Pathophysiological Role of CD6 and Its New Ligand in Disease

PATHOPHYSIOLOGICAL ROLE OF CD6 AND

ITS NEW LIGAND IN DISEASES

GOSPEL ENYINDAH-ASONYE

Submitted in partial fulfillment of the requirements for the degree of

Doctor Of Philosophy

Department of Molecular Medicine

CASE WESTERN RESERVE UNIVERSITY

May 2017

CASE WESTERN RESERVE UNIVERISTY

SCHOOL OF GRADUATE STUDIES

We hereby approve the dissertation of Gospel Enyindah-Asonye, Candidate for the degree of Doctor of Philosophy*

Committee Chair Christopher King, MD PhD

Committee Member Feng Lin, PhD

Committee Member Neetu Gupta, PhD

Committee Member Trine Jorgensen, PhD

Committee Member Brian Hill, MD PhD

Committee Member Thomas Hamilton, PhD

Date of Defense January 11, 2017

*We also certify that written approval has been obtained for any propriety material obtained therein

TABLE OF CONTENTS

LIST OF FIGURES……………………………………………………….……………iii

LIST OF TABLES……………………………………………………..………………..iv

ABSTRACT………………………………………………………….………………..…v

CHAPTER 1. Introduction………………………………………………..…………….1 i. Intestinal I/R-induced injury………………………..……………………..2 ii. Natural IgM and the initiation of Intestinal I/R-induced injury………..… 4 iii. B1 cell development and self-renewal………………………………….. 6 iv. Mechanism utilized by B1 cells to secrete natural IgM……………..…….9 v. B cells contribute to tissue homeostasis………………………….………10 vi. B1 cells are influenced by their local microenvironment…………….... .12 vii. CD6 and expression distribution………………….…………….……..…14 viii. Regulatory control of CD6 expression………………………….…… ….16 ix. CD6 interacting proteins……………………………………………….. 17 x. CD6 role in T cell development and immune response………….………18 xi. CD6 role in B cells………………………………………………….……21 xii. IOR-T1/Itolizumab (CD6 mAb)………………………………….……...22 xiii. CD6 as a therapeutic target in RA clinical studies………………….……24 xiv. IOR-T1/Itolizumab immunomodulatory mechanism of action….……….25 xv. Evidence for additional CD6 ligands…….………….…………………...27 xvi. CD318………………….………………………………………….……..28 xvii. CD318 expression profile and regulatory control…………….………….29 xviii. CD318 role in cancer………………………………….……….……… 30 xix. CD318 interacting proteins…………………….………………………...32 xx. Therapeutic potential of targeting CD318…………………….……….... 33

CHAPTER 2. Materials and Methods……………………….……………………… 36

CHAPTER 3. CD6 regulates intestinal/reperfusion-induced injury………….……49

I. Introduction……………………………………………………………...50 II. Results………………………………………………………………… ..52 III. Discussion……………………………………………………………… 66

CHAPTER 4. CD318 is a novel CD6 ligand………………………………………….73

i I. Introduction………………………………………...…………………...73 II. Results………………………………………………...………………...75 III. Discussion…………………………………………...………………….86

CHAPTER 5. Future directions……………………………...………………………..94

I. Summary and future directions part 1………………………...………….94 II. Summary and future directions part 2……………………...………….…99

References…………………………………………………………...…………………106

List of Figures

Figure `1: CD6-/- mice are protected from intestinal I/R-induced injury……………….. 53

Figure 2: Ischemic-antigen specific IgM titers are reduced in the CD6-/- mice……….…55

Figure 3: CD6 is not detectable on resting or activated B2 cells……………………….56

Figure 4: CD6 is expressed on non-peritoneal B1 cells……………………………. …...57

Figure 5: Peritoneal B1a cells gain CD6 expression outside the peritoneal cavity… …. 58

Figure 6: The B1a cell populations are reduced in the CD6 -/- mice…………………….59

Figure 7: Reduced B1a cells in the CD6-/- mice are due to impaired B1a cell self-renewal…………………………………………………………………..… …...…..60

Figure 8.: The flow cytometry gating strategy for B1 cells ………………………….….64

Figure 9. CD6 expression on Bone marrow B1 cells and its effect on B1 cell population in the bone marrow and peritoneal cavity……………………………...…….65

Figure 10. B1/a cell proliferation are not reduced in the bone marrow and peritoneal cavity of CD6-/ mice…………………………………………………………..66

Figure 11. Identification of the antigen recognized by mAb 3A11………………..…….77

Figure 12. The anti-CD318 mAb and mAb 3A11 have identical staining patterns on cells previously known to express or lack CD318 expression on the cell surface………………………………………………………………………..79

III

Figure 13. The anti-CD318 mAb and mAb 3A11 have identical staining patterns on cells engineered to upregulate or downregulate CD318 expression..,,,,,,,,,,,,.80

Figure 14. CD6 interacts with CD318…………………………………………………...82

Figure 15. CD318 is a potential biomarker for inflammatory arthritis and chemotactic for T cells………………………………………………………………85

List of Tables

Table 1: Frequency and absolute numbers of B1 cells for WT and CD6-/- mice………..60

Table 2: Frequency and absolute numbers of Bone marrow and

peritoneal B1/a/b cells for WT and CD6-/-- mice…………………………………..…….61

Pathophysiological Role of CD6 and its New Ligand in Disease

Abstract

GOSPEL ENYINDAH-ASONYE

Intestinal ischemia/reperfusion (I/R) injury is a relatively common pathological condition that can lead to multi-organ failure and mortality. Regulatory mechanism for this disease is poorly understood, although it is established that circulating pathogenic natural IgM, which is primarily produced by B1a cells outside of the peritoneal cavity, are integrally involved. CD6 was originally identified as a marker for T cells and later found to be present on some subsets of B cells in humans, however, whether CD6 plays any role in intestinal I/R induced injury and if so, what are the underlying mechanisms, remain unknown. Here we report that CD6-/- mice were significantly protected from intestinal inflammation and mucosal damage compared to WT mice in a model of intestinal I/R-induced injury. Mechanistically, we found that CD6 was selectively expressed on B1 cells outside of the bone marrow and peritoneal cavity, and that pathogenic natural IgM titers were reduced in the CD6-/- mice in association with significantly decreased B1a cell population. Our results reveal an unexpected role of

CD6 in the pathogenesis of intestinal IR-induced injury by regulating the self-renewal of

B1a cells.

Acknowledgements

I would like to thank everyone that has contributed in some way to the work presented in this thesis. Foremost, I would like to express my sincere gratitude to my thesis advisors

Dr. Feng Lin and Dr. Neetu Gupta for their continuous support through out my thesis study, for their motivation, enthusiasm, and commitment towards my scientific development.

Besides my advisors, I would like to thank the rest of my thesis committee: Dr.

Christopher King, Dr. Trine Jorgensen, Dr. Brian Hill, and Dr. Thomas Hamilton, for their encouragement, insightful comments/advice and tough questions.

I would like to thank my lab members: Yan Li and Lingjun Zhang for all their help, inputs, suggestions during lab meeting as well as providing a fun and exciting lab environment to work during last 3 years. I owe a large debt to Yan without whom the intestinal ischemia/reperfusion injury surgery and other in vivo immunization studies could not have been done. I would also like to thank our collaborator Dr. David Fox and his lab members for performing the T cell migration assay and assessing the levels of soluble CD318 in the synovial fluid of RA patients.

Finally, I would like to thank my family and friends for supporting me through out my

PhD study.

VII

List of Abbreviations

ALCAM- Activated leukocyte cell adhesion molecule B-CLL- B cell chronic lymphocytic leukemia BCL6- B cell lymphoma protein 6 BCR- B cell receptor CIA- Collagen-induced arthritis Cr- Complement receptor DSS- Dextran sulfate sodium EAE- Experimental autoimmune encephalomyelitis EGFR- Epidermal growth factor receptor FL- Full length Tdt- Terminal deoxynucleotidyl transferase HER2- Human epidermal growth factor receptor 2 HMGB1- High mobility group box chromosomal protein 1 IL-Interleukin IRF4- Interferon regulatory factor 4 I/R- Ischemia and reperfusion LDL-Low-density Lipoprotein LPS- Lipopolysaccharide

MAPK- Mitogen activated protein kinase MS- Multiple Sclerosis NMHC- Non-muscle myosin heavy chain PAX5- Paired box 5 PBMC- Peripheral blood mononuclear cells PHA- Phytohaemagglutinin PKC- Protein kinase C PMA- Phorbol myristate acetate PRR- Pathogen recognition receptor

VIII

RAG- Recombination-activating gene ROS- Reactive oxygen species SCID- Severe combined immunodeficiency SS- Sjogren Syndrome TCR- T cell receptor TEC- Thymic epithelial cell TLR- Toll like receptor Tyr- Tyrosine

CHAPTER ONE: Introduction

1 Introduction to intestinal ischemia and reperfusion induced-injury

Intestinal ischemia and reperfusion (I/R) induced-injury ensues when blood circulation to a particular tissue/organ is interrupted for a period time and subsequently restored. The restoration of blood circulation surprisingly provokes an acute inflammatory response that contributes to tissue injury [1]. It stems from diverse etiologies and affects people of all ages. In the neonatal population, intestinal I/R induced-injury occurs during necrotizing enterocolitis and midgut volvulus[2]. In adults, intestinal I/R injury often occurs during abdominal and thoracic vascular surgery, intestinal transplantation, hemorrhagic shock, and surgery using cardiopulmonary bypass.

The mortality rate can be as high as 40% especially when surgical intervention to salvage the ischemic tissue is necessary[3]. Nevertheless, without intensive management and treatment, the leakage and migration of endotoxins across the damage intestinal epithelial barrier to peripheral lymphoid organs could lead to sepsis, systemic inflammation, multiple organ failure, and eventually, death [4]. Although intestinal ischemia usually occurs in a sterile environment, there is an activation of the innate and adaptive immune system that contributes significantly to tissue injury. The immune response during the reperfusion of the tissue is very similar to a host immune response against a foreign pathogen such as the involvement of pro-inflammatory cytokines and recruitment of inflammatory cells into the ischemic tissue site[5].

The pathogenesis of intestinal-I/R induced-injury

Intestinal I/R induced injury is primarily comprised of two main phases. The first phase of intestinal I/R injury involves the disruption of normal epithelial barrier integrity as a result of intestinal epithelial cell death. The enterocytes which are nutrient-absorbing

cells found on the cell surface of the small intestine are moderately resistant to momentary hypoxic condition, however, prolong exposure to the hypoxic condition can result in irreversible cell death [6-8]. Enterocytes undergoing cell death release various intracellular components into the extracellular matrix that binds to germline encoded pathogen recognition receptors (PRR) on the immune cells inducing an inflammatory response that further aggravates tissue injury. The released intracellular components include molecules such as nucleic acid, proteases, lysozymes and heat shock proteins which are also referred to as damage-associated molecular pattern[9]. The paracellular tight junctions are also disrupted as a result of the hypoxia-induced enterocytes cell death

[10].

During the second phase of intestinal I/R-induced injury, the re-establishment of blood flow further worsens the ischemic tissue due to systemic activation of the immune system. The recognition of the accumulated ischemic cellular debris by PRRs on the immune cells induces the expression of transcription factors that activate genes of mediators that promote inflammatory response[11-13]. These mediators are mostly proinflammatory cytokines such as tumor necrosis factor –alpha (TNF-α) and interleukin

(IL)-1,6 [10, 14, 15]. These mediators promote complement activation, expression of adhesion molecules, chemokines as well as the recruitment and transmigration of activated leukocytes into the damaged tissue [16, 17]. Within the damaged tissues, these infiltrated leukocytes secrete additional pro-inflammatory cytokines and mediators including IL-6, IL-8, IL-17, nitric oxide, leukotrienes and reactive oxygen species (ROS)

[18-21]. These secreted pro-inflammatory mediators further inflict more intestinal epithelial tissue damage that further contributes to the disruption of the epithelial tight

junctions if anti-inflammatory regulatory responses are inadequate. Interestingly several studies have implicated natural antibodies as the primary initiator of the acute inflammatory response typically associated with reperfusion of the intestinal ischemic organ [22].

Introduction to natural antibodies

Natural antibodies are antibodies that are found in the serum of naïve mice or germ-free mice in the absence of any obvious antigen stimulation or immunization.

Natural antibodies are germline-encoded and predominantly of the IgM isotype [23].

They lack N-region addition and somatic hypermutation due to the absence of the enzymes terminal deoxynucleotidyl transferase (Tdt) and activation-induced (cytidine) deaminase (AID) in natural antibody-secreting B cells. The majority of the pool of natural IgM recognizes phosphorylcholine an antigen expressed on bacteria cell wall as well as on apoptotic cell membrane and oxidize lipids. Natural IgM plays a major role in providing first line defense against pathogens and participating in tissue homeostasis by regulating the clearance of apoptotic cell debris due to its inherent poly specificity.

Neoepitopes expressed on ischemic-apoptotic cell membrane can be recognized by natural IgM following reperfusion of the tissue or organ resulting in activation of complement, neutrophil recruitment, and tissue or organ injury.

Natural IgM and the initiation of Intestinal I/R-induced inflammation

Natural IgM is critical in the pathogenesis of intestinal I/R. These natural antibodies recognize neoantigens exposed on hypoxic cells after intestinal ischemia, and activate complement through the lectin pathway to produce C5a, which can facilitate

neutrophil and monocytes infiltration and inflammatory cytokine (e.g. IL-6, IL-1 and

TNFα) production [24, 25]. Circulating neutrophils infiltrate rapidly into the injured tissue producing free radicals to kill bacteria and control infection by sterilization. These free radicals can also induce bystander tissue injury further exacerbating tissue injury.

The first evidence indicating that natural IgM initiates the inflammatory processes that induce intestinal I/R-induced injury was found in studies using the immunodeficient

RAG-/- mice, which do not have any antibodies [26]. These mice are highly resistant to intestinal I/R-induced injury in association with significantly reduced IL-6 production, but become susceptible again following reconstitution with purified serum IgM from naïve WT mice, highlighting the critical role of natural IgM in the development of intestinal I/R-induced injury [22, 26-29]. In a related study, Cr2-/- mice that lack the complement receptors 1 and 2 were found to be resistant to intestinal I/R-induced injury, despite displaying similar titers of natural IgM in the serum. The adoptive transfer of natural IgM from the serum of wild-type mice into Cr2-/- mice restored intestinal- injury[30, 31]. The administration of IgG alone into Cr2-/- mice did not restore intestinal ischemia but rather enhanced the infiltration of neutrophils when infused together with

IgM[30]. These data suggested that there is a subset of natural antibody repertoire that contributes to the pathogenesis.

Select subset of natural antibodies initiate intestinal I/R induced injury

In support of the earlier observation that innate recognition of ischemia-induced self-antigens was involved in the initiation of ischemia-induced injury, a single clone of natural IgM (CM22) that could restore significant intestinal I/R-injury in RAG1 -/- was identified from a panel of hybridoma that was generated from peritoneal enriched B1-

cells[29, 32]. In a later study, non-muscle myosin heavy chain II (NMHC II) A was identified as the target for the natural IgM CM22 that could initiate intestinal and skeletal-I/R injury in RAG 1-/- mice[29, 32].

In a different study, another novel IgM mAb (mAb B4) capable of inducing complement activation, neutrophil recruitment and intestinal injury in IR-resistant

RAG1−/− mice was recently reported [33]. Annexin IV was identified as the antigen specifically recognized by this mAb B4. Annexin IV is a member of the calcium and phospholipid-binding protein family[34]. It is mainly produced by epithelial cells and highly expressed in the intestine, liver, kidney and lung[35]. The location of annexin IV varies depending on the cell type. Annexin IV can be found in the cytoplasm or the basal- apical domains of the plasma membrane[13, 36]. Importantly, the surface localization of annexin IV has been associated with cell death and apoptosis[12, 37]. The administration of recombinant Annexin IV to wild-type mice prevented intestinal I/R-injury establishing

Annexin IV as an essential ischemic-neoantigen targeted by natural antibodies during the reperfusion phase[33]. Collectively, these findings suggest a general mechanism by which stress, such as hypoxia, leads to expression of a highly conserved neo-epitopes on hypoxic cells and recognition by pathogenic natural IgM during reperfusion-phase results in complement activation, neutrophils infiltration, and tissue injury.

Introduction to B1 cells

B1 B cells spontaneously produce natural IgM in the absence of any antigen- stimulation. [38-40]. B1 cells are a unique subset of B cells that are different from conventional B2 B cells based on their function, primary localization, development and cell surface markers. B1 cells express the cell surface markers IgM, CD19, B220, and

CD43. They lack the expression of CD23, CD21 and while in the peritoneal cavity, B1 cells express the macrophage marker CD11b. B1 cells can be further subdivided into

B1a (CD5+) and B1b (CD5-) [41]. B1a cells primarily produce natural antibodies, while

B1b cells participate in T-independent memory response. The majority of the natural

IgM (80-90%) in the serum of naïve mice are produced by splenic and bone marrow B1 cells. An analysis of IgM secretion by B1 cells in different compartments indicated that peritoneal B1 cells do not spontaneously secrete IgM and that splenic B1 cells produce more IgM than bone marrow B1 cells[38]. These natural antibodies producing B1 cells are phenotypically distinct from conventional B2 differentiated plasma cells. They express the markers IgM and CD19 but lack the differentiated plasma cell hallmark marker CD138 unlike activated B2 cells that downregulate cell surface marker IgM and acquire CD138 surface expression during plasma cell differentiation[42, 43]. Peritoneal

B1 cells also contribute to the pool of natural antibodies by migrating out of the peritoneum into the spleen where they differentiate into IgM-secreting plasma cells[44,

45].

B1 cell development

B1 B cells are called “B1” because they develop before conventional B2 cells and their population in adult mice is maintained by their inherent ability to undergo self- renewal. B2 B cells are termed “B2” because they develop later in ontogeny after B1 cells and unlike B1 cells that can undergo self-renewal; B2 cells are dependent on the bone marrow[40]. The development of B1 B cells has been a debatable topic for some time and that has led to the emergence of two main hypotheses that are not entirely exclusive.

The first hypothesis proposes that B1 cells arise from a specific progenitor that is distinct from conventional B2 and T cells. B1 cells develop from Lin-CD19+B220- progenitor cells that are abundantly present in the fetal liver and bone marrow[41, 46,

47]. These progenitor cells are also present in the adult bone marrow and spleen albeit in low numbers and do not function efficiently as the fetal-derived progenitor cells. The adoptive transfer of pro-B cells isolated from the fetal liver reconstituted the B1 compartment in SCID mice efficiently but not pro-B cells isolated from adult bone marrow. Pro-B cells isolated from adult bone marrow efficiently reconstituted the B2 cell compartments and poorly reconstituted the B1 cell compartment, specifically they reconstituted only the B1b cells subset but not B1a cells [10, 48].

The second hypothesis proposes that B1 cells arise from a single B cell progenitor as a result of B cell receptor signaling strength. This hypothesis was proposed based on the observation that CD5 expression; a bona fide marker of B1a cells is induced on B2 cells following B cell receptor (BCR) crosslinking and the effect of BCR signaling strength on B1 cell numbers[42, 49]. The deficiency of certain positive regulators of BCR signaling results in reduced numbers of B1 cells [50-52], while the deficiency in negative regulators of BCR signaling results in increased numbers of B1 cells[53]. This indicates that unlike B2 cells that are deleted upon strong BCR binding to self-antigens, B1 cell development is fostered.

B1 cell self-renewal

The spleen is critically required for B1 cell homeostasis. Splenectomized mice specifically display reduced B1- but not B2 B cells in the peripheral blood and peritoneal cavity[54, 55]. These findings led to the proposal that renewal of B1 cells occurs in the

spleen. Further studies comparing the rate of B1 self-renewal in different tissues reported that splenic B1 cells have a higher turnover rate compared to B1 cells localized in other tissues[56, 57]. The ability of non-splenic B1 cells to undergo self-renewal albert lower than splenic B1 cells could also account for the reduced peritoneal B1 cells in splenectomized mice. The adult spleen also harbors B1 cell specific progenitor cells that could be activated upon stress to replenish or maintain the B1 cell population further indicating the significance of the spleen in regulating B1 cell homeostasis[58].

B1 cells utilize mechanism similar to B2 cells in secreting Natural IgM

B1 B cells in the spleen and bone marrow contribute significantly to the pool of natural antibodies in the serum of naïve mice[38]. This is consistent with reports that bone marrow residing B1 cells are phenotypically similar to splenic B1 cells but distinct from peritoneal B1 cells[42]. The mechanisms utilized by B1 cells to spontaneously secrete natural antibodies are poorly understood. However, several studies indicate that

B1 cells utilize a similar pathway for antibody production as B2 cells[38, 59]. In conventional B2 cells, IRF4 initiate a cascade of transcription factors that promotes plasma cell differentiation and antibody secretion. The induction of the transcription factor IRF4 suppresses BCL-6 expression, which results in elevated BLIMP expression[60, 61]. The increase of BLIMP-1 expression results in the suppression of

PAX-5. The suppression of PAX-5 expression permits XBP-1 levels to increase[62, 63].

The expression of XBP-1 induces the activation of genes that are involved in the immunoglobulin secretory pathway[63]. IRF4 is required for natural IgM secretion by splenic B1 cells[38]. Whether natural IgM secretion by bone marrow B1 cells is also dependent on IRF4 is currently unknown. Furthermore, peritoneal B1 B cells

spontaneously produce very little amount of natural antibodies. This finding is consistent with the lack of detectable antibody-secreting plasma cell associated transcription factors[64]. However, in response to B1 cell activation, peritoneal B1 cells rapidly egress out of the peritoneal cavity and into several peripheral tissues such as the spleen and lymph node where they rapidly upregulate BLIMP-1 and undergo differentiation into

IgM/IgA-secreting cells[44, 65]. The egress of peritoneal B1 cells mediated by TLR signaling requires the coordinated transient down-regulation of integrins and CD9, thereby allowing the detachment from the local matrix [66].

B1 cells contribute to tissue homeostasis

B1 cells are also critically important in maintaining tissue homeostasis due to the inherent ability of natural IgM to bind to self-antigens, which can be expressed on apoptotic cells. The significance of natural IgM ability to bind to self-antigens has been explored in several disease models. They play an important role in preventing the development of atherosclerosis by binding and facilitating the clearance of oxidized lipids and apoptotic cells within the carotid artery[39, 67]. The deficiency of serum IgM in LDL receptor-deficient mice resulted in larger and complex aortic root atherosclerotic lesions as well as accelerated cholesterol crystal formation[68]. Increased apoptosis was also observed in the double-deficient mice compared to LDL receptor-deficient mice alone. In a different study, the splenectomy of apolipoprotein E-deficient mice that selectively reduced the B1 cell population and titers of serum natural IgM resulted in elevated atherosclerotic lesions and increased necrotic cores[69]. B1 cells have also been implicated in reducing the severity of autoimmune pathology associated with IgG autoantibodies in Lpr mice. Lpr mice deficient in serum IgM displays enhanced

production of double-stranded DNA and histones specific IgG autoantibodies[69]. These mice also have severe glomerulonephritis and succumb to the disease at an earlier age compared to regular Lpr mice.

In addition to the housekeeping function of B1 cells mediated by natural IgM, B1 cells also secrete large amounts of IL-10 that suppresses T cell immune response[69].

B1a cells produce a high concentration of IL-10 in response to certain viruses and toll- like receptors stimulation[23]. Neonatal B1 cell secretion of IL-10 in response to CpG stimulation was reported to suppress optimal IL-12 production by dendritic cells resulting in inadequate Th1 priming in vivo[70]. In later studies, B1-derived IL-10 has been reported to reduce the disease severity of DSS-induced mouse colitis. These studies highlight an additional way for B1a cells to regulate inflammatory responses and contribute to tissue homeostasis[71].

Effector functions of B1 cells

In response to infection, several studies have shown that B1 cells can undergo two distinct immune responses and outcomes. B1 cells can secrete high titers of polyclonal

IgM within the local site of infection or produce antigen-specific antibodies that provide long-term protection depending on the antigen/pathogen[72]. Infection with influenza virus induces the rapid redistribution of B1a cells from the peritoneal cavity to regional lymph nodes in response to probably innate signals[73]. In the lymph node, these mobilized B1a cells undergo differentiation into IgM-secreting cells. This response has been reported to be polyclonal based on the observation that these differentiated B1a cells do not undergo clonal expansion and/ or secrete virus-specific antibodies[73]. The administration of LPS or streptococcus pneumonia also induces the rapid mobilization of

peritoneal B1a cells to the spleen or mucosal tissue where they undergo proliferation and differentiation into antigen-specific IgM or IgA-secreting cells[44, 74, 75]. The ability of

B1-derived antibodies to directly neutralize, induce complement activation or enhance the adaptive B-2 cell immune might contribute to the potential mechanism by which B-1 cells provide protective effector function.

B1 cells are influenced by their local microenvironment

The phenotypic, functional and transcriptional gene profile differences between splenic and peritoneal B1 cells has been a subject of controversy as to whether these cells represent the same population of B1 cells residing in different peripheral tissue compartments. In terms of surface antigen expression, one of the major differences between splenic and peritoneal B1 cells is the expression of CD11b[76]. It is well known that CD11b is exclusively expressed on peritoneal B1 cells. The presence of CD11b- positive B1 cells in the spleen represent recent migrant B1 cells from the peritoneal cavity and likewise, CD11b-negative B1 cells in the cavity represent recent arrivals into the peritoneal cavity[44, 77]. Additionally, peritoneal B1 cells express significantly elevated cell surface levels of CD80 and IgM compared to splenic B1 cells[78].

Interestingly, peritoneal B1 cells differ from splenic B1 cells in their responsiveness to BCR crosslinking. Although peritoneal B1 cells express higher levels of BCR than splenic B1 cells, engagement of the BCR results in modest intracellular calcium influx, reduced proliferation and enhanced apoptosis[79, 80]. While splenic B1 cells display robust calcium influx and signaling phenotype similar to splenic B-2 cells following engagement of the BCR [79]. It has been suggested that the unique phenotype of peritoneal B1 cells BCR-signaling and activation state are induced by chronic antigen

stimulation. These antigens are believed to be unique to the peritoneal cavity. In support of this observation, peritoneal B1 cells express significantly elevated basal levels of

CD80, phosphorylated STAT3 and ERK compared to splenic B1 cells [81]. The BCR hyporesponsiveness of peritoneal B1 cells has been attributed to the expression of the Src family kinase Lck that is exclusively expressed on peritoneal but not splenic B1 cells[82].

The expression of Lck can be induced on splenic B1 cells after adoptive transfer into the peritoneum and conferring a hyporesponsive phenotype to these adoptively transferred cells.

The production of natural antibodies is also one of the major differences between splenic and peritoneal B1 cells. B1 cells residing in the spleen and bone marrow spontaneously produce natural antibodies[38, 42]. These data fit well with the observation that anti-Gal antibodies are specifically produced by splenic B1 cells and not peritoneal B1 cells[45]. Peritoneal B1 cells also contribute to IgM production by migrating to peripheral tissues such as the spleen and lymph node where they undergo differentiation into IgM-secreting cells in response to an infection or specific innate- signals. Peritoneal cavity macrophage-mediated production of prostaglandin E2 has been attributed to the inability of peritoneal B1 cells to secrete IgM in vivo while in the peritoneal cavity[83].

B1 cells are also found in the bone marrow and these cells contribute significantly to the pool of natural antibodies. These cells have not been extensively characterized, however they express the cell surface markers CD19 and CD43 as splenic B1 cell counterpart[42]. Whether this bone marrow B1 cells extensively differ from splenic or peritoneal B1 cells remains to be explored. Overall scientific data indicates that B-1 cells

are homogenous and are heavily influenced by their local microenvironment resulting in phenotypic and functional differences between B1 cells residing in the spleen, peritoneal and perhaps bone marrow[82, 84, 85].

How these natural IgM-producing B1a cells are regulated are also poorly understood, despite the findings that antigen specificity and BCR signaling strength are critical factors in B1a cell development because deletion of BCR co-stimulatory molecules such as CD19 results in a massive reduction of B1a numbers, while deletion of the negative regulators of BCR signaling such as Siglec-G leads to a vast increase in B1a cell population [50, 86].

Introduction to CD6 and expression distribution

CD6 is a member of the scavenger receptor of family proteins. It is a cell surface glycoprotein with a molecular weight range of 105-130 kDa[87]. CD6 is expressed on all mouse and human T cells [88]. In regards to B cells in humans, CD6 is expressed on mature and human B1a B cells and its expression on mouse B cells has not been previously reported[89]. Natural killer (NK) cells express CD6, however its expression is restricted to only CD56-dim but not CD56-bright NK cells[90]. The expression of CD6 has also been reported in various regions of the brain. CD6 is also expressed in the basal ganglia and cortex region in the central nervous system[7]. Accumulating evidence in the literature throughout the years has indicated that CD6 plays a co-stimulatory role in T cell activation. However, recently published data contradicts results from the early studies and attribute an inhibitory role to CD6 in T cells. Hence, it appears that CD6 has a dual role in regulating T cell intracellular signaling.

Structural components of CD6 and CD6 isoforms

CD6 is composed of 3 extracellular domains termed domains (termed domain 1, 2 and 3 respectively) that are anchored to the cell surface via a transmembrane domain and a long cytoplasmic domain [91]. CD6 is composed of 13 exons and localized on chromosome 11 at 11q13, which is in close proximity to CD5, a related molecule[2, 92].

Exon 1 to 6 encodes for domain 1, 2 and 3 of CD6 extracellular portion. Exon 7 encodes the transmembrane sequence and the cytoplasmic domain is encoded by exons 8 to 13[2].

The analysis of CD6 mRNA transcripts from B cells obtained from chronic lymphocytic leukemia patients and PHA-activated PBMC disclosed the presence of five different transcripts. These five transcripts result from variable splicing of CD6 cytoplasmic domain encoding exons[2]. The presence of these spliced isoforms has also been reported in T cells[93]. However, the functional significance of these alternatively spliced isoforms is unknown. The lack of functional information may be due to the lack of abundance of these spliced isoforms as previously shown by northern blot analysis[2].

CD6 has been proposed to contribute to the diversification of TCR signaling by interacting with proteins containing SH2 and SH3-domains[94-96]. Hypothetically, it’s possible that some of these spliced isoforms might be able to interact with both SH2 and

SH3-domain containing proteins, while others can only selectively interact with either

SH2 or SH3-domain containing proteins. Future studies characterizing the intracellular signaling molecules that bind to the various CD6 spliced isoforms upon binding of specific CD6 ligands will be critical in elucidating their signaling capabilities and function on immune cells that predominantly expresses a specific CD6 spliced isoform.

An isoform of CD6 resulting from exon 5 skipping has also been observed. The skipping of exon 5 results in an isoform of CD6 that lacks the reported membrane proximal domain that the well-described CD6 ligand CD166 binds to. The skipping of exon 5 is mediated by the splicing factor SRF1[92].

The T cell expression profile and immunological synapse localization of this domain-3 lacking isoform are remarkable different from full-length CD6. During antigen presentation full-length CD6 but not domain-3 lacking isoform targets to the T cell/APC interface [97]. Analysis of different rat thymic T cell subsets, revealed that domain-3 lacking isoform constitutes a small percentage of double-positive thymocytes but constitutes about 50% of CD4 or CD8-single positive thymocytes [97]. This switching between full-length CD6 to domain-3 lacking isoform may play an important role in T cell development and immune response. In the peripheral, domain-3 lacking isoform is significantly upregulated on activated T cells in rat, human PBMC as well as Jurkat cells

[97]. The reproducibility of these finding in different cell types suggests that the replacement of full-length CD6 with domain-3 lacking isoform upon following T cell activation could be a potential mechanism of regulating signals derived from CD6 ligand interaction.

CD6 is a risk gene for multiple sclerosis based on the single nucleotide polymorphism rs17828933 in the CD6 gene that was identified and confirmed as a risk factor for multiple sclerosis (MS) in different cohorts [98]. The T cells isolated from these patients preferentially express domain-3 lacking isoform over the full-length CD6.

The proliferation of these T cells is significantly impaired in response to anti-CD3/CD28 stimulation[99]. The observation that T cells from the “disease-risk” patient proliferated

less compared to T cells from “normal” patient is not entirely surprisingly given that CD6 lacking domain-3 is excluded from the immune synapse and thought to provide inhibitory signals during T cell activation.

Regulatory control of CD6 expression

The promoter region of CD6 on human T cells contains multiple transcription factors binding sites. The naive expression of CD6 on human T cells is dependent on the expression of the transcription factors RUNX1/3 and Ets1[100]. The cell surface expression of CD6 is enhanced during T cell activation indicating that CD6 may regulate

T cell immune response. In response to PMA stimulation, the CD6 surface expression on immature and mature T cells is greatly enhanced [101]. This observed increase in CD6 surface expression is dependent on protein kinase C (PKC) activation. The inhibition of

PMA-induced CD6 expression in the presence of cycloheximide demonstrated that the substantial increase of CD6 surface expression is dependent on new protein synthesis.

PMA had no effect on the CD6 surface expression on the mature B cell lines that were investigated [101]. The upregulation of CD6 expression on thymocytes has also been observed following in vitro stimulation with anti-CD2 mAb [88]. Whether this upregulation is also dependent on PKC signaling or CD6 mRNA transcriptional activity is unknown.

CD6 interacting proteins

The signaling pathway that is activated as a result of CD6 and its ligand interaction is not fully understood in T cells. Furthermore, until recently, whether CD6 is a positive or negative regulator of TCR signaling was very controversial. The

intracellular cytoplasmic domain of CD6 is composed of tyrosine, serine and threonine residues that can be phosphorylated. These tyrosine-residues are phosphorylated upon T cell receptor crosslinking [102]. The levels of phosphorylation can be augmented by crosslinking CD3 and CD4, indicating that CD6 may regulate TCR signaling transduction pathways that are important for T cell effector function [103]. However, the exact kinase that phosphorylates CD6 tyrosine residues is unknown. The rat homolog of CD6 co- precipitated with various kinases such as Lck, Fyn and ZAP70 [104]. The significance of this association was confirmed in later studies. The specific crosslinking of CD6 with three different mAbs (161.8, SPV-L14.2 and MAE1-C10) on normal and leukemic human T cells induced ERK, p38 and JNK activation [105]. The cytoplasmic domain of

CD6 and Src tyrosine kinase Lck and Fyn were required for these CD6-induced events.

In a later study, the phosphorylation of these intracellular molecules (ERK, p38 and JNK) in T cells was significantly reduced in the presence of the soluble anti-CD6 mAb

(Itolizumab). These data further supports the previous reports that the MAPK pathway is downstream of CD6-mediated signaling. Recently CD6 has been proposed to initiate its own T cell signaling pathway by recruiting the cytoplasmic adaptor protein SLP-76 and

Vav1[106].

On the cell membrane, CD6 also interacts with specific cell surface adhesion molecules. One of the extensively studied and well-characterized CD6 ligands is activated leukocytes adhesion molecule (ALCAM) also known as CD166 [107, 108].

CD166 is a member of the immunoglobulin superfamily proteins and was the first ligand identified for a scavenger receptor family protein. CD166 is broadly expressed on a variety of cells and tissues such as mononuclear cells, epithelial cells, stromal cells, bone

marrow progenitors, neurons and microglia in the brain[107, 109]. Through the membrane-proximal domain of CD6, CD6 interacts with domain 1 of ALCAM[107,

110].

In a recent study Galectin (Gal)-1and Galectin-3 were reported as two novel ligands for CD6 [11]. Gal-1/3 are the most abundantly expressed and extensively studied members of the galectin family. They are co-expressed on many components of the immune system including dendritic cells, macrophages, T and B cells as well as thymic epithelial cells (TEC) [17, 19-21, 111]. The binding of Gal-1/3 with CD6 appears to be dependent on the CD6 domain-3 because the binding of CD6 to CD166 is inhibited in the presence of Gal-1/3 [11]. From a functional perspective, WT CD6 transfectant 2G5 cells were significantly protected from Gal-1 induced apoptosis compared to parental 2G5 cells that do not express CD6. This CD6-mediated protection requires the cytoplasmic domain since 2G5 cells expressing CD6 cytoplasmic-truncated mutant were susceptible to apoptosis in the presence of high concentrations of Gal-1/3 [11]. Gal-3 hastens the de- adhesion of thymocytes to TEC suggesting that the binding of Gal-1/3 with CD6 could be of functional importance since the binding of CD6 to its other ligand CD166 may be important in thymocytes/TEC interaction during T cell development [17, 88].

CD6 role in T cell development and immune response

The report that ALCAM is expressed on thymic epithelial cells provided the first clue that CD6-derived signaling might be important in T cell development. However, the first study that suggested a role for CD6 in thymocyte survival and selection in mice and humans was based on the observation that CD2-ligation could selectively upregulate CD6 expression on thymocytes but not on mature T cells[101]. It was proposed that CD2-

induced CD6 expression might provide additional regulatory signals required for T cell differentiation. In a subsequent study, the role for CD6 in T cell development was further explored. The expression of CD6 is developmentally regulated in human and mouse thymocytes. The expression level of CD6 correlates with the positive selection marker

CD69 [88]. CD6 expression increases as double-positive thymocytes are selected to single positive CD4+ and CD8+ T cells. Additionally, CD6 is expressed on immature thymocytes that are resistant to apoptosis[88]. Collectively, these findings suggested that

CD6-derived signals might be involved in thymic survival and functional avidity of selection.

All of the preceding studies support a putative involvement of CD6 in T cell development. However, the lack of CD6-deficient mice delayed further understanding of its role in vivo during T cell development. Recently the work by Lozano and co-workers provided the first in vivo analysis of CD6-mediated function in T cell development, homeostasis, activation and immunological response to self-antigens on the C57BL/6 background[112]. The deficiency of CD6 had a significant impact on normal T cell development by triggering an increase in the negative selection of single positive CD4 and CD8 T cells during the transition from double positive thymocytes T cells to single positive mature T cells[112]. As a result of this increased negative selection, the frequency of single positive CD4 and CD8 T cells are reduced in the thymus. The analysis of peripheral tissues revealed an increased frequency of effector/memory T cells in the spleen and lymph node of CD6-deficient mice indicating a role for CD6 in the restriction of antigen-experienced peripheral T cells [112].

Prior to the deciphering of CD6 ultimate function in T cells, in vitro experiments using soluble CD6 or CD6 blocking mAb as well as engineered cells overexpressing CD6 were employed to determine the function of CD6. Not entirely surprising, the results from these in vitro studies were contradictory. Several studies reported that soluble CD6 or recombinant ALCAM-Fc could significantly reduce T cell activation and proliferation.

Consistent with this finding, antigen induced T cell proliferation and IL-2 cytokine secretion were also inhibited by recombinant CD6 and ALCAM [95, 113]. These reduced

T cell responses in the presence of soluble CD6 and recombinant ALCAM highlights the important role of CD6-ALCAM interaction in T cell activation. Consistent with a co- stimulatory role, anti-CD6 monoclonal antibodies also significantly impaired T cell proliferation and cytokine secretion[114, 115]. Interestingly, the CD6 mAbs used in these studies specifically binds to domain-1 of CD6 and do not block ALCAM binding to

CD6. Collectively, these in vitro data suggest a co-stimulatory role for CD6 in T cell activation and further supports the results from the clinical studies in which anti-CD6 mAb was utilized.

The belief that CD6 is a co-stimulatory has been contested by several in vitro studies, which indicated that CD6 might be a negative regulator of T cell activation. The overexpression of wild-type or rat CD6 molecule significantly reduced CD3 mAb induced calcium influx in primary human T cells or Jurkat T cells[94]. In a later study,

E.6.1 T cells that do not express CD6 naturally were observed to secrete more IL-2 compared to CD6-positive T cells [94]. Additionally, these CD6-negative T cells also displayed elevated calcium influx compared to CD6-positive T cells. This enhanced mobilization of calcium influx in CD6-negative T cells was also observed in T cells

overexpressing CD6 cytoplasmic truncation mutant [102]. Finally, after more than 30 years since the identification of CD6, the Lozano group recently published the first study addressing the role of CD6 in TCR signaling using primary T cells isolated from CD6- deficient mice. Their TCR signaling experiments using CD6-deficient T cells supported a role for CD6 as a negative regulator of T cell activation. The analysis of TCR-induced

Calcium mobilization showed that CD6-deficient double positive T cells display elevated calcium-influx compared to WT double positive T cells [112]. Subsequent analysis indicated that this elevated TCR-induced calcium influx was undetectable in single CD4 and CD8 positive thymocyte[112]. Interestingly analysis of in vitro CD3-mediated proliferation of peripheral single positive CD4 and CD8 T cells showed an increased proliferation in CD6-deficient T cells [112]. Although this study provided a great understanding of CD6 in vivo role in TCR signaling, it failed to address the effect of

CD6-deficiency on CD6 activated pathways that have been previously reported.

Additionally, the molecular mechanism by which CD6 curtails TCR signaling remains speculative. There are two potential mechanisms by which CD6 could restrict TCR signaling that could be explored in future studies. (1) It is possible that CD6 could recruit tyrosine phosphatases to the TCR signalosome to restrain the amplitude of TCR signaling. (2) Alternatively, CD6 could transduce inhibitory signals through its association with CD5 given the previous report that these two molecules interact at the T cell surface.

Nevertheless, the current belief is that CD6 is a negative regulator of TCR signaling. The existence of CD6-deficient mice presents an exciting opportunity to fully understand the mechanisms by which CD6 enact its regulatory functions in T cells.

CD6 role in B cells

In contrast to the extensive studies addressing the role of CD6 on T cells, there are only a few studies addressing the potential role of CD6 on B cells. CD6 is selectively expressed on mature, memory and B1a B cells in humans[89]. The frequency of CD6+ B cells in the peripheral blood of patients Sjogren Syndrome (SS) is reduced compared to patients with rheumatoid arthritis [89]. This observed decrease in the peripheral blood is a result of the accumulation of CD6+ B cells in the salivary gland [89]. This finding led to the hypothesis that perhaps CD6 was facilitating the transmigration of CD6-expressing B cells into the salivary gland and thereby contributing to the pathogenesis of SS. In support of this hypothesis, the salivary gland epithelial cells were reported to express the

CD6 ligand ALCAM.

In addition to human PMBCs, CD6 is expressed on chronic lymphocytic leukemia

B cells (B-CLL), which are derived from B1a precursor cells. An analysis of CD6 expression on 26 leukemic cells from progressive and non-progressive B-CLL patients did not reveal any significant difference in the expression level of CD6 between the groups [116]. The CD6 expression on B-CLL increases in response to staphylococcus aureus Cowan or TPA stimulation. Furthermore, CD6 is not involved in B-CLL proliferation but integrally important in their protection from anti-IgM induced apoptosis by modulating Bcl2/BAX-2 ratio [116]. The expression of CD6 has also been analyzed on two established B cells originating from patients with Burkitt lymphoma (Daudi and

Raji) and no expression was detected in these cells [101]. Although CD6 is expressed on different B cell subsets in human peripheral blood, the B cell immunological function of

CD6 and expression profile on mouse B cell subsets remains unknown.

The observation that CD6 is primarily expressed on T cells and potentially has a dual role as a modulator of intracellular signaling has made it a therapeutic target using mAb in several diseases such as multiple sclerosis, psoriasis, rheumatoid arthritis and attaining several levels of success. Initially, the rationale for targeting CD6 in early clinical studies was to deplete T cells given that CD6 is primarily expressed on T cells that play a pathogenic role in several autoimmune diseases [117]. However, recent studies targeting CD6 that has achieved great success utilizes a non-T cell depleting CD6 mAb [114, 118, 119].

Introduction to IOR-T1 (murine derived anti-human CD6 mAb)

The IOR-T1 is a murine anti-human CD6 mAb that was developed in the 1980s in

Cuba for the immunologic diagnosis, prognosis and treatment of patients diagnosed with leukemia and lymphomas[120, 121]. This mAb was generated in BALB/c mice immunized with the PBMC isolated from a patient diagnosed with Sezary’s syndrome

[122, 123]. The major rationale for the utilization of IOR-T1 in an autoimmune clinical setting particular psoriasis and rheumatoid arthritis (RA) was based on early studies in which depletion of CD6-positive T cells by IOR-T1 from donor bone marrow prevented the graft-versus-host disease[124]. In psoriasis, the application of IOR-T1 topically in combination with IV infusion showed good results in long-lasting psoriasis vulgaris condition[125]. In RA patients, IOR-T1 mAb reduced the number of tender and swollen joints[126].

Introduction to Itolizumab (humanized CD6 mAb)

The clinical results following treatment with IOR-T1 were very promising, however there several undesirable side effects necessitating the need to humanized the

CD6 mAb. In many of these patients, human antibodies directed towards the infused mouse IOR-T1 mAb were detected in the serum and the median half-life of the antibody ranged from 13.9 to 19.6 h post-infusion[127, 128]. The humanized mAb was termed

Itolizumab and reported to be less immunogenic, while also displaying similar affinity as the parental murine IOR-T1 mAb[129]. Itolizumab displayed a longer half-life in the serum following infusion and this was attributed to the rate at which the Fab fragment of

Itolizumab disassociates[130]. The Fab fragment of the humanized mAb dissociates six- fold lower than IOR-T1, resulting in a longer half-life and as a result, patients were infused less frequently compared to the parental IOR-T1 mAb[130].

RA is a chronic inflammatory disease that affects about 0.5- 1% of the adult population in the western hemisphere with an estimated 3-fold greater incidence in females resulting in impaired movement, deterioration of life and lower life expectation[131, 132]. The hallmark of this disease is the chronic inflammation of synovial tissue in the joints that eventually leads to the destruction of bone and cartilage.

Although the exact early events that triggers RA is unknown. There is strong evidence that T cells play an important role in the pathogenesis of RA[132]. The depletion of T cells using specific antibodies has been reported to reduce disease severity in the collagen-induced arthritis models in rodents[90, 133, 134]. Additionally, the proinflammatory cytokines secreted by T cells such, as IFN-gamma and IL-17 are associated with synovial inflammation[7, 132].

CD6 as a therapeutic target in RA clinical trials

Itolizumab is currently approved for the treatment of psoriasis in India following the clinical data from two large randomized multicenter studies in both Cuba and India that were carried out to assess its effect on psoriasis. The clinical efficacy of Itolizumab in patients with psoriasis led to clinical studies that evaluated its effect on patients with rheumatoid arthritis. In the first open-label, and dose-finding phase 1 trial with 18 patients with rheumatoid arthritis, the anti-CD6 IOR-T1 mAb showed great potency of therapeutic immunosuppression[92]. The patients received three different daily doses of

0.2 mg/kg, 0.4 mg/kg or 0.8 mg/kg of IOR-T1 mAb via IV infusion and the therapeutic effects were immediate. The patients within four days of receiving the infusion reported significant improvement in their tender joints and swollen joints. There was a dose- dependent effect of IOR-T1 that was observed following infusion. Contrary to the expectation of the investigators, patients that received low doses of IOR-T1 reported greater benefits than those receiving the highest dosage. The patients receiving the 0.4 mg/kg dose showed long-lasting clinical improvement, which resulted in the selection of this dosage as the Optimum Biological Dose[92]. Adverse events correlating with the dosage of IOR-T1 were observed and could be controlled by medication.

The immediate and long-lasting clinical improvement observed with the parental murine IOR-T1 led to the subsequent clinical studies to evaluate the safety, immunogenicity and preliminary efficacy of the humanized anti-CD6 mAb (Itolizumab) in 15 patients with active rheumatoid arthritis in a phase 1, open-label and dose-finding study[92]. The majority of these patients were women (73%) with a median duration of the disease of 10 years. These patients had previously failed to response adequately to

disease-modifying therapies (DMTs). The patients received a weekly infusion of itolizumab ranging from 0.1 to 0.8 mg/kg for a period of 6 weeks. The objective clinical response as assessed by the ACR (American College of Rheumatology criteria) score was achieved in more than 80% of the patients and these responses were sustained for a period up to 6 months after the last infusion. There was no observed anti-idiotypic response in the serum of these patients, which correlated with the lack of adverse events compared to the parental murine IOR-T1 mAb. Additionally, Itolizumab infusion did not alter the lymphocyte population count during the course of the study. This finding is consistent with several in vitro experiments that indicated Itolizumab does not induce T cell depletion by complement-dependent cytotoxicity or antibody dependent cell- mediated cytotoxicity but rather induces CD6 internalization in T cells within minutes[97]. Collectively, these studies suggest an alternate mechanism of action that is independent of CD6+ lymphocyte depletion.

In a later clinical study, the safety and efficacy Itolizumab in combination with methotrexate was assessed in patients with active RA in an open-label, phase 2 study[2].

The patients received three different doses of 0.2, 0.4 or 0.8 mg/kg of Itolizumab on a weekly basis for a 6-month time period. At the end of 12 weeks, 50% of the patients

achieved ACR20. The Itolizumab infusion was well tolerated and no anti-Itolizumab idiotype antibodies were detected in the serum. The OBD was unable to be defined properly due to the low number of participants in the study. Nevertheless, this study provided further evidence for the therapeutic benefits of Itolizumab in patients with RA.

Although results from the clinical studies evaluating the efficacy of Itolizumab in the treatment of RA were promising, there were several limitations to the study design

that preclude definite conclusions from being ascertained. One of the major concerns is the small number of patients that were enrolled in the clinical studies. The open-label nature of the Cuban studies may not be as rigorous as open-label clinical studies in the

USA. Additionally, the lack of placebo-control arm group linked with open-label design potentially may result in the overestimation of Itolizumab treatment effect. The recent thaw in USA-CUBA relations has spurred a lot clinical trial related collaboration between the two countries. The developers of Itolizumab are planning on filing an investigational new drug application with the FDA upon the identification of a suitable partner with the expertise and funding needed for the approval in the U.S for the treatment of psoriasis.

They are optimistic that their high quality data from the clinical studies in Cuba and India would shorten the application process in the U.S.

IOR-T1/ Itolizumab potential mechanism of action

One of the major issues with patients receiving DMTs is immunosuppression and increased risks of infection. In RA patients that received anti-CD6 mAb treatment, the effect of targeting CD6 on the immune system was investigated given that CD6 is primarily expressed on T cells and plays an important role in T cell immune response.

IOR-T1 infusion did not alter the PBMC lymphocyte count, which correlated with the absence of opportunistic infection during the clinical trials[2, 119]. In a 52-week study examining the effect of Itolizumab on psoriasis, the immune cell population in the peripheral blood was also not affected[102]. Regarding its effect on immunosuppression, many participants in the study had urinary tract infection and the common cold. These infections were mild or moderate and in majority of the cases, these infections were

considered not related to the assayed drug[102]. Nevertheless, in comparison to other biological agents, Itolizumab appears to have reduced incidence of infection.

The lack of lymphocyte depletion in the peripheral blood of patients infused with

Itolizumab provided some clues to its potential immunomodulatory mechanism. The use of in vitro binding and blocking assay mapped the binding domain of Itolizumab to the membrane distal domain 1 of CD6[129, 130]. The binding of Itolizumab to CD6 did not interfere with the binding of soluble CD166 to CD6 domain 3. In a later study, T cell proliferation and inflammatory cytokine secretion such as IL-6, IFN-γ and TNF-α were impaired in the presence of Itolizumab following stimulation of PBMCs from healthy with anti-CD3 mAb and soluble CD166[115]. The activation of intracellular signaling proteins such as MAPK, STAT3 and AKT were also impaired in the presence of

Itolizumab[105, 135]. These signaling pathways are also activated following CD6- crosslinking. The dose-dependent inhibition of Itolizumab was also associated with reduced CD25 expression [115].

These in vitro inhibitory effects of Itolizumab are consistent with the observation made in clinical studies targeting CD6 in patients with psoriasis that received Itolizumab infusion[99]. Isolated T cells from these patients, when restimulated in vitro, displayed reduced numbers of IFN-γ secreting T cells and also reduced T cell proliferation capacity. Additionally, the analysis of the serum from these patients also showed reduced levels of IL-6, TNF-α and IFN-γ. Furthermore, the analysis of T cell infiltration in the epidermal layer of these psoriatic patients from skin biopsies showed reduced CD3+ and

CD6+ T cells[99]. This reduction in T cell infiltration correlated with significantly reduced epidermal thickness.

The recently deciphered CD6 role as a negative regulator suggests that perhaps

Itolizumab binds to domain -1 of CD6 and induce inhibitory signals that modulate the intensity of T cell immune response. Alternatively, Itolizumab may inhibit the interaction of a novel ligand of CD6 that binds to the membrane distal domain 1 of CD6. Although

Itolizumab has been postulated to be a non-blocking mAb, this assumption was derived from in vitro blocking experiments showing that Itolizumab does not inhibit the binding of soluble CD166 to domain-3 of CD6. Further indicating that the potential mechanism of

Itolizumab inhibiting the binding of a novel a CD6 ligand to domain 1 of CD6 is a plausible hypothesis.

Evidence for additional CD6 ligands

Several studies have suggested the existence of additional CD6 ligands other than

ALCAM including 1) in vitro blocking studies using anti-ALCAM antibodies only partially inhibit the binding of ALCAM-expressing thymic epithelial cells to CD6 overexpressing COS cells[103], 2) an anti-ALCAM mAb failed to completely inhibit

CD6- dependent binding of Hut 78 cells to IFN-γ-stimulated keratinocytes[104], and 3)

CD6 protein immunoprecipitated 3 polypeptides of surface proteins from synovial fibroblast lysate including a 130-kDa band that is distinct from ALCAM[107, 136].

Additionally, several studies also suggest that the expression of this other ligand(s) is upregulated after IFN-γ stimulation[107]. These earlier results initiated the quest to identify and characterize these other CD6 ligands.

In a subsequent study, a mAb 3A11 recognizing this 130-kDa protein that could be a new ligand of CD6 was developed[136]. This mAb was developed by serially immunizing BALB/c mice intraperitoneally using HBL-100 cells. The hybridoma

supernatants were screened for differential reactivity to resting and IFN-γ stimulated epithelial cells by flow cytometry. The antigen recognized by mAb 3A11 is expressed in human thymus, skin, synovium and cartilage and its expression is enhanced by interferon-γ stimulation. The 3A11 antigen can be immunoprecipitated using either mAb

3A11 or CD6 protein indicating that the 3A11 protein is a CD6 ligand[136].

Additionally, siRNA knockdown of ALCAM expression did not alter mAb 3A11 antigen expression, therefore confirming that the mAb 3A11 antigen is distinct from the previously identified CD6 ligand, ALCAM. Although 3A11 and CD166 are two distinct proteins, they both colocalize at the cell surface[136].

Introduction to CD318

Although the antigen recognized by mAb 3A11 has been proposed as a distinct

CD6 ligand, the identity of this protein was unknown. We identified CD318 also referred to as TRASK, SIMA135 and gp140 as the 3A11 antigen by Mass spectrometry. CD318 is a 135-150 kDa cell surface glycoprotein that is highly tyrosine-phosphorylated and overexpressed in several cancers[110, 137-139]. Structurally, it has 3 extracellular CUB

(complement protein subcomponents C1r/C1s, urchin embryonic growth factor and bone morphogenetic protein 1) domains termed CUB1, CUB2 and CUB3, a transmembrane domain and a cytoplasmic domain[110, 137, 138, 140]. The intracellular domain of

CD318 contains several tyrosine residues that are phosphorylated upon CD318 CUB1 crosslinking, mediated detachment from the extracellular matrix or protease cleavage of the distal extracellular CUB domain[117, 135]. The intracellular cytoplasmic domain also contains SH3-containing proteins binding residues. The protease cleavage of CD318 results in the generation of a 70-85 kDa membrane-anchored form of CD318[141-143].

The naive expression of either full-length (FL) or cleaved form of CD318 is dependent on the cell type [144]. The majority of the cells preferentially expresses the FL CD318 compared to the cleaved form. Following cleavage of FL CD318, several studies have reported several soluble fragments ranging from 25-110 kDa product sizes[145].

CD318 expression profile and regulatory control

CD318 is highly expressed in ovarian, lung, kidney, gastric and colon cancers[137, 139, 140, 145, 146]. Elevated expression of CD318 by immunostaining correlates with poor prognosis in patients with pancreatic, lung, ovarian and kidney cancer[147, 148]. In non-cancer cells, CD318 is abundantly expressed on non-immune cells such as mesenchymal stem cells, neuronal progenitor cells, hematopoietic stem cells and various cancers of epithelial and stromal origin in humans[149-151]. CD318 is not expressed on B cells, T cells or myeloid cells in humans[152]. The cell-type restricted expression of CD318 is dependent on the regulated interaction between transcription factors and chromatin DNA. Lymphocytes, granulocytes and Jurkat cells have transcription factors that can induce CD318 expression, however there are several heavily methylated CpG sequences around the promoter region of CD318. The demethylation of these CpG sequences results in the expression of CD318 in lymphocytes[153]. The

CD318 expression on human colorectal carcinoma cells can be enhanced by HIF-1/2α transcription activity in response to hypoxia[154-156]. Oncogenic Ras/ERK signaling also regulates CD318 expression. The expression of CD318 was observed to be significantly elevated in non-small cell lung cancer (NSCLC) cell lines with Ras mutations[120]. In support of this finding, CD318 promoter contains three activator protein-1 (AP-1) binding sites. Recently the ADAM9, a transmembrane protein of the

ADAM family has been reported to enhance CD318 expression by suppressing miR-218 expression[121].

CD318 function in cancer is dependent on its tyrosine phosphorylation

CD318 cytoplasmic domain contains five tyrosine residues sites (Tyr 707, 734,

743, 762 and 806) that are phosphorylated by Src family kinases (SFK) including Src,

Fyn and Yes. Tyr 734 is the major tyrosine phosphorylation site of CD318 and is critically important for CD318 mediated function[157]. The phosphorylation of CD318 at

Tyr 734 by SFK results in the binding of SFK to this site, which also results in the tyrosine phosphorylation at Tyr 743 and Tyr 762[157]. Both the FL and cleaved CD318 are tyrosine phosphorylated by SFKs. Following Tyr 762 phosphorylation, Protein

Kinase C δ is recruited to this site and subsequently activated. CD318 induced PKC δ and protein kinase B (Akt) activation, has been associated with cancer cell anoikis resistance and cancer cell migration[158]. CD318 can also facilitate cancer cell

migration/invasion through a variety of signaling pathways that include EGFR and β1 integrin signaling[159, 160].

CD318 was first identified in several NSCLC cell lines in suspension because it was highly tyrosine-phosphorylated compared to other proteins[137, 138]. This observed phosphorylation was attributed to the loss of integrin binding to the extracellular matrix.

In a later study, the inhibition of α6β4 mediated keratinocytes adhesion to laminin 5 using inhibitors increased the cytoplasmic tyrosine phosphorylation of CD318[161].

Furthermore, keratinocytes that were treated with trypsin predominantly expressed the cleaved membrane-anchored 80-kDa tyrosine phosphorylated CD318[161]. Proteolytic cleavage of CD318 has also been reported to induce cytoplasmic tyrosine

phosphorylation of a 70-kDa membrane-anchored CD318 and the subsequent recruitment of Src and PKC δ to the cytoplasmic domain[162]. Dephosphorylation of the CD318 cytoplasmic domain has also been reported to occur following reattachment of these cells to the extracellular matrix[143]. The transition between attachment and detachment from the extracellular matrix plays an important role in cancer resistance to anoikis, cell migration and invasion. The switch of CD318 cytoplasmic tyrosine phosphorylation and dephosphorylation suggest that regulation of CD318 phosphorylation state might play an important role in cancer cell anoikis, migration and invasion.

The significance of CD318 tyrosine phosphorylation in cancer migration and metastasis was first explored in the highly invasive scirrhous gastric cancer[163]. The specific knockdown of CD318 did not affect the survival and proliferation, however it significantly impaired the invasion and dissemination of these cancer cells to the liver and peritoneal tissues when orthotopically implanted in nude mice[163]. In a later study, the over expression of CD318 on Hela cells conferred these non-aggressive cells with higher metastatic and invasive properties[164]. The suppression of CD318 expression by siRNA resulted in reduced pancreatic cell migration and invasion. In the same study, the tyrosine-phosphorylation of CD318 was implicated in pancreatic cell migration and invasion. The overexpression of a CD318 mutant (Tyr734F) reduced pancreatic survival, migration and invasion[164]. In a different study, the shRNA knockdown of CD318 expression in renal carcinoma cells reduced the migration and invasion of these cells[155]. The impairment of PKC activation in the absence of CD318 or presence of

CD318 mutant was attributed to the observed reduction in cell migration and invasion in these different studies[155]. Interestingly in CD318-deficient mice, the loss of CD318

resulted in the acceleration of mammary and skin tumors driven by the PyMT and

SmoM2 oncogene respectively confirming a tumor-suppressing function[165].

CD318 interacting proteins

The structural features of CD318 make it an ideal candidate that is capable of initiating its own signalosome in response to CD318 ligation/crosslinking. Interestingly, the specific ligand(s) that binds to CD318 and induces its activation/tyrosine phosphorylation is currently unknown. Proteomic analysis of CD318 containing complexes reveals that CD318 interacts with cell surface molecules that are involved in cell-cell, cell-matrix adhesion such as CD9, Syndecan-1 and Syndecan-4, N and P-

Cadherin, metalloproteinase 14 (MMP-14) and Galectin 1[159, 166, 167]. Galectin 1 selectively interacts with FL CD318 specifically the carbohydrate moieties containing N- terminal domain of CD318[159]. However, whether the association of these molecules induces or influence CD318-dependent signaling remains to be determined. CD318 has also been reported to interact with several cell surface receptors. For example, the EGFR receptor interacts with CD318 to form a multi-protein complex that includes Src that induces cell detachment and disrupt breast cancer cell adherens junctions[159]. Recently

CD318 was reported to interact with HER2 via its intracellular domain and promotes

HER2 driven tumorigenesis in breast cancer[168]. Crosslinking of the CD318 N- terminal domains using a CUB1 mAb can initiate CD318-dependent signaling. The

CUB1 mAb-mediated crosslinking promotes Src-induced CD318 Tyr 734 phosphorylation, which is essential for CD318 mediated function[117]. The identification of a surface molecule that can initiate CD318 intracellular signaling upon binding is

critical in understanding the biological function of this protein in normal epithelial and non-epithelial expressing cells.

Therapeutic potential of targeting CD318

Circulating levels of soluble CD318 ranging from 25-110 kDa in molecular weight has been detected in the urine of patients with prostate cancer. Using quantitative mass spectrometry, the level of soluble CD318 was demonstrated to be significantly elevated in high-risk prostate cancer patients compared to low-risk patients [145]. This observation highlights the therapeutic potential of targeting CD318. In a later study, Siva and colleagues investigated the therapeutic potential of targeting CD318 in tumor migration using the murine anti-CD318 antibody 25A11. This antibody inhibited tumor migration and invasion in vitro[169]. In a mouse xenograft model, an anti-CD318 mAb termed C20Fc efficiently blocked primary tumor growth and metastasis of PC-3 cells[170]. These finding highlight the therapeutic potential of anti-CD318 mAb in

CD318-mediated diseases such as cancer.

Summary and Aims

CD6 is a cell surface glycoprotein that is expressed on all T cells and a subset of

B cells in humans. CD6 is a negative regulator of TCR signaling and critically important in peripheral T cell homeostasis. In B cells, the biological function of CD6 is currently unknown. The finding that CD6 is an important modulator of T cell immune response has made it a therapeutic target in several autoimmune-related clinical studies. Itolizumab, a humanized mAb targeting CD6 was recently approved for the treatment of chronic plaque psoriasis in India. Itolizumab binds to domain-1 of CD6 indicating that it does not

interfere/inhibit the binding of CD6 and ALCAM suggesting the presence of a novel ligand that binds to domain 1 of CD6. Additionally, several in vitro blocking and immunoprecipitation studies suggest the existence of additional CD6 ligands other than

ALCAM. The antigen recognized by the 3A11 mAb is a novel CD6 ligand that is distinct from ALCAM. However, the identity of this antigen is unknown. We identified

CD318 as the antigen recognized by the 3A11 mAb by Mass Spectrometry. CD318 is a

135-150 kDa cell surface glycoprotein that is primarily expressed on epithelial cells. The role of CD318 has been extensively studied on cancer cells and its expression is associated with poorer prognosis. The knockdown of CD318 inhibits cancer cell migration and invasion, which is dependent on CD318 tyrosine phosphorylation.

Although CD318 has been extensively studied in cancer, its role in immune response is unknown.

This thesis has two major aims.

Aim 1: To understand the role of CD6 on mouse B1a cells and its contribution to the pathogenesis of intestinal ischemia reperfusion-induced injury. For this aim, using

WT and CD6-/- mice, we studied the potential role of CD6 in regulating intestinal I/R- induced injury by comparing mucosal histopathology, local IL-6 production, and serum

IgM titers. We explored the underlying mechanism by examining the distribution, regulation, and effect of CD6 on B1a cells.

Aim 2: To determine the identity of 3A11 antigen and confirm whether it’s a CD6 ligand. For this aim, using mass spectrum techniques, we identified the antigen recognized by mAb 3A11 as CD318. To validate the proteinomics results, we probed proteins pulled down by mAb 3A11 using established anti-CD318 antibodies and probed

recombinant CD318 protein by mAb 3A11 in Western blots. We also compared staining patterns of both mAb 3A11 and established anti-CD318 mAbs using cells that known to be positive or negative for CD318, and engineered cells with upregulated or downregulated levels of CD318. In addition, we confirmed the binding of CD318 to CD6 by using soluble CD6 protein as bait in pull-down assays and by staining WT and CD166 deficient cells with the soluble CD6 followed by flow cytometric analyses. Finally, we examined CD318 expression on synovial fibroblasts from rheumatoid arthritis patients, measured levels of soluble CD318 in synovial fluids from RA patients, and studied its potential roles in recruitment and retention of T cells in synovial tissue.

CHAPTER 2:

Materials and Methods

Methods:

Mice: CD6-/- mice were developed by manipulating ES cells using a conventional homologous recombination strategy and blastocyst injection of the identified target ES cells. The resultant CD6-/- mice were bred onto the DBA/1 background by more than 12 generations of backcrossing. PCR and Southern blot assays verified the deletion of the targeted loci, and flow cytometry analysis of CD6 protein on T cells confirmed the deficiency of CD6 in the CD6-/- mice. All mice were maintained in the animal facility at the Cleveland Clinic and all animal care and experimental procedures were approved by the Institutional Animal Care and Use Committee of the Cleveland Clinic.

Mouse intestinal I/R model: The intestine ischemia and reperfusion was performed on mice at 8-12 weeks of age. Surgical instruments were sterilized by autoclave prior to surgery. The mice were anesthetized with 4% isoflurane inhalation for induction and

0.5% for maintenance of anesthesia using an approved anesthetic gas machine. The mouse abdomen skin was cleaned and entered via a midline laparotomy incision. The superior mesenteric artery was identified and isolated, and then the VASCU-STATT Plus bulldog clamp (Scanlan, Saint Paul, MN) was utilized to perform the vascular exclusion for 30 min. A heating pad was applied to keep the mouse body temperature around 37oC,

After 30 min of mesenteric ischemia phase, the clamp was removed, and the midline laparotomy incisions were closed with 4-0 silk suture, the intestine was reperfused for 3 hours before the tissue (jejunum) was harvested. For some experiments, CD6-/- mice were reconstituted with 100 ug of mouse IgM (Rockland, PA) intravenously 30 minutes prior to the start of the intestinal ischemia-surgery

Local IL-6 assay: The jejunum tissue was mechanically homogenized in NP-40 lysis buffer (Invitrogen) containing a tablet of protease inhibitors (Roche). The homogenized tissue was incubated on ice for 30 min, with brief vortexing every 5 min. Tissue lysates were centrifuged at 13000 rpm for 30 min at 4 °C and pellets were discarded and protein concentration of the supernatant was measured using the Pierce BCA Protein Assay Kit.

For the detection of IL-6 levels in the supernatant, 96 well high binding plates were coated with 2 μg/mL of rat anti-mouse IL-6 (Biolegend, CA) diluted in 1X PBS. Protein lysates were diluted 1/20 and loaded onto the coated ELISA plate. Bound IL-6 were detection using biotin anti-mouse IL-6 (Biolegend, CA) and subsequently using avidin-

HRP (Biolegend, CA). The ELISA color reaction was initiated using TMB substrate

(Thermo Scientific, MA). 2M H2SO4 was used to stop the TMB reaction and absorbance at 450 nm was measured. IL-6 concentration was normalized to the starting initial protein concentration.

Histology and injury scoring: Histopathological examination and tissue injury score was performed in a blinded manner by a pathologist on 4-μm-thick paraffin-embedded sections stained with hematoxylin and eosin. Digital images were captured with a Leica-

DM5500 B microscope camera (Buffalo Grove, IL) and analyzed with Image Pro software (Media Cybernectics, Silver Spring, MD).

Flow cytometry: Single-cell suspensions of spleen, peritoneal lavage from adult mice or day 7 old neonatal liver were incubated for 10 min on ice with anti-mouse CD16/32 Fc blocking antibody (Biolegend, CA). Following Fc blocking, cells were incubated with varying combinations of the following antibodies: anti-CD19, anti-CD43, anti-CD5, anti-

CD6 clone 34 (prepared in the lab), anti-CD11b, anti-CD80, anti-CD86, anti-IgM, anti-

CD23 and anti-CD21. All antibodies were purchased from Biolegend. Cells were stained in FACS buffer and expression of cell surface markers was acquired on both BD

LSRFortessa and FACSCalibur flow cytometer. Acquired flow cytometer data was analyzed using the Flowjo software. For splenic B1a intracellular staining, splenocytes from WT and CD6-/- mice were first incubated with antibodies to CD19, CD43 and CD5 as previously described. Following cell surface staining, the cells were fixed and permeabilized using the BD permeabilization kit according to manufacturers instruction.

The permeabilized splenocytes were incubated with anti-pERK (T204/Y202)

(eBioscience, CA), NFATc1 (Biolegend, CA), mouse IgG1 isotype control (BD

Bioscience, CA) or rat isotype control antibody (Biolegend, CA).

Splenic B2 cells in vitro stimulation: Single-cell suspensions of spleen were plated at a density of 106 cells/ml in a 96-well plate. For the detection of CD6 expression on splenic

B2 cells, splenocytes from WT and CD6-/- mice were harvested after 72 h in vitro

stimulation with 10 μg/ml of goat F(ab’)2 anti-IgM and IL-4 (1000 Units/mL)

(Peprotech, NJ) or 5μg/ml of LPS (Sigma, MO) and CD6 expression was analyzed on activated CD19+ (B2 cells) by flow cytometry.

Peritoneal B1a cells migration to the spleen: Peritoneal cells were isolated from WT and

CD6-/- mice by peritoneal lavage. 5 x 106 cells were labeled in vitro with CFSE

(Invitrogen, CA) and transferred into the peritoneal cavity of CD6-/- recipient mice. Two hours post-adoptive transfer; 5 μg of LPS (Sigma, MO) was also administered into the peritoneal cavity. After 72 h, splenocytes were isolated and CD6 expression was analyzed on CFSE positive cells in addition to gating on CD19 and CD5 double positive cells

(CFSE+CD19+CD5+).

BrdU incorporation: WT and CD6-/- mice were intraperitoneally injected with 500 μl of

BrdU (BD biosciences, CA) at a diluted concentration of 1 mg/ml every other day for 7 days. At the end of the study, splenocytes, Bone marrow and peritoneal cavity cells were isolated and stained with various antibodies (CD19, CD43, CD5), (CD19, CD43), and

(B220, CD11b, CD5) respectively. After cell surface staining on ice, the cells were fixed and permeabilized. Following permeabilization, the cells were intracellular stained with

FITC-labeled anti-BrdU antibody (BD biosciences, CA) and analyzed by flow cytometry.

ELISA: For the detection of total IgG or IgM serum levels, 96 well high binding plates

(Greiner bio-one, NC) were coated with 0.34 μg/ml of rabbit anti-mouse IgM+IgG (H+L)

(Jackson Immunoresearch, PA). Serum samples were diluted 1/1000. Bound antibodies were detected using either goat anti-mouse IgM HRP (Southern biotech) or rat anti- mouse IgG-HRP (Southern biotech, AL). For the detection of PC-specific IgM, plates were coated with 10 μg/mL of PC-BSA (Biosearch technologies, CA). Serum samples were also diluted 1/1000 and bound PC-specific IgM antibodies were detected using goat anti-mouse IgM-HRP. For the detection of Annexin IV-specific IgM, plates were coated with 1ug/mL of recombinant mouse Annexin IV (LSBio, WA). Serum samples were diluted 1/50 and bound Annexin IV-specific IgM were detected using goat anti-mouse

IgM-HRP. The ELISA color reaction was initiated using TMB substrate (Thermo

Scientific, MA). 2M H2SO4 was used to stop the TMB reaction and absorbance at 450 nm was measured.

Statistical analysis: Data were compared by unpaired t test or paired t test using the

GraphPad Prism software with a two-tailed Student’s t test of equal variance. The difference between the groups was considered significant at p<0.05.

Cell culture

The HBL-100, Raji, A549, Molt4 and MCF, wild type (WT) HT-1080 and CD166 knockout (KO) cell lines were cultured in RPMI supplemented with 10% FBS, L- glutamine, penicillin/streptomycin and Na-pyruvate. WT MDA-468, and CD318 knockdown cell lines as well as transfected CHO cells expressing human CD6 on their surface[171] were cultured in DMEM supplemented with 10% FBS, L-glutamine, penicillin/streptomycin, Na-pyruvate and 300 μg/ml of G418. MDA-468 expressing empty vector or doxycycline-inducible CD318 were also cultured in the same media described above with Zeocin in place of G418. Caco-2 cells were also cultured in the same media described in the absence of selection pressure. MDA-468 expressing vector control and doxycycline inducible CDCP1 were stimulated with 100 ng/ml of doxycycline overnight[172].

CD166 knockout cell line development

CD166 was knocked out in the HT-1080 cells using CRISPR/Cas 9 technology.

In brief, RNA (AGACGGTGGCGGAGATCAAG, Horizon Discovery, UK) was transfected into cells by lippofection. Transfection efficiency above 20% is considered successful. Efficiency was monitored by the surrogate marker, GFP. Cells lacking cell- surface CD166 were selected by fluorescence activated cell sorting using an antibody against the extracellular domain of CD166 (R&D Systems, MAB656). Cell population was further purified by single-cell colony formation in soft agar. After identification of pure, CD166-negative clonal populations, 5 clones were selected at random and mixed to create the CD166-KO cell line.

Antibodies

3A11 mAb was previously developed and characterized [136] [173]. mAbs against human CD166, human CD318 (clone: CUB1) and mouse IgG2b isotype control were all obtained from ( Biolegend, San Diego, CA). The polyclonal anti-CD318 antibody was obtained from Thermo Scientific, (Waltham, MA). Recombinant human

CD6 was obtained from R&D,(Minneapolis, MN). Recombinant mouse CD6 was described previously [173]. Purified human IgG1 was obtained from Sigma-Aldrich, (St.

Louis, MO). Alexa 488-conjugated donkey anti-mouse IgG and Alexa 488-conjugated donkey anti-human IgG were both obtained from ImmunoResearch, (West Grove, PA).

Alexa 488-conjugated polyclonal anti-human CD6 was obtained from R&D,

(Minneapolis, MN). FITC-conjugated mouse IgG isotype control was obtained from

Biolegend, (San Diego, CA).

Immunoprecipitations

3A11 mAb IP:

HBL-100 breast carcinoma cells were biotinylated using E-Z link sulfo-NHS-LC biotin and subsequently lysed in NP-40 lysis buffer (Invitrogen, Carlsbad, CA, USA) containing 0.1 % SDS (Fisher Scientific, Waltham, Ma), 0.1% deoxycholic acid (Sigma-

Aldrich, St. Louis, MO, USA) and one complete tablet of protease inhibitor (Roche,

Mannhein, Germany) on ice for 30 min. Immunoprecipitation was performed overnight at

4° C using either mouse IgG1 or 3A11 mAb. Antigen-antibody complexes were pulled down using protein (A+G). Following antigen-recombinant protein complex pull down,

the samples were boiled for 5 min in 2X Laemmli sample buffer (Biorad, Hercules, CA).

For Western Blot electroporesis: samples were loaded onto an SDS-PAGE and following electrophoresis, proteins were transferred to a polyvinylidene difluoride (PVDF) membrane for western blotting. The membrane was incubated for 1 hour at room temperature with blocking buffer containing 5% BSA and 0.05% Tween 20. Following blocking, the protein was detected using streptavidin-HRP conjugate and visualized using the chemiluminescent substrate ECL (Amersham Biosciences, Buckinghamshire, U.K.).

In some experiments, HBL-100 carcinoma cells were not biotinylated prior to the preparation of cell lysates for immunoprecipitation with either mouse IgG1 or 3A11 mAb as described above. Following blocking of the PVDF membrane, the membrane was incubated with CDCP1 (Thermo Scientific, Waltham, MA) primary antibody and followed by HRP-conjugated goat anti-rabbit (Southern biotech, Birmingham, AL). The protein was visualized using ECL Western Blot detection reagent (Amersham

Biosciences, Buckinghamshire, U.K.).

Recombinant soluble CD6 IP:

HT1080 CD166-KO cells were lysed in 0.5% NP-40 (Roche, Mannhein,

Germany) lysis buffer on ice for 30 min. Lysates were immunoprecipitated with either purified human IgG1 or recombinant mouse CD6 overnight at overnight at 4°C. Protein

A/G-agarose beads was then added to the samples for 3 hr. Following antigen- recombinant protein complex pull down, the samples were boiled for 5 min in 2X lamini sample buffer (Bio-rad, Hercules, CA). For Western Blot electroporesis: samples were loaded onto an SDS-PAGE and following electrophoresis, proteins were transferred to a

PVDF membrane for Western Blotting. Following blocking of the PVDF membrane, the membrane was incubated with an anti-CDCP1 (Thermo Scientific, Waltham, MA) primary antibody and followed by HRP-conjugated goat anti-rabbit IgGs (Southern biotech, Birmingham, AL).

In-gel Digestions and Mass Spectrometry

Proteins immunoprecipitated by mAb 3A11 were separated by SDS-PAGE.

Bands at the appropriate size were excised and destained with 50% acetonitrile in 100 mM ammonium bicarbonate followed by 100% acetonitrile. Cysteine residues were first reduced by incubating the sample with 20 mM DTT at room temperature for 60 min, and then alkylated with 50 mM iodoacetamide for 30 min in the dark. The gel pieces were washed with 100 mM ammonium bicarbonate, dehydrated in acetonitrile, dried in a

SpeedVac centrifuge, and then rehydrated in 50 mM ammonium bicarbonate containing sequencing grade modified trypsin for overnight digestion at 37°C. The resulting proteolytic peptides were extracted from the gel with 50% acetonitrile in 5% formic acid, dried and reconstituted in 0.1% formic acid for LC-MS/MS analysis.

The digests were analyzed by LC-MS/MS using Orbitrap Elite Hybrid Mass

Spectrometer (Thermo Electron, San Jose, CA), equipped with a Waters nanoAcquity

UPLC system (Waters, Taunton, MA). The spectra were acquired in the positive ionization mode by data-dependent methods consisting of a full MS scan at 120,000 resolution and MS/MS scans of the twenty most abundant precursor ions in ions trap by collision-induced dissociation at normalized collision energy of 35%. A dynamic exclusion function was applied with a repeat count of 2, repeat duration of 30 s, exclusion

duration of 45 s, and exclusion size list of 500. The obtained data were submitted for a database search using Mascot Daemon (Matrix Science, Boston, MA).

Carbamidomethylation of Cys was set as a fixed modification, whereas oxidation of Met was selected as variable modifications. The mass tolerance was set as 10 ppm for precursor ions and 0.8 Da for productions. SwissProt (July, 2014) database (546000 sequences; 194259968 residues) was used for searching against the taxonomy of human

(20210 sequences). The significance threshold p value was set to < 0.05. Proteins hits with at least two unique peptides at Mascot score > 20 were considered to be identified.

Flow cytometric staining

For CD318, CD166 and 3A11 mAb cell surface staining, cells were stained with anti-human CD318, anti-human CD166, and 3A11 mAb respectively on ice for 30 min.

Following 3A11 mAb cell surface staining cells, these cells were washed and subsequently stained with the secondary antibody Alexa 488-conjugated donkey anti- mouse IgG and analyzed by flow cytometry. For soluble CD6 cell surface staining,

HT1080 CD166-KO cells were incubated with 1μM recombinant human CD6-Ig or human IgG1 at 4 °C for 45 min. After 45 min, the cells were washed and subsequently stained with Alexa 488-conjugated donkey anti-human IgG at 4°C for 30min, washed and analyzed by flow cytometry. In some experiments, HBL-100 cells or synovial fibroblasts were stimulated with 1000 U/ml of human IFN-γ for 72 hr prior to analyzing the expression of CD318 by flow cytometry.

For rCD318 binding to control or human CD6-expressing CHO cells, these cells were incubated with rCD318 at 4°C for 45 min. After 45 min, the cells were washed and

subsequently stained with PE-conjugated mAb against human CD318 at 4°C for 30 min, washed and analyzed by flow cytometry

Production of recombinant CD318 extracellular domains and Western Blots:

Gene sequence encoding for the extracellular domains of CD318 with a C- terminal 6XHis-tag, was synthesized (Genscript, NJ) and cloned into the expression vector pcDNA3.1. After transfection of the expression construct into 293 cells for transient expression, recombinant CD318 in the culture supernatant was purified by nickel affinity chromatography following published protocols. For western blots, the same amount of either recombinant CD318 or BSA was separated by SDS-PAGE and transferred to a PVDF membrane, then probed with either the 3A11 mAb or a rabbit anti- human CD318 antibody, followed by either rat anti-mouse HRP conjugate or goat anti- rabbit HRP conjugate respectively. Protein bands were visualized using the chemiluminescent substrate ECL.

Synovial tissue and synovial fluid specimens:

Synovial tissue specimens were obtained from patients with rheumatoid arthritis

(RA, n = 13) and osteoarthritis (OA, n = 20) at the time of arthroplasty. Normal synovial tissues were obtained from cadavers (n = 17). A portion of each tissue was homogenized for ELISA assays. Synovial fluids were obtained at therapeutic arthrocentesis from patients with RA (n = 36), OA (n = 28) and juvenile inflammatory arthritis (JIA, n = 10).

In all cases synovial tissue and fluid specimens were excess material obtained at procedures performed for clinical indications.

Enzyme-Linked immunosorbent Assay (ELISA):

Levels of soluble CD318 in sera and synovial fluids were measured by an ELISA kit (R&D Systems, MN) per the manufacturer’s protocol.

T-cell Chemotaxis assay.

CD3+ T-cells were isolated using a RosetteSep lymphocyte isolation kit (Stem

Cell Technologies, Danvers, MA) and instructions were followed per the manufacturer’s protocol. Cytospin examination was performed to ensure that the isolated cells were lymphocytes. Chemotaxis membranes were coated with type IV collagen (Sigma- l of varying concentrations of CD318 (100, 200, 400, 800, and 1600 pg/ml) were added to wells in the bottom of the chamber. T-cells (1.0 × 106 cells/ml in PBS) were placed in the top wells of a 48-well Boyden chemotaxis chamber. PBS served as the negative control in this experiment. After 18 hours, the membranes were then removed and stained with Diff-Quik (Thermo Fisher Scientific, Kalamazoo, MI). Readings represent the number of cells migrating through the membrane (the sum of three high power 40x fields/well, averaged for each quadruplicate well).

Adhesion Assay:

CD3+ T-cells were isolated using a RosetteSep lymphocyte isolation kit (Stem

Cell Technologies) and were stained with carboxyfluorescein succinimidyl ester (CFSE)

(Thermo Fisher Scientific, Waltham, MA ) at 5 uM . RA synovial fibroblasts at culture

(Cell Signaling Technologies, Danvers, MA) in 10% FBS in culture media or in 10%

FBS for 3 days. Fibroblasts were then incubated with mouse IgG, anti-CD318, anti-

CD166, or both anti-CD318 and anti-CD166. T-cells were added at 50,000 cells per well onto fibroblasts and incubated for 1 hour at room temperature. Intensity was measured with a Synergy plate reader.

CHAPTER 3:

CD6 receptor regulates intestinal ischemia/reperfusion-

induced injury by modulating natural IgM-producing

B1a cell self-renewal

This work has been published. J Biol Chem. 2017 Jan 13. PMID: 27909060.

Introduction

The intestine is probably the most sensitive internal organs for ischemia/reperfusion (I/R)-induced injury [1]. Intestinal I/R injury often occurs during abdominal and thoracic vascular surgery, intestinal transplantation, hemorrhagic shock, and surgery using cardiopulmonary bypass. Without intensive management, it leads to intestinal damage, sepsis, systemic inflammation, multiple-organ failure, and eventually, death [4]. Currently available options for preventing or treating intestinal I/R induced injury are limited, therefore, understanding the regulatory mechanisms would help to develop novel therapies for this life-threatening condition [1, 4, 174].

Many inflammatory cytokines contribute to the breach of gut epithelial barrier during intestinal I/R induced injury. Among them, IL-6 locally produced in the intestine has been identified as a key inflammatory cytokine that plays an essential role in damaging the tissue during intestinal I/R. In support of this, it has been demonstrated that 1) both in humans [10] and in mice [175, 176] , local levels of IL-6 in the intestine are markedly elevated in association with severe tissue damage after intestinal I/R, 2) IL-

6-/- mice developed significantly attenuated intestinal damage after I/R, and 3) blocking

IL-6 activity in WT mice using anti-IL-6 mAbs greatly retained intestinal tract integrity after I/R [15] .

Natural IgM has been found to be critical in the pathogenesis of intestinal I/R.

These natural antibodies recognize antigens exposed on hypoxic cells after intestinal ischemia, and activate complement through the lectin pathway to produce C5a, which can facilitate neutrophil and monocytes infiltration and inflammatory cytokine (e.g. IL-6) production [24, 25]. The first evidence indicating that natural IgM initiates the

inflammatory processes to induce intestinal I/R-induced injury was found in studies using the immunodeficient RAG-/- mice, which do not have any antibodies [26]. These mice are highly resistant to intestinal I/R-induced injury in association with significantly reduced IL-6 production, but become susceptible again following reconstitution with purified serum IgM from naïve WT mice, highlighting the critical role of natural IgM in the development of intestinal I/R-induced injury [22, 26-29]. Additionally, recent studies have demonstrated the initiation of intestinal I/R-induced injury is not an inherent property of all natural IgM, but a subset of natural IgM termed pathogenic natural IgMs

[29-31, 33]. Despite the established role of natural IgM in the pathogenesis of intestinal

I/R-induced injury, regulatory mechanisms underlying the production of natural IgM are inadequately studied.

Natural IgM is spontaneously secreted by B1 B cells [38-40]. B1 B cells can be further subdivided into B1a (CD5+) and B1b (CD5-) [41]. Although still debatable, it has been demonstrated that B1a cells outside of the peritoneal cavity produce majority of the natural IgM in naïve mice independent of T cells [38, 42, 177], while B1b cells are responsible for T-independent IgM memory response [178], but are not a major source of natural IgM. How these natural IgM-producing B1a cells are regulated are also poorly understood, despite the findings that antigen specificity and B cell receptor (BCR) signaling strength are critical factors in B1a cell development because deletion of BCR co-stimulatory molecules such as CD19 results in a massive reduction of B1a numbers, while deletion of negative regulators of BCR signaling such as Siglec-G leads to a vast increase in B1a cell population [50, 86].

CD6 is a cell surface glycoprotein receptor originally discovered as a marker of T cells, and latter was found present on a subset of human B cells [89, 179]. The precise function of CD6 in T cells is still uncertain. Previous studies suggested that CD6 is a costimulatory molecule that can synergize with the T cell receptor to enhance and/ or inhibit T cell activation [94, 113, 115, 180]. Compared to the undefined role of CD6 on T cells, its role in B cells is even less clear. There has been only one report providing in vitro evidence suggesting that CD6 could regulate apoptosis of chronic lymphocytic leukemia B cells [116]. The distribution of CD6 on murine B cells, whether it has any role in natural IgM production, and, in the development of intestinal I/R induced injury is completely unknown.

In this study, using WT and CD6-/- mice, we studied the potential role of CD6 in regulating intestinal I/R-induced injury by comparing mucosal histopathology, local IL-6 production, and serum IgM titers. We explored the underlying mechanism by examining the distribution, regulation, and effect of CD6 on B1a cells. Our results showed the first evidence that CD6 is expressed on mouse B1a B cells and that CD6 regulates intestinal

I/R-induced injury by modulating natural IgM-producing B1a cell self-renewal.

Results

CD6-/- mice are protected from intestinal I/R-induced injury

To explore whether CD6 has any role in the gut epithelial barrier breaching and mucosal damage after I/R, we induced intestinal I/R-induced injury in sex- and age- matched WT and CD6-/- mice following previously published protocol [26], and compared the clinical scores of the jejunum as well as local levels of IL-6 in these

intestinal segments. We found that CD6-/- mice showed marked increased epithelial cell layer integrity within the intestinal villi ( Fig 1b) with 3.5-fold lower tissue damage in the intestine after I/R than WT as quantified by clinical scores assigned in a blinded fashion by a Pathologist (Fig 1a). In correlation with these histopathological analysis results in the CD6-/- mice after intestinal I/R, ELISA analyses of the jejunum tissue lysate also found 22-fold lower levels of IL-6 than those in WT mice (Fig 1c). Thus, the severity of intestinal I/R-induced injury is significantly attenuated in the CD6-/- mice, demonstrating a previously unknown role of CD6 in this pathological condition.

Pathogenic natural IgMs are integrally involved

It has been previously reported that natural IgM plays a critical role in inducing mucosal damage in this model of intestinal I/R-induced injury [26]. To understand the mechanism by which deficiency of CD6 protected mice from intestinal I/R-induced injury, we measured serum levels of total IgM in the serum of naïve WT and CD6-/- mice.

We found that while the titers of total IgG in the serum were similar between WT and

CD6-/- mice (Fig 2a), the titers of natural IgM were reduced 1.9-fold in the CD6-/- mice compared to WT mice (Fig 2b). Since natural IgM is reported to be polyreactive and only the pathogenic natural IgMs, e.g., anti- phosphorycholine (PC) IgM and anti-Annexin IV

IgM, are critical in initiating intestinal I/R-induced injury [27, 181], we specifically measured titers of these pathogenic IgMs in the WT and CD6-/- mice that were used in the

I/R studies and found that a 1.2 and 1.4-fold reduction of Annexin IV- and PC-specific

IgM titers respectively in the sera of CD6-/- mice (Fig 2C & Fig 2D). To determine that reduced titers of pathogenic natural IgMs protected CD6-/- mice from I/R-induced injury, we reconstituted IgM in the CD6-/- mice by tail vein i.v. injection of IgMs purified from naïve WT mice before I/R procedures. We found that reconstitution restored the titers of the PC and Annexin V-reactive IgMs in the CD6-/- mice to those in the WT mice (Fig 2C and Fig 2D) and that these reconstituted CD6-/- mice showed increased local inflammation after I/R (Fig 2E).

CD6 is selectively expressed on B1 cells outside the bone marrow and peritoneal cavity

The above-described results suggest that CD6 could have a previously unknown role in regulating B1a cells, which are the source of natural IgM. To test this, we first examined the expression of CD6 on all B cells and, then focused on the B1a cells. We found that CD6 is not expressed on splenic B2 cells (specifically Marginal and Follicular

B cells ) neither constitutively (Fig 3a), nor after in vitro activation (Fig 3b).

Interestingly, using B1 cells from CD6-/- mice as controls, we found that CD6 is expressed on B1 cells in the spleen (Fig 4a), neonatal liver (Fig 4b), and peripheral blood

(Fig 4c), but not in the peritoneal cavity (Fig 4d) and bone marrow ( Fig 9a). Among the

B1 cell subsets in the spleen, neonatal liver and peripheral blood, we found that the expression of CD6 was highly expressed on B1a cells compared to B1b cells. The full flow gating strategy for B1 B cells in the spleen, neonatal liver, peripheral blood, peritoneal cavity and bone marrow are shown in Fig (8a,b,c, d and 9b respectively).

Peritoneal B1a cells gain CD6 expression outside the peritoneal cavity

Previous studies showed that B1a cells homeostatically migrate back and forth between the peritoneal cavity and the spleen [44, 85]. To explore the mechanism by which CD6 is not present on B1a cells in the peritoneal cavity, we tested whether these

B1a cells will gain the expression of CD6 after they move out the peritoneal cavity. To do this, we first examined the expression of CD6 on “migrant” peritoneal B1a cells

(CD19+CD5+CD11b+) and found that CD6 is expressed on these “migrant” peritoneal

B1a cells in the spleen (Fig 5a). To further confirm this result, we isolated peritoneal cells from WT mice, labeled them with CFSE, then injected them i.p. into naïve CD6-/- recipient mice. To induce the adoptively transferred B1a cells to migrate out of the peritoneal cavity, we injected the recipient mice with LPS i.p. immediately after [44]. At

72 h, we analyzed CD6 expression on the CFSE+ peritoneal cells in the CD6-/- recipient mice and found that these adoptively transferred peritoneal B1a cells gained CD6 expression in the spleen (Fig 5b), suggesting that CD6 expression is induced on peritoneal B1a cells after they exited the peritoneal environment.

The B1 cell population is reduced in the absence of CD6 in select tissue compartments

Our data so far showed that 1) lack of CD6 protected mice from intestinal I/R injury, in which natural IgM plays a critical role in the pathogenesis; 2) blood levels of natural IgM were significantly reduced in the CD6-/- mice, and 3) CD6 is expressed on B1 cells outside of the peritoneal cavity and bone marrow, which are one of the major sources of natural IgM. To determine why deficiency of CD6 leads to reduce IgM production, we compared the B1a cell population in the neonatal liver, peripheral blood and spleen of WT and CD6-/- mice by flow cytometry. We found a 4 and 2.5-fold reduction in the B1a cells frequency and absolute numbers in the CD6-/- mice neonatal liver respectively (Fig 6a). The decrease of the B1a cell population in the CD6-/- neonatal

liver was compensated with a 1.7 and 2.9-fold increase in the B1b cell frequency and absolute numbers as anticipated (Fig 6a, table 1). In the peripheral blood, the frequency of B1a cells was decreased by 1.5-fold in the CD6-/- mice (Fig 6b, table 1). The frequency of B1b cells in the peripheral blood was increased by 1.4-fold in the CD6-/-mice (Fig 6b, table 1).

In the spleen, we found a 2 and 2.5-fold decrease in the frequency and absolute numbers of B1a cells in the CD6-/-mice (Fig 6c, table 1). In the bone marrow, we found no significant difference in the frequency and absolute numbers of B1 cells ( Fig 9c, table

2). In the peritoneal cavity, we made some interesting observations. We found a 1.2 and

1.6-fold reduction in the B1 cell frequency and absolute numbers respectively in the CD6-

/- mice. Upon further analysis of the B1 cell subsets, we found a 1.8-fold increase in the

B1a cell frequency in the CD6-/- mice. Although there was a 1.3 fold increase in the total

B1a cell numbers in the CD6-/-, there was no significant difference compared to WT mice.

Additionally, in the CD6-/- peritoneal cavity we found a 1.2 and 1.7-fold compensatory reduction in the B1b cell frequency and absolute numbers respectively (Fig 9d, table 2).

Together, these data indicate that CD6 is required to maintain a normal B1a cell population in these tissues and the significantly reduced population of B1a cells in CD6-/- mice could explain the markedly reduced blood IgM levels that we found in above experiments.

Reduced B1a cell population in CD6-/- mice is due to impaired B1a cell self-renewal

We then explored the possible mechanism by which the absence of CD6 resulted in the reduced numbers of B1a cells in the CD6-/- mice. The B1a cell population is maintained by their unique ability to undergo self-renewal [182, 183]. In asplenic mice, the B1a cell population is significantly reduced in multiple tissue compartments, highlighting the importance of the spleen in B1a self-renewal [184]. To determine if the reduced B1a cell population in the CD6-/- mice is caused by their impaired ability to undergo self-renewal, we measured B1a cell proliferation between WT and CD6-/- mice using a BrdU incorporation assay. Incorporation of BrdU by splenic B1a cells was reduced by 1.5-fold in the CD6-/- mice (Fig 7a). To determine if CD6 plays an intrinsic role in B1a cell proliferation, we also assessed BrdU incorporation of peritoneal B1a and bone marrow B1 cells that do not express CD6. We found no significant difference in either peritoneal B1a ( Fig 10a) or bone marrow B1 cell proliferation between WT and

CD6-/-mice (Fig 10b). In addition to impaired self-renewal, reduced B1a cell population in CD6-/- mice may result from enhanced apoptosis of these cells given that CD6 has been implicated in the survival of B1a-derived B-CLL cells [116]. We compared the frequency of apoptotic splenic B1a cells in WT and CD6-/- mice before and after LPS stimulation by Annexin V staining. There were no differences in splenic B1a apoptosis basally or upon activation (Fig 7b). These data indicate that the reduction of B1a cell population in the CD6-/- mice is due to impaired self-renewal of B1a cells and not enhanced apoptosis.

Elevated basal Erk phosphorylation is one of the hallmark features of naive B1a cells [185]. To determine whether the reduced splenic B1a cell self-renewal in CD6-/- mice is associated with impaired Erk phosphorylation, we compared the basal Erk phosphorylation of WT and CD6-/- splenic B1a cells. CD6-/- splenic B1a cells displayed a slight increase in basal Erk phosphorylation compared to WT, however, the difference was not significant (Fig 7c). We also analyzed the expression levels of NFATc1, a transcription factor that has been shown to be important for B1a cell development/survival [186]. A slightly elevated but insignificant increase in NFATc1 expression levels was detected in CD6-/- splenic B1a cells (Fig 7d). Taken together, these data suggest that reduced self-renewal of CD6-deficient B1a cells is independent of impaired Erk signaling and NFATc1 activity.

Discussion

In this report, we studied CD6-/- mice in a model of intestinal I/R-induced injury, a devastating and likely lethal pathological condition. We found that CD6-/- mice were protected from intestinal I/R-induced injury as shown by significantly attenuated histopathology of the intestine and reduced levels of local IL-6. In mechanistic studies, we found that total and PC-specific IgM titers were reduced in the CD6-/- mice, and that reconstitution of CD6-/- mice with IgM from naïve WT mice significantly restored intestinal I/R-induced inflammation. In addition, we demonstrated that CD6 was selectively expressed on natural antibody producing-B1 cell outside of the peritoneal cavity and bone marrow. Finally, we found that CD6-expressing B1 cells were significantly reduced in select tissue compartments in the CD6-/- mice in association with reduced B1a cell proliferation.

Intestinal I/R-induced injury is a condition frequently associated with high morbidity and mortality in patients with trauma and surgeries, especially small intestine transplantation [1]. The unexpected results that CD6-/- mice were protected in the intestinal I/R-mediated injury demonstrated a previously unknown role of CD6 in the pathogenesis of this condition. CD6 was originally identified as a marker for T cells, and has been associated with multiple autoimmune diseases including rheumatoid arthritis and psoriasis [118, 119]. Quantitative proteomics analysis of signalosome dynamics in T cells identifies CD6 as a Lat adaptor–independent TCR signaling hub, suggesting that

CD6 modulates TCR signaling [106]. A recent study using CD6-deficient mice found exacerbated collagen-induced arthritis in the CD6-/- mice [112], a surprising result since anti-CD6 mAbs showed efficacy in treating patients with rheumatoid arthritis and psoriasis. Compared to the debatable role of CD6 on T cells, information on the potential role of CD6 on B cells is even more limited. So far, it only has been reported that proportion of blood CD6+ B cells is reduced in primary Sjögren's syndrome patients

[187], and that CD6 ligation modulates the Bcl-2/Bax ratio and protects chronic lymphocytic leukemia B cells from apoptosis[188]. Our results that CD6 deficiency led to impaired B1a cell self-renewal, reduced B1a cell population and natural IgM titers in the blood showed a new role for CD6 in regulating B cells for the first time.

The discovery that CD6-/- mice have reduced titers of pathogenic natural IgM and are protected from intestinal I/R-induced injury is consistent with previous reports that natural IgM is critical in I/R-induced injury [26, 28]. RAG1-/- mice, which do not have any antibodies, are highly resistant to intestinal I/R-induced injury but become susceptible again following reconstitution with purified serum IgM from naïve WT mice

or, an IgM mAb alone (clone B4), which specifically recognizes Annexin IV [33]. In addition to Annexin IV, natural IgM reactive to PC has also been implicated in the pathogenesis of intestinal I/R-induced injury [16, 27, 33]. Consistent with these previous reports, we found that natural IgM specific for PC and Annexin IV were significantly reduced in the blood of CD6-/- mice compared to WT mice in association with decreased intestinal I/R-induced injury, and that supplementing IgMs isolated from naïve WT mice into CD6-/- mice increased titers of both PC- and Annexin IV specific IgMs, leading to exacerbated intestinal inflammation after I/R.

Beside nature IgM, several studies has implicated a role for T cells in this acute inflammatory model [18, 189, 190]. The recent report of impaired Treg function in the

CD6-/- mice along with enhanced susceptibility to a model of T cell mediated autoimmune disease would suggest that CD6-deficiency should result in exacerbated intestinal I/R- induced injury if CD6-mediated function on T cell is critical in this acute inflammatory model. However, the finding that CD6-/- mice are protected from intestinal I/R-induced injury in association with reduced natural IgMs and that reconstitution with IgM in CD6-/- mice restored their susceptibility to intestinal I/R-induced injury emphasizes the significant contribution of CD6-mediated function on natural antibody secreting B1a cells in the pathogenesis.

The absence of CD6 expression on mature B2 cells in mice was an interesting finding because CD6 is reported to be present on mature B2 cells in humans [89]. CD6 is closely related to the lymphocyte receptor CD5 [191]. These two genes are immediately adjacent to each other and are highly homologous members of the group B scavenger receptor cysteine-rich (SRCR) superfamily of protein receptors, previously reported to be

co-expressed on the same cells [191, 192]. In mouse B cells, CD5 is not expressed on naïve B2 cells; however anti-IgM stimulation of B2 cells induces CD5 expression [193].

In our studies, in vitro stimulation of mouse B2 cells did not induce appreciable expression of CD6. These findings suggest that while CD5 and CD6 are closely related, the mechanism regulating their expression in mouse B cells is distinct and different from

T cells wherein expression of CD5 and CD6 appears coordinated.

The discovery that CD6 is differentially expressed on mouse B1 cells, adds CD6 to a list of unique markers that are differentially expressed between peritoneal and splenic

B1 cells [194]. This finding also indicates that bone marrow B1 and splenic B1 cells may also express differential markers. Differential expression of CD6 raises the possibilities that CD6+ B1 and CD6- B1 cells may arise from two distinct B1 cell lineages and/or that

CD6 expression is heavily influenced by the local microenvironment. The expression of

CD6 on “migrant peritoneal B1a cells” in the spleen and the discovery that WT peritoneal

B1a cells acquire CD6 expression in the spleen of CD6-/- recipient mice after adoptive- transfer strongly supports the notion that the microenvironment greatly influences expression of CD6 and drives phenotypic differences in B1a cells [82, 84].

The deletion of certain BCR signaling positive regulators results in the massive reduction of B1a cells [182]. Reduction of the B1a cell population outside the bone marrow and peritoneal cavity in the CD6-/- mice, suggest that CD6 is a positive regulator of BCR signaling. In T-cells, CD6 can initiate its own signaling hub and tune overall T cell receptor signaling threshold thru its ability to recruit signaling molecules such as

SLP-76 and Vav1 [106]. Given that enhanced BCR signaling is one of the critical factors

in B1a development, our data support the hypothesis that CD6 signaling contributes to the overall BCR signaling threshold that shapes the B1a development process.

The B1 cells in the bone marrow and peritoneal cavity do not express CD6. As anticipated, the deficiency of CD6 did not alter the B1 cell population in the bone marrow. In contrast, we found reduced B1 cells in the peritoneal cavity of CD6-/- mice.

These findings are consistent with the recent study indicating that serum IgM-deficient mice display reduced B1 cell population in the peritoneal cavity and normal B1 cell population in the bone marrow [195]. These serum IgM-deficient mice also harbor large amounts of anergic CD5+ B cells, which potentially could account for the elevated CD5+

B1a cells population in the peritoneal cavity of CD6-/- mice. The discovery that the B1 cell population in the CD6-/- mice bone marrow are not altered despite altered B1 cell population in the peripheral tissue suggest a potential defect in the renewal/proliferation rather than in the development of B1 cells. We assessed the proliferation of B1a cells and discovered that CD6-deficient mice displayed reduced proliferation compared to WT.

Specifically, the defect was only found in CD6-expressing B1a cells. These data suggest that CD6 plays an intrinsic role in B1a cell self-renewal.

Thus, our working model is that CD6 deficiency impairs the self-renewal of natural IgM-secreting B1a cells, therefore reducing the B1a cell population. The reduction of the B1a cell population results in reduced pathogenic natural antibodies

(Annexin IV and PC-specific IgM) in the blood. Reduced recognition of ischemic neoantigens by natural IgM subsets in the CD6-/- mice lead to impaired inflammatory responses such as IL-6 production during reperfusion of the intestine, and, consequently, attenuated mucosal damage. This study provides new insight into the role of CD6 in the

inflammatory process that induces intestinal I/R-induced injury and on B1a cells that produce natural IgM, which drives the onset of intestinal I/R-induced injury.

Chapter Four:

CD318/CDCP1 is a novel CD6 ligand

Introduction

CD6 is a marker of T cells and an important T cell regulator[179]. Recent genome-wide association studies also identified CD6 as a risk gene for multiple sclerosis

(MS)[196-199], an autoimmune disease in which T cells play a vital role in the pathogenesis. CD6 is composed of three extracellular domains (domain 1, 2 and 3), and it functions by interacting with its ligand (s) [171]. The domain 3 of CD6 has been shown to be the site that the identified CD6 ligand, CD166, a.k.a., ALCAM (activated leukocyte cell adhesion molecule) binds to[107]. However, anti-CD166 antibodies only partially blocked the binding of thymic epithelial cells to CD6 overexpressing COS cells, and mAbs blocking CD6-CD166 interactions do not abolish CD6 function [115, 180].

Itolizumab, an anti-CD6 mAb developed in Cuba and approved in India for treating psoriasis, reduces pathogenic T cell responses in patients with psoriasis, but this mAb binds to domain 1 of CD6 instead of domain 3, and it does not interfere with the CD6-

CD166 interaction . Interestingly, UMCD6, a mouse anti-human CD6 mAb that we found highly effective in treating EAE in CD6 humanized mice (manuscript submitted), also fails to block the CD6-CD166 interaction. All these studies suggest the existence of an additional CD6 ligand, other than CD166, that binds to domain 1 of CD6, and could be critical for CD6 function in autoimmune conditions. Further studies using a CD6 fusion protein as a bait to pull down CD6-binding proteins from synovial fibroblast surface proteins showed the binding of 3 polypeptides[173]. One of these polypeptides was identified as CD166 and the identities of other two were unknown [136]. A mAb termed 3A11 was developed and the antigen recognized by this mAb was identified as

the new ligand of CD6 that binds to its domain 1[136] [91]. However, attempts to identify the antigen recognized by mAb 3A11 were not previously successful.

CD318 (a.k.a. CDCP1, TRASK, SIMA135, or gp140) is a cell-surface glycoprotein with an apparent molecular weight of ~140 kDa [135, 137, 200]. It consists of 3 extracellular CUB domains, a transmembrane domain, and an intracellular domain.

CD318 can be proteolytically cleaved between the two distal CUB domains by certain serine proteases, resulting in different ratios of the ~140 kDa intact molecule and the ~80 kDa cleaved product on different cells. Cleaved CD318 is phosphorylated and activated by Src kinase, then the activated CD318 forms a complex with activated β1 integrin and activates FAK/PI3K/Akt motility signaling to promote early tumor dissemination [160].

Under normal conditions, CD318 is present on many epithelial cells,[201] some hematopoietic cells,[151] and mesenchymal stem cells[150]. CD318 is also present on many tumor cells.[202] Upregulation of CD318 expression is associated with a poor prognosis for many cancer patients[137, 138, 163, 203-205]. Interestingly, a recent study using CD318 KO mice showed that two different oncogene-driven tumors grow much faster in CD318 KO mice than in wild-type (WT) control mice.[206] Lack of CD318 in these mice enhances tumor growth potentially by liberating integrin signaling and growth factor receptor cross-talk in unanchored tumor cells.[206] So far, all studies on CD318 have been limited to its direct signaling effect in tumor cells, and its possible role in regulating immune responses has never been examined.

In this report, using mass spectrum techniques, we identified the antigen recognized by mAb 3A11 as CD318. To validate the proteomics results, we probed proteins immunoprecipitated by mAb 3A11 using established anti-CD318 antibodies and

probed recombinant CD318 protein by mAb 3A11 in Western Blots. We also compared staining patterns of both mAb 3A11 and established anti-CD318 mAbs using cells that are known to be positive or negative for CD318, and engineered cells with upregulated or downregulated levels of CD318. In addition, we confirmed the binding of CD318 to CD6 by using soluble CD6 protein as a bait in pull-down assays and by staining WT and

CD166 deficient cells with the soluble CD6 followed by flow cytometric analyses.

Finally, we examined CD318 expression on synovial fibroblasts from rheumatoid arthritis (RA) patients, measured levels of soluble CD318 in synovial fluids from arthritis patients, and studied its potential roles in recruitment and retention of T cells in synovial tissue. Our results showed that CD318 is the new CD6 ligand recognized by mAb 3A11, and suggest that CD318 could be a new biomarker and target for the diagnosis and/or treatment of inflammatory arthritis.

Results

Identification of the antigen recognized by mAb 3A11:

It had been previously established that mAb 3A11 recognizes an uncharacterized

CD6 ligand that binds to domain 1 of CD6[136], the same domain that Itolizumab, the anti-CD6 mAb approved for treating psoriasis in India[207] binds to, suggesting that this new CD6 ligand might be more important than the already identified CD6 ligand,

CD166, for CD6 function in disease. To determine the identity of the antigen recognized by mAb 3A11, we investigated HBL-100 cell surface proteins pulled down by this mAb by mass spectrum (MS) analysis. We found that CD318-related peptides were abundant in the mAb 3A11 precipitates (data not shown), indicating that the protein recognized by

3A11 could be CD318. We then probed whole HBL-100 cell lysate with an anti-CD318

Ab in western blot and assessed CD318 expression levels on HBL-100 cells by flow cytometry before and after IFNγ stimulation . We found that CD318 met the previously established characteristics of the potential mAb 3A11 antigen candidate [136] such as 1) it has a molecular weight of ~130 kDa (Fig 11A), and 2) its expression can be upregulated by IFNγ stimulation (Fig 11B).

Western blots using a commercial anti-CD318 antibody and recombinant CD318:

To validate the MS results, we performed western blotting of the above immunoprecipitates using a commercial anti-CD318 antibody (Pierce, IL) and found that these antibodies detected three bands (Fig 11C), including a ~140 kDa band and a

~80kDa band, in the mAb 3A11 immunoprecipitates, but not the control IgG1 immunoprecipitates. In addition, we prepared recombinant soluble CD318 (rCD318) by synthesizing an artificial gene coding for the extracellular domains of CD318 with a C- terminal 6XHis-tag, and cloned it into the expression vector pcDNA3.1. After transfecting the expression construct into 293 cells, we purified the rCD318 in the culture supernatants by nickel affinity chromatography following published protocols[135], and verified the protein by western blot using an anti-His tag antibody (data not shown). We then probed the rCD318 and the same amount of BSA with mAb 3A11 or an established anti-CD318 antibody in western blots, and found that both the mAb 3A11 and the anti-

CD318 antibody selectively recognized rCD318 but not the BSA (Fig.11D).

Flow cytometric analysis of cells normally expressing or not expressing CD318 using both mAb 3A11 and a commercial anti-CD318 mAb

CD318 has been reported to be present on A549[137], HBL-100[208] and

Caco2[209] cells, but not on MCF-7[210], Molt-4[149] or Raji cells[137]. We analyzed all of these cells with a commercial anti-CD318 mAb (Fig 12A) or mAb 3A11 (Fig.

12B), and found exactly the same staining pattern, suggesting that mAb 3A11 and the anti-CD318 mAb recognize the same antigen.

Flow cytometric analysis of engineered cells with CD318 overexpression or downregulation

We cannot exclude the slight possibility in the above-described flow cytometry experiments that it was a coincidence that mAb 3A11 and the anti-CD318 mAb might recognize different antigens that happen to have the same expression pattern on the various examined cell lines. To address this issue, we studied the transfected MDA-468 cells that overexpress CD318 after doxycycline induction[135, 172] and the transfected

MDA-468 cells knocked down for CD318 expression using shRNA[172] by flow cytometry using the commercial anti-CD318 mAb and mAb 3A11. We found that, consistent with previous reports[172], following doxycycline treatment the expression of

CD318 increases above basal levels in MDA-468 expressing CD318 inducible system and not in cells expressing empty vector (control) (Fig 13A).These assays also showed that staining of these cells expressing CD318 with mAb 3A11 resulted in exactly the same pattern as the anti-CD318 mAb (Fig 13A), while, in CD318 knocked down MDA-

468 cells, which are CD318-negative, staining with either anti-CD318 mAb or mAb

3A11 was also negative (Fig 13B ). These additional data confirmed that mAb 3A11 recognizes CD318.

CD6 binding analysis on cells expressing both CD166 and CD318, or CD318 alone

After confirming that CD318 is the protein recognized by mAb 3A11 in the above experiments, we tested whether CD318 indeed binds to CD6, as suggested by previous studies. We have already shown that soluble CD6 protein can be used to stain cells expressing CD6 ligands in flow cytometric assays. The human fibrosarcoma cell line HT-

1080 expresses both CD318 and CD166[109, 211], and we generated an HT-1080 CD166

KO cell line by CRISPR/Cas9 technology to exclude the previously known CD6-CD166 interaction (Fig 14A). We first confirmed that indeed our HT-1080 CD166 KO cell line expresses CD318 but not CD166 (Fig 14B), then stained the WT and CD166 KO cells with the soluble CD6 protein. We found that, in the absence of CD166, the binding of

CD6 to the surface of these cells was significantly reduced but still evident (Fig 14C), further evidence that CD6 has ligand(s) other than CD166. We then carried out a competitive binding assay using our prepared soluble rCD318 protein and found that binding of CD6 to the CD166 KO cells was reduced by rCD318 in a dose-dependent manner (Fig 14D). In addition, we incubated soluble CD6-Fc protein or the same amount of purified human IgG1 with the CD166 KO cell lysates, and probed the CD6- participated proteins with a commercial anti-CD318 antibody in Western Blot. We found that CD6-Fc protein, but not the control human IgG1 protein, pulled down a protein that recognized by the anti-CD318 antibody (Fig. 14E). Finally, we stained transfected CHO cells expressing human CD6 on the surface and control CHO cells with the soluble rCD318 and found that rCD318 binds to human CD6- expressing CHO cells but not the control CHO cells (Fig 14F). These results demonstrate that CD6 binds to CD318.

Measurement of CD318 levels in synovial tissues from RA and OA patients by

ELISA

We have established previously that the antigen recognized by the mAb 3A11

(now shown to be CD318) is highly expressed in synovial fibroblasts from RA patients after IFNγ stimulation. To explore a potential role of CD318 in the pathogenesis of arthritis, we homogenized synovial tissues from patients without or with RA or OA and measured levels of total CD318 in them by ELISA. We found that CD318 is present in synovial tissues from all the synovial tissues examined.While levels of CD318 were comparable between the control groups without arthritis and with OA, levels of CD318 in synovial tissues from RA patients were significantly elevated (Fig. 15A).

Measurement of soluble CD318 levels in sera and synovial fluids from RA and OA patients by ELISA

Soluble CD318 has been found in cancer cell culture supernatants and in urine samples from men with high-risk prostate cancer[135]. We first measured levels of soluble CD318 in sera from RA patients and healthy donors. We found that levels of soluble CD318 in the sera were very low, barely above the sensitivity of the ELISA assay that we used (Fig. 15B). We also examined synovial fluids from patients with RA, JIA and OA by the same ELISA, and found that levels of soluble CD318 were significantly higher in synovial fluids than in plasma, and that levels of soluble CD318 were significantly higher in synovial fluids from both RA and JIA patients than in those from the OA patients (CD318 was not detectable in synovial fluids from OA patients) (Fig.

15B). These data suggest that soluble CD318 is produced within the joints and that it could be used a novel biomarker in synovial fluids to distinguish OA from RA and JIA.

T cell chemotaxis in response to soluble CD318

The gradient between serum and synovial fluid levels of soluble CD318 and the elevated levels of CD318 in synovial fluids from patients with RA and JIA but not in OA led us to assess the possibility that soluble CD318 might be chemotactic for T lymphocytes. Using a modified Boyden chamber assay we found that indeed, at a concentration that approximated the difference between the mean RA serum and RA synovial fluid soluble CD318 levels, peripheral blood T cells migrated in response to soluble CD318 as a single stimulus (Fig. 15C).

Contributions of CD6 ligands to adhesion between T cells and synovial fibroblasts

Previous work has suggested that a CD6 ligand other than CD166 could contribute to adhesion between human T lymphocytes and various IFNγ-treated nonhematopoietic cell types [136, 173, 212]. To evaluate the roles of CD166 and CD318 in interactions between T cells and synovial fibroblasts, we performed adhesion assays using fluorescently tagged T cells and synovial fibroblasts that were or were not pre- cultured with IFNγ, (a step that is required to re-induce expression of CD318 on these cells, which is lost between harvesting of the cells from synovial tissue and use of pure synovial fibroblast lines at passages 4-8 in culture). The expression of CD318 following exposure to IFNγ corresponds to the clear expression of this molecule in fresh synovial tissue (data not shown).

In these assays we found that CD6 ligands were significantly involved in the adhesion of T cells to synovial fibroblasts (Fig. 15D). Without IFNγ pre-treatment of the synovial fibroblasts, only CD166 was functionally important in these adhesion assays, consistent with the minimal expression of CD318 on these cells. Interestingly, when

IFNγ-treated synovial fibroblasts were used, both ligands were functional, and adhesion was substantially interrupted only when both were simultaneously masked with monoclonal antibodies. These results are consistent with important functional roles for both CD6 ligands in synovial tissue in vivo.

Discussion

In this report, using MS analysis, we identified abundant CD318-derived peptides in mAb 3A11-immunoprecipiated proteins. CD318 also met the previously published criteria of a 3A11 antigen, suggesting that CD318 is the antigen recognized by mAb

3A11. We confirmed the MS results by probing the mAb 3A11-immunoprecipiated proteins with a commercial anti-CD318 antibody, and by producing and probing recombinant CD318 with mAb 3A11 in western blots. In addition, we found that mAb

3A11 and the established CD318 mAb have the same staining patterns on cells known to naturally express or lack CD318, and on cells engineered to overexpress or knockdown

CD318 expression. Using soluble CD6, and soluble CD318 in a competitive binding assay, we demonstrated the binding specificity of CD318 to CD6. Additionally, we confirmed the binding of CD318 to CD6 by assessing the binding of soluble CD318 to transfected CHO cells expressing CD6 using flow cytometric analyses. Finally, utilizing pull-down experiments, we further validated that CD318 does bind to CD6. We found that CD318 is abundant in synovial tissues and while levels of total CD318 are similar in synovial tissues from controls and OA patients, they are significantly elevated in RA patients. Moreover, a soluble form of CD318 can be readily detected in synovial fluids from patients with inflammatory (RA, JIA) but not non-inflammatory (OA) arthritis, and these levels are appreciably higher than those in the serum. Of particular interest, both the membrane-bound and soluble forms of CD318 are functionally active in vitro, in adhesion and chemotaxis assays, respectively, that are arguably relevant to components of the pathogenesis on joint inflammation in vivo.

The distinct patterns of expression of CD318 compared to CD166 are striking – while CD166 is expressed widely on a broad range of hematopoietic and nonhematopoietic cells, including activated T lymphocytes, CD318 expression appears to be confined to non-hematopoietic lineages, such as fibroblasts, keratinocytes, epithelial cells and a variety of neoplastic cells [136, 173, 211, 212]. The one possible exception to the lack of CD318 expression on hematopoietic cells is a subset of cord blood hematopoietic progenitors [213]. Thus, engagement of CD6 by CD318 is an unusual example of a ligand-receptor interaction between a lymphocyte-specific cell surface glycoprotein that can participate in T cell activation (CD6) and a molecule (CD318) that is found only on cells that are traditionally considered not to be components of the immune system. This interaction points to an ability of T cells to specifically receive and recognize distinct signals from “non-immune system” tissue cells that may be important in organ-targeted autoimmune diseases, such as synovial fibroblasts and keratinocytes.

The functional roles of CD318 versus CD166 must be in part distinct given the marked differences in the distribution of these molecules. With the identification of

CD318 as a second ligand of CD6, exploration of the consequences of its binding to CD6 is still at an early stage. Membrane-anchored CD318 appears to be a mediator of T cell adhesion to tissue cells, such as fibroblasts and keratinocytes, and our data suggest that under some conditions both CD6 ligands, CD166 and CD318, can cooperatively participate in adhesion to T cells. These two CD6 ligands appear to recognize distinct epitopes of CD6, raising the possibility that CD6 can be engaged simultaneously by both of its ligands to form a tri-molecular complex. Structural demonstration of such a complex during interactions between intact cells is an important goal for future experiments. Early studies of CD6 identified distinct immunologic epitopes that appeared to mediate different functional effects on T cells[214], and more recent work has localized the binding domains on CD6 and CD166 that are involved in the CD6/CD166 interaction[215].

There are multiple isoforms of CD6 resulting from alternative splicing of cytoplasmic domain-coding exons that have been described, but with no distinct functions assigned. A novel isoform of CD6 lacking the CD166 binding domain has been described[92, 97]. This domain-3 lacking isoform of CD6 is significantly upregulated on activated T cells and does not localize to the T cell: antigen presenting cell interphase during antigen presentation unlike full length

CD6[97]. The identification of a CD318 as a novel CD6 ligand raises new perspectives to the potential function of this domain-3 lacking isoform of CD6 in the pathogenesis of RA. It is possible that the inflammatory microenvironment within the synovial tissue may enhance the activation state of the infiltrated T cells resulting in the upregulation of domain-3 lacking isoform. If CD318 indeed does interact with CD6 via domain-1 (See Summary and Future

direction part 2, structural domains that mediate CD6/CD318 interaction section), this presents a unique scenario in which switching from full length CD6 to domain-3 lacking isoform may induce CD6 –expressing T cells to transition from interacting with CD166-expressing antigen presenting cells to interacting with CD318-positive inflamed synovial fibroblast. This switch in ligand interaction may induce a different CD6-mediated intracellular signaling and function in T cells.

The signal transduction pathways that are activated by CD318 binding to CD6, with or without concurrent binding of CD166 to CD6, remain to be elucidated, both downstream of CD6 in the T lymphocyte and downstream of CD318 and CD166 in non-T cells that express these ligands. Signal transduction events in synovial fibroblasts generated by cell-cell adhesion events that involve CD6 and CD318 are likely to be distinct from pathways activated by anti-adhesive effects of the intracellular domain of CD318 that have been described in cancer cells[216]. In cancer cells CD318 is phosphorylated and can enhance downstream phosphorylation events, in part linked to signaling through the epidermal growth factor receptor [217, 218]. Moreover, cleavage of full-length 135kD CD318 by serine proteases creates a smaller membrane-retained 70 kD form that either associates with membrane integrins or homodimerizes, and generates downstream signaling events that enhance cancer cell invasiveness and metastasis[160, 218].

Cleavage of CD318 can be blocked by dexamethasone[167]. Like cancer cells, RA synovial fibroblasts are locally invasive and are stimulated by a variety of growth factors. Moreover, intra-articular injection of corticosteroid ameliorates joint inflammation. It will be important to elucidate the potential roles of CD318 in migration and invasion of synovial fibroblasts, and to assess the effect of engagement of CD318 by CD6 on these processes.

On synovial fibroblasts and keratinocytes CD318 is upregulated by IFNγ. However, on cancer cells its expression is increased following engagement of the epidermal growth factor

(EGF) receptor by EGF[219]. In cancer CD318 is also upregulated by hypoxia-inducible factor

2alpha, which is also potentially relevant to CD318 expression on synovial fibroblasts since inflamed synovium is a hypoxic environment. Methylation of sites near CD318 has been proposed as a critical element of epigenetic control of its expression. In bone marrow stromal cells reciprocal CD146+CD318- and CD146-CD318+ subsets of marrow fibroblasts have been identified that have distinct patterns of gene expression[152]; whether this is also true in synovium or other tissues is as yet unknown.

The elevated levels of soluble CD318 in inflamed synovial tissue and fluid (RA and JIA) raise questions regarding its function, and identify this molecule as potentially a measurable biomarker. Our data indicate that soluble CD318 is chemotactic for T cells, which are not present in normal synovial tissue, but which accumulate in large numbers in RA and JIA synovium through mechanisms that are as yet not fully defined. Importantly, the concentration at which soluble CD318 is chemotactic corresponds to the in vivo concentration gradient between RA serum and RA synovial fluid, indicating that this in vitro assay is likely to be physiologically relevant. However, there are several observations that call into question the reliability of this finding that soluble CD318 is chemotactic for T cells. Interestingly, we observed that increasing the concentration of soluble CD318 seemed to decrease its chemotactic effect on T cells.

Additionally, CD318 does not show structural resemblance to conventional chemokines and the proteolytic cleavage of full-length CD318 results in a soluble form that is about 6 times larger than the average molecular weight of a typical chemokine. Whether soluble CD318 is derived by protease-mediated shedding from the synovial fibroblast surface or by secretion of soluble CD318 from the synovial fibroblasts is as yet unknown. Nevertheless, the chemotactic effects of soluble

CD318 resemble in some respects chemotactic properties of CD13, another membrane protein on synovial fibroblasts that also is present at high concentrations as a soluble molecule in inflammatory joint fluid[220]. Although CD13 does not show structural resemblance to

conventional chemokines, there is evidence that CD13, like classical chemokines signals through a G-protein coupled receptor[220]. Moreover, whether soluble CD318 chemotactic effect on T cells is dependent on CD6-mediated signaling remains to be investigated. Overall, these observations indicate that more rigorous future studies are warranted to validate the T cell chemotactic effect of soluble CD318. These issues are also addressed in the summary and future direction part 2 (Determine the contribution of CD6 to the chemotactic effect of soluble CD318 section).

Although biologic therapeutics have led to important improvements in the treatment of RA and JIA, these agents impair host defenses to various pathogens and do not selectively target molecular interactions that are more important in pathogenic autoimmunity compared to normal immune responses. Identification of CD318 as a ligand of CD6 creates a potential therapeutic target at the level of the T cell/synovial fibroblast interaction that is not relevant to T cell interactions with professional antigen- presenting cells in lymphoid organs. CD318 has been proposed as a novel molecular target for treatment of malignant neoplasms[140, 210, 221]; the realization that it is engaged by CD6 will create a new perspective from which to assess such possibilities.

Moreover, our new data could also prompt consideration of CD318 as a novel therapeutic target in autoimmune diseases.

CHAPTER FIVE Summary and Future directions

Summary and Future directions Part 1

In summary of the research work presented in chapter 3, we found that CD6-/- mice were protected from intestinal I/R-induced injury as shown by significantly attenuated histopathology of the intestine and reduced levels of local IL-6. In mechanistic studies, we found that total and PC-specific IgM titers were reduced in the serum of CD6-/- mice, and that reconstitution of CD6-/- mice with IgM from naïve WT mice significantly restored intestinal I/R-induced inflammation. In addition, we demonstrated that CD6 was selectively expressed on natural antibody producing-B1 cell outside of the peritoneal cavity and bone marrow. Finally, we found that CD6-expressing

B1 cells were significantly reduced in specific tissue compartments in the CD6-/- mice in association with reduced B1a cell proliferation. While this research work provides great insight into the role of CD6 on B1a cells and its contribution to intestinal I/R-induced injury, there are still several questions to be addressed.

Factors that induce/repress CD6 expression are unknown We found that the differential expression of CD6 contributes to the phenotypic differences between splenic and peritoneal B1 cells. While peritoneal B1 cells do not express CD6 in the peritoneal cavity, they acquire the expression of CD6 in the spleen.

Peritoneal B1 cells have been reported to also migrate to the lymph node and differentiate into IgM-secreting cells in response to a specific pathogen[72, 222]. It would be interesting to determine whether these peritoneal B1 cells in the lymph node also acquire similar CD6 expression level as those that enter the spleen. Characterizing the expression of CD6 on B1 cells in the lymph node is crucial in determining whether CD6 is truly restrictive to splenic B1 cells. Additionally the tissue-specific signal that induces

expression of CD6 on B1 cells in the spleen remains unknown. An alternate hypothesis for the lack of CD6 expression on B1 cells in the peritoneal cavity is that the activation of

B1 cell represses CD6 expression.

The distinct basal activation state of peritoneal B1 cells is one of the major phenotypic differences between peritoneal and splenic B1 cells[76]. This unique peritoneal B1 cell basal activation state is induced by chronic antigen stimulation of the

B1 cells in the peritoneal cavity. This suggest a model in which B1 cells entering the peritoneal cavity lose CD6 expression and acquire CD11b expression as they transition into the distinct activation state typically associated with B1 cells in the peritoneal cavity.

It would be interesting to determine if splenic B1 cells lose the expression of CD6 when stimulated in vitro or when adoptively transferred into the peritoneal cavity.

Additionally, the correlation between the loss of splenic CD6 expression and the acquisition of CD11b, CD80 and CD9 should also be investigated post-adoptive transfer.

Role of CD6 on peritoneal B1a cell homing/migration into peripheral tissues

The acquired expression of CD6 on peritoneal B1 cells outside in the peritoneal cavity could potentially be important for the effective homing of B1a cells to the splenic or lymph node naively or during infection. A similar role has been previously attributed to CD6 in the accumulation of memory B cells at the local site of inflammation in patients with sjogren syndrome, necessitating the need to further investigate the role of

CD6 and its ligand(s) interaction in the homing of B1 cells to secondary lymphoid organs.

Characterization of the elevated peritoneal B1a cells in the CD6-/- mice

In the peritoneal cavity of CD6-/- mice, we found a significant increase in the frequency of CD5+ B1a cells compared to WT mice. Recent studies utilizing serum IgM- deficient mice suggest that these accumulated CD5+ B1a cells are not B1a cells but rather an accumulation of short-lived anergic B2 cells. These anergic B2 cells express CD19,

B220, and CD5 but lack the expression of CD43. The observed elevated CD5+ B1a cells in the peritoneal cavity of CD6-deficient mice should be further characterized to determine if CD6 indirectly regulates the accumulation of anergic CD5+ B2 cells in the peritoneal cavity.

Contribution of CD6 to splenic B1 BCR hyper responsiveness is unknown

The BCR signaling responses of splenic and peritoneal B1 cells are very distinct.

Following BCR ligation, splenic B1 cells respond similarly as B2 cells[79, 82]. They display robust calcium mobilization unlike peritoneal B1 cells that are hypo responsive, however the molecular mechanism(s) that contribute to these phenotypic differences are poorly understood. CD6 was recently identified as important regulator of TCR signaling.

The cytoplasmic domain contains several tyrosine and serine residues that interact with signaling transducing effector such as SLP-76[106]. Deficiency of CD6 on T cells results in exacerbated calcium mobilization in response to TCR-crosslinking[112]. The restricted expression of the scavenger receptor CD6 on splenic B1 cells suggest that perhaps CD6 may be a component of the signaling mechanism that confer the BCR hyper responsiveness typical of CD5+ splenic B1 cells. In this study, we investigated the effect of CD6-deficiency on basal splenic B1a cell signaling, It would interesting to determine if CD6-deficiency alters splenic B1 BCR signaling following engagement of the BCR.

Furthermore, a future study addressing the effect of CD6-overexpression on peritoneal

B1 BCR hypo responsiveness should be investigated in order to address CD6 role in modulating the signaling and cellular fate of B-1 cells.

Effect of CD6-deficiency on B1a cell development

Our data demonstrate a novel function of CD6 in regulating the self-renewal of

CD6-expressing B1a cells. It remains to be determined as to whether the reduced B1a cell compartment is intrinsic to genetic defects in B1a precursor cells or due to CD6- deficiency in other cell types. It has been reported that cytokines such as IL-5 and IL-7 which can be secreted by CD6-expresing cells such as T cells are critically important for

B1 cell homeostasis[223-225]. Therefore a conditional knockout of CD6 in B cells needs to be generated in order to fully exclude an indirect effect of CD6 –deficiency on the reduced B1a cell population. During fetal development, B1 cell progenitors in the fetal liver give rise to mature B1a cells. Although we found that neonatal B1a cells in the liver express CD6, the expression of CD6 on B1 cell progenitors needs to be investigated.

Characterization of CD6 expression on B1 cell progenitors is essential in order to explore the possibility that impaired differentiation/expansion of B1 cell progenitors potentially may have contributed to the decreased frequency/numbers of B1a cells in the liver of neonatal CD6-/- mice.

Effect of CD6-deficiency on splenic B1a cell ability to secrete IgM

We attributed the reduced total IgM titers in the CD6-/- mice to lower splenic B1a cell population in the CD6-/- mice. However whether the CD6-deficiency has any effect on the ability of splenic B1a cells to secrete IgM on a per cell basis was not investigated

and should be explored in future studies. The small frequency of CD6-expressing splenic

B1a cells has made it technically challenging to isolate these in vitro and assess their ability to secrete IgM on a per cell basis. Recently it has been reported that splenic B1a cells expressing CD138 spontaneously secrete more IgM in naïve mice compared to non-

CD318 expressing splenic B1a cells[177]. Additionally, these splenic B1a CD318 expressing cells contribute a significant portion of the natural IgM in naïve mice. The identification of splenic B1a cells that express CD318 raises the question of whether CD6 cell surface expression differ between CD138+ and CD138- B1a cells, and whether CD6- deficiency alters the frequency and population of CD138+ splenic B1a cells.

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Summary and Future directions Part 2 In summary of the research work presented in chapter four, we identified abundant CD318-derived peptides in mAb 3A11-immunoprecipiated proteins. CD318 also met the previously published criteria of a 3A11 antigen suggesting that CD318 is the antigen recognized by mAb 3A11. We confirmed the MS results by probing the mAb

3A11-immunoprecipiated proteins with a commercial anti-CD318 mAb, and probing recombinant CD318 with mAb 3A11 in Western blots. In addition, we found that mAb

3A11 and the established CD318 mAb have the same staining patterns on cells known to naturally express or lack CD318, and on cells engineered to upregulate or downregulate

CD318. Using soluble CD6, soluble CD318, cells express both CD166 and CD318 or

CD318 alone in flow cytometric analyses and pull-down experiments, we confirmed that

CD318 does bind to CD6.

Finally, we found that CD318 is expressed on synovial fibroblasts from RA patients after stimulation and that levels of CD318 are significantly higher in synovial fluids from RA and JIA patients than those from OA patients. Moreover, a soluble form of CD318 can be readily detected in synovial fluids from patients with inflammatory

(RA, JIA) but not non-inflammatory (OA) arthritis, and these levels are appreciably higher than serum levels of soluble CD318 in RA or healthy controls. Of particular interest, both the membrane-bound and soluble forms of CD318 are functionally active in vitro, in adhesion and chemotaxis assays, respectively, that are arguably relevant to components of the pathogenesis on joint inflammation in vivo. While this research work provides great insight into the identification of CD318 as a novel CD6 ligand and its role as a potential biomarker for RA, there are still several questions to be addressed.

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Determine the structural domains that mediate CD6/CD318 interaction and the biological effect of this interaction

We identified CD318 as novel CD6 ligand, however the specific extracellular domain of CD6 that directly interacts with CD318 is unknown. We hypothesize that CD6 interacts with CD318 via its membrane distal domain (domain-1) and preliminary

experiment using the T12-mAb (IgM mAb for CD6 domain 1) supports this hypothesis.

T12-mAb has been proposed to either block the interaction of CD6 and a novel ligand that binds to domain 1 or non-specifically block cell-cell adhesion due to its relatively large size[137, 138]. Therefore to validate our preliminary results, several deletion mutants of

CD6 extracellular domains should be generated and their binding to rCD318 should be assessed. Additionally the extracellular domain of CD318 that directly binds to CD6 is also unknown and should be investigated in future studies.

Identifying the structural domains that mediated CD6/CD318 interaction is crucial in understanding the biological effects of CD6/CD318 ligation on lymphocyte and epithelial cell activation/function. Ligation of CD6 using CD6 mAbs or recombinant

ALCAM has been reported to induce MAPK signaling cascade in T cells[105]. The

MAPK signaling pathway has been implicated in a variety of T-cell function. The effect of rCD318 binding to CD6 expressing T cells on these signaling pathways should also be investigated. Crosslinking of CD318 CUB1 domain using CD318 specific mAb has been reported to induce Src phosphorylation of CD318 cytoplasmic tyrosine residues resulting in PKC δ activation[117]. Whether crosslinking of the other CUB-domains can also initiate CD318-mediated signaling is currently unknown. Mapping the extracellular

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domain of CD318 that binds to CD6 will be very important in determining whether CD6 binding to CD318 can potentially initiate CD318-mediated intracellular signaling.

Determine the contribution of CD6 to the chemotactic effect of soluble CD318

During the progression of RA, effector T cells and other immune cells are recruited from circulation into the synovial, where they secrete proinflammatory cytokines that contributes to the pathogenesis of RA[131]. We found that CD318 is elevated in the synovial fibroblast of RA patients compared to healthy controls and the levels of soluble CD318 are also elevated in the synovial fluid of RA and JIA patients.

Additionally we show that these elevated levels of soluble CD318 may potentially recruit

T cells into the synovium. CD318 has been reported to interact with a variety of cell surface molecules, which may be expressed on T cells. Whether the chemotactic effect of soluble CD318 on T cells is entirely dependent on CD6-mediated signaling in response to

CD318 binding remains to be addressed. Assessing the chemotactic effect of soluble

CD318 on non-CD6 expressing immune cells will also help in determining the contribution of CD6 to the chemotactic effect of soluble CD318. The use of antagonist to and mice-deficient in CD318 will be essential for defining the role of CD318 in the pathogenesis of RA.

Determine the role of CD318 as a potential biomarker in other autoimmune diseases

In addition to RA, CD6 has been a therapeutic target for the treatment of MS and we found that CD6-deficient mice are protected from experimental autoimmune encephalomyelitis (EAE), a mouse of MS[226]. Additionally, we also observed that

CD318 is expressed on human brain microvascular endothelial cells and this expression

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is enhanced with IFN-gamma stimulation. Preliminary data suggest that levels of CD318 are elevated in the cerebral spinal fluid of MS patients compared to control, however the samples size were very small. The levels of soluble CD318 in the CSF should be investigated with a larger sample size to determine if CD318 is also a potential biomarker for the diagnosis and/or treatment of patients with MS. The significance of soluble

CD318 in the CSF in respect to T cell transmigration across the blood brain barrier as well as local T cell activation within the CSF should also be investigated in future studies.

Determine the immunological role of CD318 in T cell development and maintenance of thymic epithelial compartment

Preliminary studies indicate that mouse immune cells do not express CD318 naively or upon activation. The naïve expression of CD318 seems to be restricted to epithelial cells and we found that CD318 is expressed on thymic epithelial cells. The dynamic interaction between developing thymocytes and thymic epithelial cells are essential for the development of both the T cell and thymic epithelial compartment[227,

228]. Thymic epithelial cells regulate T cell differentiation into mature T cells by interrogating developing T cell self-reactivity; in return T cells contribute to the development and maintenance of the thymic architectural compartment by secreting trophic factors[229]. Thymic epithelial cells are classified into two groups (medullary and cortical thymic epithelial cells)[230]. The characterization of CD318 expression among the thymic epithelial cell subsets in future studies will be critical in determining what potential roles CD318/CD6 interaction might play in T cell development

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(positive/negative selection) and in the maintenance of the thymic epithelial compartment.

Determine the role of CD318 on B1 cell development and positive selection for self- antigens

As reported in chapter 3, the deficiency of CD6 resulted in reduced natural IgM titers as well as titers of IgM specific to self-antigens such as PC and Annexin IV. BCR signaling strength and positive selection for self-antigens play an important role in B1 cell development. The reduced B1a cell population in the CD6-/- mice suggests that CD6- mediated signaling perhaps in response to CD6 and its ligand(s) interactions contributed to the overall BCR signaling threshold essential for B1a cell development and /or positive selection for self-antigens. Preliminary data indicates that CD318-deficiency does not alter the titers of total IgM in the serum of naïve mice indicating that perhaps the B1a cell population may not affected, however this warrants further investigation. The titers of

PC-specific IgM was found to be reduced in the CD318-/- mice indicating a possible role in the positive selection of B1 cells given that the titers of total IgM were not reduced.

The titers of IgM-specific for other self-antigens such as Annexin IV, Myosin IIA and oxidized lipids should be investigated in future studies in order to fully address the role of

CD6-CD318 interaction in B1 cell positive selection.

Determine potential CD318 mediated functions that are independent of CD6

CD318 has been reported to interact with other cell surface molecules, however the biological significance of those interactions has not been reported. These previous findings suggest that CD318 may have CD6-independent functions. In support of this

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hypothesis, we found that the titers of total IgG are reduced in the serum of CD318-/- mice, a phenotype that was not found in the serum of CD6-/- mice. Neutrophils have been reported to contribute to the production of total IgG by stimulating B cells via secretion of cytokines[231]. The neutrophil population in the lymph node of CD318-/- mice is significantly reduced, while those in circulation are unaffected. The effect of CD318- deficiency on the B cell compartment needs to be characterized in future studies in order to understand the contribution of neutrophil/B cell interaction to the reduced IgG titers in the serum of naïve CD318-/- mice.

The reduction of the neutrophil population in the lymph node suggests perhaps

CD318 is critically important for the homing of neutrophils into the lymph node. CD6 is not expressed on neutrophils and the homing of neutrophils to the lymph node has been reported to be dependent on CD11b[232]. It would interesting to determine if CD11b mediated homing of neutrophils to the lymph node is dependent on CD318. Finally, an extensive analysis of both CD318 and CD6-deficient mice will be crucial in determining

CD318-mediated immune functions that are independent and dependent on CD6.

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