and mature DC adhesion to resting or TNF--activated HBMEC. Cytokine activation time is 5 hours. The graph represents the quantification of DC adhesion to HBMEC by light microscopy. Error bars indicate standard errors of the mean. The asterisks (“*“) represent significant differences in adhesion (p < 0.05).  107  Fig. 5c  CJ  E E  —  Immature DC  0.  Activated EC  C) G) G) •D  Mature DC Immature DC Mature DC  .  ‘I  0  15Mm.  30Mm.  —  Resting EC  60 Mm.  Adhesion Assay Duration  Figure 5c: Immature and Mature DC adhesion to resting and 24-hour-activated HBMEC increases with time. The difference between immature and mature DC adhesion to activated HBMEC is statistically significant at 30 minutes (p <0.01) and 60 minutes (p <0.001), but not at 15 minutes, as determined by ANOVA and Bonferroni’s post tests.  108  3.0 2.5 cs1  C)  0  2.0 1.5  *  CU  .0  1.0’  0 U)  .0  0.5’ An, None  12h  24 h  48 h  24h (No Ab)  24h (a-ICAM-1)  TNF-a Incubation Time  Fiure 6: Relative surface expression of ICAM-2 by resting and TNF-cL-activated HBMEC as measured by ELISA. Values represent mean absorbance ± SEM (n = 3). “*“ indicates a statistically significant difference in expression as determined by ANOVA and Bonferroni post tests.  109  7a  E  I  No Blocking  ICAM-1  ICAM-2  PECAM-1  VCAM-1  E-Selectin  VCAM-1  E-Selectin  Blocked Molecule  7b  40  E  E a)  30  0.  C) C  20  a) I..  .  .4-  0 41:  10  III No Blocking  ICAM-1  I  ICAM-2  PECAM-1  Blocked Molecule  Figures 7a and 7b: DC adhesion to resting HBMEC in the presence of blocking Abs against eCAMs. Figure 7a shows immature DC adhesion and figure 7b represents mature DC adhesion. Bar graphs represent quantification following light microscopy. Error bars indicate standard errors of the mean.  110  8a  40 c1  E  E a)  30  0 .1  20  a) a) .  10  LJ  II II F  ‘I  0  No Blocking  CD18  DC-SIGN  PECAM-1  U  VLA-4  s-Le”  Blocked Molecule  8b 40  E I-  a)  30  C) 20 a) .  10  15  [‘I  No Blocking  i’i I CD18  DC-SIGN  PECAM-1  Blocked Molecule  Figures 8a and 8b: DC adhesion to resting HBMEC in the presence of blocking Abs against eCAM ligands. Fig. 8a shows immature DC adhesion and Fig. 8b represents mature DC adhesion. The graphs represent quantification of adhesion following light microscopy. Error bars indicate standard errors of the mean.  111  9a  4  4  —  .4  0  .‘  P  ‘%  i•  I  •1  •  .  4  \  .  *.. 2  S •  4  4  .  4.,  4’  4.,.  4  4  07  .  4  I  7  2  4  4__  .4  .,..  I  4•_  •t  ‘r .  4 4  •  *  -  4;  .  4  0  S  4  I  4  4.  4  ..e. , •—1  I  •  -‘  .5—  *  4  . ,  4.  4.  4 ->  0  -. -  -  4-  •  4 4I  I -5  II  —  •  .**  —  .-  I • .  4 —  ..“  .  0  0.  -  -  0  . .  4.  -  Pl4 53_  •  . ,.,_ 5  -,-  VCAM-1 Blocking  PECAM-1 Blocking  .4•5,  S  I  ..  • •1’  4  I  4  4  S  .  .  4  V 4  4.,  4  4  -  •....  I  .  4  ICAM-2 Blocking  I  -  S  S  —  ICAM-1 Blocking i  .  •  4  .  .‘  .  -  ‘  a  4.  4  —.  No Blocking  4  4  —  4  5  • •4I  s—  I  E-Selectin Blocking  9b 80.  *  C”  E E  I  p<O.OOl  60.  100% 40.  60%  60%  a-ICAM-1  a-ICAM-2  1  20.  I No Blocking  cz-PECAM-1  c-VCAM-1  cc-E-Selectin  lgGl Isotype Control  IgG2a lsotype Control  Ab Treatment  Fkures 9a and 9b: Adhesion of immature DCs to TNF--activated HBMEC in the presence of blocking Abs against eCAMs. Light micrographs in Fig. 9a illustrate HBMEC as stained by haematoxylin (blue background) and DCs as stained by anti-CD45 (brown). Scale bars = 60jim. Bar graphs in Fig. 9b represent quantification following light microscopy. Error bars refer to standard errors of the mean. Percentages refer to adhesion relative to baseline conditions (i.e. “no blocking”, which is 100%). 112  lOa  .  •S. 1 b_  —.  :  ,.._:  a  4  -  :  .  -  4  —  -  ‘A  .-,.-  a  •..  -at  ‘b  ‘.  •  — •.  a  .  -4..  ..‘  •  *  DC-SIGN Blocking ,  a.  .  ‘.  ‘  •  S  a  -:  •.  CD1B Blocking a  .  —  —  .1 —  -  .  . ..  a;.’  ...L•  No Blocking —-  •  S  •_ I  —  a  -  :.  a  9  •  .  a  a  I  ..  -  ‘  a.  :..  a,  9  .  a  ,,  4  •  •4  4-  b  a.  a  A I  • •  •  •  4  •  —  -  4  .  5•  a7 ‘  a  4  -  .•  .‘ .  4  4  4  •  •  .  a  a  S  I  —  • .  •  .•  PECAM-1 Blocking  sLex Blocking  VLA-4 Blocking  *  lOb 80. C,’  E E I  a)  60.  100% 63%  0 C-)  49% 40.  p<.O1  a)  I p<.O01  20. I’  0 No Blocking  a-CD18  a-DC-SIGN  a-PECAM-1  a-VLA4 Control  IgG2a Isotype Control  lgG2b Isotype Control  Ab Treatment  Figures lOa and lOb: Adhesion of immature DCs to TNF--activated HBMEC in the presence of blocking Abs against eCAM ligands. Light micrographs in Fig. lOa illustrate HBMEC as stained by haematoxylin and DC as stained by anti-CD45. Scale bars 60p.m. Bar graphs in Fig. lOb represent quantification following light microscopy. Error bars show standard errors of the mean. Percentages refer to adhesion relative to baseline condition (i.e. “no blocking”, which is 100%). =  113  ha  I  4  4  a *,  ,  4  —  .4  4’  ¶  .  1  4  /  4—.  .4  4  ‘ 4  a  .  —  .4,—..—  No Blocking 4  4  ...  .4—.—— ICAM-1 Blocking  ICAM-2 Blocking  4  .  4  4  .-  -4  -.4  -,  4  4  4  4  &  *4,  .  :  44  (  4  -.,  4  ;.  ‘—  a  4  4 ••  ‘44  0)  4 .-  4*,  -  4  “4  .  4  *  F  —  PECAM-1 Blocking  -  4  .•  VCAM-1 Blocking  —  E-Selectin Blocking  lib N  80  E E a)  0.  60  C.) 4-  *  40  C  a) a) .C  .  I p<O.OO1 20 35%  0 A  No Blocking  cx-ICAM-1  a-ICAM-2  a-PECAM-1  a-VCAM-1  a-E-Selectin  IgGi Isotype Control  IgG2a Isotype Control  Ab Treatment  Figures ha and hib: Adhesion of mature DCs to TNF--activated HBMEC in the presence of blocking Abs against eCAMs. Light micrographs in Fig. 1 la illustrate HBMEC as stained by haematoxylin and DCs as stained by anti-CD45. Scale bars 60iim. Bar graphs in Fig. 1 lb represent quantification following light microscopy. Error bars indicate standard errors of the mean. Percentages refer to adhesion relative to baseline conditions (i.e. “no blocking”, which is 100%). The “*“ indicates a significant difference in adhesion (p < 0.05). =  114  12a —  I  4  •  —.  •  --  I  .  .  I  a  •  S  4  a  .  I S  .  •  •  -•• a.  4--  •  a  ‘  N  a  I  S.  I  S  S  •  S  0•  a•  I  -  N’  .-  .  •.  .  -:  -  -4  S S  •  S  ‘-  “ • a  No Blocking S  *4.  •.•  • ‘  5  :  S ,  ?  I  ,  a  F  5  p  •  4  S  •  S  -  •  a  •  S  I  $  5  b  .  5.  —  4  1  S  5*  4  ..  •  ,  .  DC-SIGN Blocking  CD18 Blocking  ,4  ;_5_  •  a •  •55  4  S  444  5  *  — 5’  •  S.  -4  j  S -  S  PECAM-1 Blocking  sLex Blocking  VLA-4 Blocking  12b 80  *  c.l  60  40  Ip<O.OO1 100% 46%  2: —  No Blocking  I 54%  61%  EE[IJ1_J a-CD18  a-DC-SIGN a-PECAM-1  a-VLA-4  Ab Treatment  Figures 12a and 12b: Adhesion of mature DCs to TNF-a-activated HBMEC in the presence of blocking Abs against eCAM ligands. Light micrographs illustrate HBMEC as stained by haematoxylin and DCs as stained by anti-CD45. Scale bars 6Oiim. Fig. 12b represents quantification of adhesion following light microscopy. Error bars indicate standard errors of the mean. Percentages refer to adhesion relative to baseline conditions (i.e. “no blocking”, which is 100%). The indicates a significant difference in adhesion (p <0.05). =  “*“  115  13a  :t  -Fr  *  a  •%l  .,  ,.(  4  ad  J  ‘  •  4  (  •  -4  -  -  F  (  a  41  r  1  S  — I  a.  .4  •4  .  S  I  .  4 •I..  P  d  •a  I  a  ‘  pS. •.  t 45  4 I  I  No Blocking  ICAM-1/CD18 Blocking  . •  a  0%  s.•  ICAM-210C-SIGN Blocking  .c  •  I.  ,  I .  .  .  9%  .4, 1  .  4  •  . .4 •,.,,A  4  —  4  I  ‘,  .  4  4  -  ‘:  a •  -• •  F  I.  •  ; p  a  •  4  I I  •I  ••  •  1  4  •  0 ___  VCAM-1NLA-4 Blocking 5,  •—s. II  a  I  I  V •  •  •  4  1.  4.  $  I  •  4  .—.  I  •  I  4  ,%4 -1  44  PECAM-1/PECAM-1 Blocking  ‘j  4  ESelectin/sLex Blocking  a  I  •  I  a I..  p  a  4  I  a  -  I—.  All Molecules Blocked  *  13b 1  1  Molecules  Blocked Molecules  Fi2ures 13a and 13b: Adhesion of immature DCs to TNF-cL-activated HBMEC in the presence of blocking Abs against eCAMs and their ligands. Fig. 1 3a illustrates HBMEC as stained by haematoxylin and DCs as stained by anti-CD45. Scale bars 60!Lm. Fig. 13b represents quantification of adhesion by light microscopy. Bars show mean values ± SEM. Percentages refer to adhesion relative to baseline conditions (i.e. “no blocking”, which is 100%). 116  ..  .•\  14a  .  *  -  a  ,  • •  4  ‘  I  .  -  4  •  -p  —  4  .‘  S.  ‘  .,  1  —  ICAM-1ICD18 Blocking  I  ICAM-2/DC-SIGN Blocking  S  1”  :  r p-p..  -‘  p  4._•  .40 •_  .•  4  ‘ •  .4  p  p  P -  —  •  d  a  ••  P  .  a  ,  9 I  4• p  4  4.  p  •  •  •  £•  S  p  %  p  p  ••  4.J  P  •  b  ,  •  -  No Blocking --  4 4  p  1  I,.  b  :  •  p  4  •,  %  St  S  4...  a  .  p  ‘  0  •  ..-..  .  p  •,  ‘  p  —.  .1  .‘‘•.  .  •  ‘-  p  —  / •_‘4  VCAM-1NLA-4 Blocking  P  p  p  .4  -.  PECAM-1/PECAM-1 Blocking  ESelectinIsLex Blocking  ‘4 4  P..  P  ,  *  p.  p  .4. —  All Molecules Blocked  14b  I 1  I  Lj No Blocking  ICAM-1/ CD18  ICAM-21 DC-SIGN  PECAM-1/ PECAM-1  VCAM-1/ VLA-4  E-Selectin/ SLeX  All Molecules  Blocked Molecules  Figures 14a and 14b: Adhesion of mature DCs to TNF--activated HBMEC in the presence blocking Abs against eCAMs and their ligands. Fig. 14a illustrates HBMEC as stained by haematoxylin and DCs as stained by anti-CD45. Scale bars = 60p.m. Fig. 14b represents quantification of adhesion by light microscopy. Bars show mean values ± SEM. Percentages refer to adhesion relative to baseline conditions (i.e. “no blocking”, which is  117  100%).  of  a)  b)  live IffQflic MS Plaque  ‘ -  —  *  -  —  .  •  .•  -4  U—.,  c)  • •  •  •  .  •  ••.•  d)  . ._  .  •.  . .••  __•.••  .  .0  S  • •  .  •.I,  •  . •  •‘  .  4’.  .  •  •  e)  Fhiure 15(a-e): DC-SIGN expression in situ in normal CNS and in CNS inflammatory diseases. Light micrographs show DC-SIGN-positive cells (stained red). Scale bars 6Ojim =  118  •1  g)  1)  •‘.•  .,..  .•  J •  •4  4  •  •  ‘I ••• ‘4  ,•.  4  4., •t, ••  •  •  .  •;  •  4,  ‘‘.•.  •  ‘ci’ 4•,% 9 •  • .•  ‘:  114  ‘ :.‘  •  •  a I  •  •  • •,  I  ‘  a •  *  “  ;•  ‘,a  •..  •l  •:•  —  •  •  • ••.  •.:  :.  ‘.  —  ‘  ‘r’  4’%ê -  i)  VZV Encephalitis  a  I  .  1?  .‘  ii-su1 -.  Aspttrilus In)ection’  4•_•  -g.  •,,,  •‘ .1  *  ‘a  e  S  •  /,.••••.  •j•  .  I-  :  1  I’.  ••  -  ‘a  F,  t  ..•l  .——  ..  4  ,  -I  :  •  .  •i•&  ••  .•‘.  •,,  h)  a.  •: ;%  :4  J  4, *  •4  I  ••  11  ‘4  a  DC-S*LN  ;‘  k)  j) •  •  b  •  •  a  ,_.  .a  1  I.  •  a  •ft  ‘  .  •  4.  •.. .  V.  . 4 p a  : ‘  .•*•  • •  :“ • 4II4 ••, -  4,  a•%.•  .‘  -.— .‘—..  •  .‘  ...  .4  • ‘  -  •  c•  •.  •  V  q  4  ••  •a;.  •  •  •• •  •  •  •  •-  1  •  4’  a a... •  ••  _4 I  •  ..  •  a.  I,  •.  ••  .  •  ,_.•  *  •.4  a.  •  -  .p•  4•_  a. • •• •• a •••• ;•  .•.  ,  a  .  .‘  •• •  • •—.• •• ..  •  a  •.  DC-SIGN  •.•  • •  ( ••  a•  •  4  .4  -•--  •..(  :.  •  S. ;  Figure 15(f-k): DC-SIGN expression in situ in infectious CNS pathologies. Light micrographs show DC-SIGN-positive cells (stained red). Scale bars = 6Ojim  119  1)  —-  •  ... —h,-  ,  r ..  a  m)  i  f,iberewsi •  4 •  .  -  .  . •  —a.  •.  •  •. ..• .•4.  .$•_ ——  ‘  C.,•  •  •  •,  •  —.•.  -  a.— •••  •  .4  .  •  a.  —  •  •.•t•  —.  f••.  0 •  •  • ••  It •..  a  .  ..  . : .••. •4, • .,a• •  •  a.  .  •_(  ‘  • a  •  ••  a  .‘  _.  •1  • -  .  ••:•  ‘•...,4. •. •  .1.1. •  •,.  •  ...  •  • I.  •  •  La  g aa•4. •  a  a.  -  I  .‘  •:  .  a  j•  I 44  0  .  ••  •••  4  —.  b  b  ••••  4••I  •.• ! ,:•  c-::  A  n)  ‘t  •  o) Alzhêimer’s IiIase.. I  •:: -  a  a ‘a  ,  •  ,•  -  •  •1  ‘p  j  h  -  (  .  4  •  a  0’  -..  •  a.  • -•• •• .  ••  4• C.  -  a  .4  %  .  —  .  .  ••.  4  I.  •.  a  •  •••  a•. .  •  —.•  -•  .  ••  a.  4.  -  —  •%  •0 I  •  a  •: •  a.  Metastic Caiinoma  •  *  .:  a •  S  •  a..  .•  ‘•I  •  4 a.  d .45  •  •  :: :  •-  I. .0•  ;.  -  •  4  IL  •  •  •  S  •,  C  I  a  ••_  C  q)  I  •‘  -:  •  Gliomatosis C&ebri  •.  •-..‘ I  bC-SIGN p)  t  •..  La  v  a.  4  :  —  •4••)  a. •  .4  .  a’  a’.’  -  ‘L.I  a  a , I  .  a  *4 .4  4  a  a  S  .  :  _•  •• •  -.  DCiN: a  •  •  ,•  .II  DC-SIGN  Figure 15(1-g): DC-SIGN expression in situ in the CNS of patients with tuberculosis, sarcoidosis, neurodegenerative diseases, and tumours. Light micrographs show DC-SIGNpositive cells (stained red). Scale bars = 6Ojtm  120  120’  çioo *  80•  =  60’  *  * *  40  *  20  1  _L  MS  Ischemia Vasculitis  * *  =  =  *  *  Vasculitis Meningitis with CAA  * —  —  I II I  I I Normal Brain  *  1i Abscess  Viral Infections  —  I II I  Fungal ToxoChronic ALS Infections plasmosis Granulomatous inflammation  AD & AD with CAA  Disease Category  Figure 16: DC-SIGN expression in situ in normal and pathological CNS. Bar graph represents the quantification of DC-SIGN expression following light microscopy. Error bars indicate standard errors of the mean. The “*“ mark above bars indicates a significantly greater expression of DC-SIGN compared to normal brain sections (p <0.05).  121  Tumour  a)  Nor  s  _;_•,.  am  i’  •.•  :.-  b)  •-.‘•  •  •‘.  •‘:  .. •  •:  -  ..•  S •  ,  A  t  —  •‘e  -  —‘  S  -  ‘III. .  •  a.  %S  .-  ‘-‘j  !ac1n c)  Acjj..t  d)  :., S ‘..  -d: *s  -  ..  : Vascwii-, a ••: •;  ,  •  a..’.  -  -:  •.  ...-  •  -  .  •‘:  •.  -  ..  .  ,.  .--  jI  ...  --  _.••  .•  :.4,-.  i.: ‘p  C.  .  a  .‘.jI.  .,‘_  .  .-.•..  :‘ -:  %... I  .  •  .  ;  .••  .  -..  —.  •  .Li.  •i  .:  -  -  •  ‘•.•  ..  ,,.-  •  .•• •  ,  •:‘‘-  e)  Fascin  :-  .  y$socraed with ‘.  • : I •  I I  •  F...  •  I •  I .  •.‘  I..  .. ——S  .  . •s  A  ;.I.•  I  .-‘. .5.’  ‘  b  4a.  ‘.  •  a  —  , •.;  4-  $  •+:3  h’  Fàsêin  .‘  Figure 17(a-e): Fascin expression in normal CNS and in CNS inflammatory diseases. Light micrographs show fascin-positive cells (stained red). Scale bars 6Otm in  situ  =  122  p  ...  ..‘  S  0  g)  h)  i)  Infeçtioñ. I.e  -  •  .  4  :  ;. ,  ft  -.4  4  •.  r  I  :  •. •  .  I..  .  I, 4.  S.  if 5  5’  £1  je  C-  Facin  I.  ççyptococus 1tirn;  j)  I  S  .  %  S  I  4  S  .1  e  a  • ..  :‘  .  TôcoIamos1s  k)  •. 4.  ,  ‘a’.  a  .4.  S  -. .:  .  .q.’%  ‘.•  •.  ••  ,•.!Cf  as .Ie..  .  Fascin  •  ,  a  C .  :  I  .—; a .r 1 •  I  a  —  4  b  0 _  a  -  rJ ;  ;,•  If  •  •  Figure 17(f-k): Fascin expression in situ in infectious CNS pathologies. Light micrographs show fascin-positive cells (stained red). Scale bars 60p.m =  123  1)  m) •s  lI,b:. V1  •  • V  .  :... .  •4•  .  ‘V  ..‘  ‘‘ p  ,:  .  .  :  .  4’_,  •A  -  •  I:  =  V  •_:i.S*.f :  ‘‘  • V’:  4— • VV  ••  —  3-L  4  p_a.  •  :  ;•-  p  V•  •‘-  •  ,•.  •.  ,•.  4.  ••  ,_  —-  -  4V•  V:/  2,  -  :  •‘  4— -.  n)  &_  o) 7  .•  4  .  •V  zp  ;:: .;  %  ‘V  ZVV  *•  •  -  ‘:  a  .•.  -  .  q)  Glioffiatosis. Cerebi’  p)  •V.V V •V•’  ,  V  V  ••V  —.:-  •  •  •  9  •.•-‘.  •• a. •  .• S’  p  I’  p•  •  p  •i  •  I  V  •  V  •  a  ES  .‘.PV  •• V  P  ‘  I  .  4  • •  .1 •  ‘ 4.  •  :  • S  .  •  .  -.1  4  •;  jV  —  -  a  _,  •  •  •=,  •- T  •V•  •••  •  •  —.:  ,•‘—.  .4  p \V  *  :  •;  ••V  •  4  ‘•  a V  r.  •.:  •  ‘  ‘,  — ••,  •k-•’:  4  .Fascin 1 a  •  •  .:..-  ••  :•.;•• ‘1  a  •  ‘  0’  •.  4  .  •*.  -  •..‘•  - •• .•  Vp  •  %  •.  S 1  •VVVV  i•  I  •  •‘I- j  a  p_.  —  •  •- •  •‘  S.  ‘-  :  V  Fasqn,  Figure 17(1-g): Fascin expression in situ in the CNS of patients with tuberculosis, sarcoidosis, neurodegenerative diseases, and tumours. Light micrographs show fascin-positive cells (stained red). Scale bars = 6Opm  124  120 C%1  E E I  100 80  Q) 60 (I,  C)  * *  40 *  20  *  iii— Normal Brain  MS  Ischemia Vasculitis Vasculitis with CAA  Meningitis  Abscess  Viral Infections  Fungal Infections  Toxo. plasmosis  I  I  —  Chronic ALS Granulo matous inflammation  AD & AD with CAA  Disease Category  Figure 18: Fascin expression in situ in normal and pathological CNS. Bar graph represents the quantification of fascin expression in perivascular areas following light microscopy. Error bars indicate standard errors of the mean. The “*“ marks above bars denote a significantly greater expression of fascin compared to normal brain sections (p <0.05).  125  Tumour  a)  b)  .Jrma1 Brain •  .  .••  :  S  S  -  —  •  S  S  •:  • •  0  S 0  9.  •  .9  ••?;, : T •  : -  . p 1 (c r •J’  .  :  :: •  e)  • 0,  •  d)  . *  ••  CD4O•  Vascijlitis —  *  •  :-.-9:  -  •  .5 ‘  .  ..  e  •  •  :  •.•.  ‘••  :.  •:  I.S  .  ....  ••  I  ,  4?  ;.•.  .  .  S  .9  £.  :•cD4e  S  .‘..  •‘  •  0  •  •  •  •  S  LiteJnfaiç4 .:  :  ;‘•.:  .9  ‘I  c)  ••••  .5  •  0  S  !  €D40  -  *  V940  Vaseii1iti Associate4ith CA -  ••  ,-—  -  -  •  •9•l  S  J•  •_4  .59%  .  .  ..*  0_  -  •S  .0  a  •b  *  -  -  ..  I  S  •  •••  ,  9  \  _5*  •,  --s-:,  •  •  •• — ., ‘ •  .  .  ‘  1  •  ••  :  •• :. .  fr.çD4o’  Figure 19(a-e): CD4O expression in situ in normal CNS and in CNS inflammatory diseases. Note the CD4O-positive cells (stained red) in vasculitis with CAA. Scale bars = 60pm  126  g)  I)  s?es’.’j’r • •  .•  -  .  .  .  •  V•  •  .  :..  I  1 •I  .  •  •..•  1  .  ••I  • •  •  ..  •I.  “  .  ‘  L.  ••.  •  •t  •  I.  •  •t’  •,  “:•  I  •:.  I.  —  ••.  ,  •  .?  *  •.  ‘ :—  ••..—  •  ‘•  •_  -.  qS,•  •  •  dc  —.  :  :  ,  •  .  ‘.  -  •  •  ‘!  ‘•..  •  .•“..‘  •  S  Aspi11usiiiQection •  h)  •  I  ••.  V•  -,  •..  •  ¶  ••  •_•.,•  S  11  S.  I  ••  ••I•  ••  j  :.nl.F.  !.  •  •  .  •  .W•.  —  1  I  • ,‘.:  ,.  •._•.  b  —,  _..  •.,•.  .  I  —  ‘  .  •  •,..  —  I •  I  4 •  —  I  V  a  Crytoocçvdpfecijn  j)  S  k)  •  5  •  •  •  •  ê  •S  ••)  •  ••  .  •  S  -  IS •  5  .  a  •  •  • :  •  •  •  •  CD4p  t  I  S  •  I  •5 p.  •  -  •  •.••  •  S  •  •  S  .  —  ••  I. ••  ••  ••  ‘-•  I.....  .• .  •.  .  I •  ••  ••..  .. •  • I  r  •  .  ••  S  •  S.  C) •  .‘ •  55 .  •• a  Ct4 •...O.I’  Figure 19(f-k): CD4O expression in situ in infectious CNS pathologies. Light micrographs show CD4O-positive cells (stained red). Scale bars = 60j.tm Note: There are no CD4O positive cells in toxoplasmosis.  127  .  S  S  •  •  ••.  •  Sq.  •  :  I’  •1.  .5  ••  IIe  •  0 —  ‘I  •  I  •  i  ‘I.  • I  IS S  15  9  •  S  S  SLe.  •  4  .  —  •  •  I,  ‘.  • —  ‘  1.  V  •  •:  •  •.  •  I Uj  •  •  ‘  S  •  S.  •  -  S  •. S  •  5  1. 5 .i•  •  I  ‘3  5’  (..b.  •.  S  •S  •.•I.•  5.  •  .---  •  3••  •  •  •  • •  ‘I  ••  .  ‘  •I’.  •• •  U •  i_  S  S.  •  ••  1•1  • •  •  S  . •.  •  •‘ •1  I,  —  .  4•••  1)  t. S.  4.  •‘.l?._t. ,•*  44  •  I,,  •  •  4  .  F t —.-.  -  4  — 4  -  .-.  • •  . —  •4  4*•  •  a  •  • :  *  4  a  •  ,  I.  .*•  l,  4  ••  I  n)  o)  Fârnilial ALS  A1zheimjr’s Ifease  I  •  I  a.  a.  a  4  I  \  a..,  •  a,  4  4  4  I’  I a  I  4  •  • a  I  •.  .  a-  at  •*  1•  •  •  •  —  I..  •  IF  a  4 a  .•e.  CD40  •  a  •  •,  4  -  a  I —  P  -  CD4O  p)  11:o  :: *  •  :  .  .‘D4i  Figure 19(1-g): CD4O expression in situ in the CNS of patients with tuberculosis, sarcoidosis, neurodegenerative diseases, and tumours. Light micrographs show CD4O-positive cells (stained red). Scale bars = 6Oim Note: There are no CD4O-positive cells in familial ALS.  128  *  120  +  cloG E80 60 4)  40  2G —  Normal Brain  MS  Ischemia  I—I  Vasculitis Vasculitis with CAA  Il  li  Meningitis Abscess  Viral Infections  ..  III Fungal Infections  —  ToxoChronic ALS plasmosis Granulomatous inflammation  AD & AD with CAA  Tumour  Disease Category  Figure 20: CD4O expression in situ in normal and pathological CNS. Bar graph represents the quantification of CD4O expression in perivascular areas following light microscopy. Error bars indicate standard errors of the mean. The “*“ mark indicates a significantly greater expression of CD4O compared to normal brain sections (p <0.05).  129  a) •  ‘  -.;m.  Acute Infarct  c)  ‘•••‘  ;q  d) 4 —  d  g  p  I.  £  I  1 p  • p  ..:  • .rç, •  .‘  .  .  •  .‘  t  HLADR  e)  Figure 21(a-e’): MHC class II expression in situ in normal CNS and in CNS inflammatory diseases. Light micrographs show MHC class IT-positive cells (stained red). Scale bars = 60pm  130  I)  h)  i)  AspirgiIius Inction ‘I 4  •  • —  /  •  I  /  . L  ‘  -.  I  • ‘I • —-  pp  /  )  —.•  •  t-.  •  ‘a  -Ir  j)  •  ,•  ,  .‘  HLADR.,  A  k ipf::.: ‘ .  S  •  •. —  •  a  -,  ,  .:.  -.  :  ‘ê. 1. -  •  :•  .  •  :  ..  .  ::‘  .;  .-  .  ...  .  ..r  Figure 21(f-k): MHC class II expression in infectious CNS pathologies. Light micrographs show MHC class Il-positive cells (stained red). Scale bars 6Oiim in situ  =  131  •.  •  4._  ‘.---  1)  ‘.iI  4’.  .5•_-•  a ,...4.14_4:”  ‘,—  —_S..(  :c*1s, —-  -.  ,; t:  .:  -.  .9..  -.  .•  •  • S  ...  :-  •  -  9•. —  -  .  •  •.:  ‘ — —  a  -••  a..  •-•  —  “‘:;—..: ..  •  c.,  -..  ALS  •  —  .  ‘  9-. •  •  HLAPR%i -4  4  c  ..  •c:  i -  o)  S.3  —  •  .;  . • ..‘  .,  ••.  •  •  .  ••.  •  :  :  ••.  •1  I’.  -  ..-..  ‘  .  9  •  i  ,  •  J.  .  -  :-.  •  •  •  :4  b.  ‘  -  • :-.  s% .  , •  •  •  LAIDR  a  •J  —  ••  9’  ..•  -  -a  .•.  •  b  ••..  -  •  .b  •.  • .  a ,.a. ,  %  •  -  -.9  ••  •  •.•  .,  ‘  s___.  ..  ‘  .  ..  .  )  -  --c  p..  ••—-  •  4 • t  -  •  q)  Gliomatosis çeiebrt’ 4  • S  3’S  • :  S  /  *  S  •  b  a  a  •.  • %,a  S  p .)  *  :1 9  •  •I 4-  a a  9  .  ‘4  V.._ •  4•9  f’ji — .:v a  a  .  9•.  -  4..’  . 4  •  *1  5  :  -  Figure 21(1-ci): MHC class II expression  a -.  •  I4 ILA-DR r  .  —  95  4  ••  a,  .  .—, •-, -  9  t.  -  .  ‘  .  9  •  *  a;,.  ,.  4 I  -—  .  -‘a  • a  •k  4.*•  •  V  4  ‘4  •  ...•  ••  •  — •-  4  ¶••  .-  .5.  —  :  •0  ••  •5  I  •‘  b  •  .m  _:  I  a  ..  ‘a-.  1 p)  .  ,-  •  I  •-;  .•  •  *  b  J  ‘  •A  .-•.  a•-’  ..‘e  .•  —  I  .-1•  ••-.-  .  •.  L  ,  •  4  •  *  4’. •  —  I’  :  :td.!  •  a.. • •%•• ••  • •1  a  •.. ‘*,ø_ . •..  _•.  •  ,  a.  9.  ••W.44%  .:“  •_  W’  •  •,  . —  .-•.•s—..’  I  —  .--—  ••4  ..‘I  n)  :.  -  • -:‘L,  m)  •.  1ILA!DR  -  in situ in the CNS of patients with tuberculosis, sarcoidosis, neurodegenerative diseases, and tumours. Light micrographs show MHC class IIpositive cells (stained red). Scale bars = 6O.tm  132  *  c1  E  E *  0 0.  *  *  *  CI)  *  0 C)  *  *  *  Fungal Infections  Toxoplasmosis  *  Normal Brain  MS  Ischemia Vasculitis  Vasculitis Meningitis sith CAA  Abscess  Viral Infections  Chronic ALS Granulo. matous inflammation  AD & AD with CAA  Disease Category  Fi2ure 22: MHC Class II expression in situ in normal and pathological CNS. Bar graph  represents the quantification of MHC class II expression in perivascular areas following light microscopy. Error bars indicate standard errors of the mean. The “*“ mark above bars indicates a significantly greater expression of MHC Class II compared to normal brain sections (p < 0.05).  133  Tumour  120.  E E I  0  a.  100. 80. 60.  0  40.  0 41:  20.  C-)  n.  —.1Li Normal Brain  MS  -‘-  Viral Ischemia Vasculitis Vasculitis Meningitis Abscess Infections with CAA  Fungal Infections  Disease Category  Toxoplasmosis  Chronic ALS Granulomatous inflammation  —.  AD & AD with CAA  Tumour  Fi2ure 23a: Immature DC participation in normal and pathological CNS tissue in situ. Bar graph is the result of subtracting the number of fascin-positive cells from the number of DCSIGN-positive cells (see Figs. 14 & 16) which were quantified following light microscopy. Error bars indicate standard errors of the mean. The “*“ mark above bars indicates a significantly greater value compared to normal brain sections (p <0.05).  *  *  *  C1  E E I  0  a.  * *  U) 0  C-) ‘4-  0 4*:  LII  Normal Brain  MS  Ischemia Vasculitis  Vasculitis with CAA  Meningitis Abscess  Viral Infections  Fungal Infections  ToxoChronic ALS plasmosis Granulo matous inflammation  AD & AD with CAA  Disease Category  Fhwre 23b: Immature vs. mature DC participation in normal and pathological CNS tissue in situ. For each condition, the left-hand bar depicts immature DC numbers (calculated as above) and the right-hand bar represents mature DC numbers (same as fascin-positive cells). Error bars represent standard errors of the mean. denotes a significant difference between immature and mature DC presence in each disease category (p <0.05). “*“  134  Tumour  

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