SLIT/ROBO SIGNALING IN MONOCYTE CHEMOTAXIS AND FUNCTION: A ROLE IN VASCULAR INFLAMMATION

by

Ilya M. Mukovozov

A thesis submitted in conformity with the requirements for the degree of Master of Medical Science

Institute of Medical Science

University of Toronto

© Copyright by Ilya M. Mukovozov. 2010.

ii

Slit/Robo Signaling in Monocyte Chemotaxis and Function: A Role in Vascular Inflammation.

Ilya M. Mukovozov

Master of Medical Science (MSc)

Institute of Medical Science

University of Toronto

2010

ABSTRACT

Vascular inflammation and associated leukocyte influx is a hallmark in the pathogenesis of

atherosclerosis. In both animal models and human subjects, inhibiting monocyte

recruitment is beneficial in preventing atherosclerosis and its clinical manifestations. The

trafficking signals that recruit cells to areas of inflammation are provided by small secreted

proteins called chemokines. Chemokines play a major role in the pathogenesis of

inflammation, and redundancy among the chemokine signaling pathways means that

blocking one pathway could result in another assuming its function. Therefore, we aim to

block a cell’s response to a range of chemokine-induced directional migration signals. Slit2

treatment inhibits monocyte migration in vitro using transwell migration assays, and in vivo,

using a murine peritonitis model of inflammatory cell influx. This inhibition is shown to be

dose- and time- dependent. Furthermore, Slit2 inhibits monocyte adhesion to activated

endothelial cell monolayers. These data may suggest a therapeutic role for Slit2 in

atherosclerosis.

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ACKNOWLEDGEMENTS

I am most grateful to my supervisor and mentor Dr. Lisa Robinson for her

constant encouragement and guidance through these past years. Lisa’s

extraordinary passion for science, her positive energy and commitment to motivate

young minds through the Kids’ Science program, have been a constant source of

inspiration. Her patience and understanding have made my transition and

subsequent experience in the lab very enjoyable. I would also like to thank the

members of my program advisory committee, Dr. Sergio Grinstein and Dr. Thomas

Waddell for their encouragement, advice, and constructive criticism. I am especially

lucky to have worked with the incredible individuals that make up the Robinson lab,

and am thankful for their continued support. In particular, I am grateful to Dr. Yi Wei

Huang and Guang Ying Liu for their patience with me when I started working in the

lab. I would also like to thank Sajeda Patel and Dr. Swasti Chaturvedi whose

suggestions, feedback and advice made my experience all the more rewarding. I

would also like to thank all the members of Dr. Grinstein’s lab and Dr. Brumell's lab,

for their technical help, constructive criticism and friendship. I would also like to

acknowledge the friendly and welcoming environment on the 4th floor, especially

Shahab Shahnazari and the rest of the bear pack, which brought joy and laughter to

my day-to-day experiences. I would also like to thank Thomas Sabljic, Danielle

Baribeau and Stephanie Byun for their support and motivation. Finally, I would like

to thank my parents, my sister Irina and Buddy, for their love, dedication and

support throughout my life.

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DATA ATTRIBUTION

The data presented here was performed in collaboration with a number of

individuals. The purification of Slit2 was performed by Dr. Durocher. Dr. Huang and

Sajeda Patel were responsible for performing the experiments presented in Fig. 3.3.

In addition, Guang Ying Liu, Shirin Chahtalkhi and I performed the immunoblotting

experiments presented in figure 3.4, while I performed the subsequent analysis. Min

Rui-Crow performed the experiments presented in Fig. 3.10. Finally, the data

presented in Appendix 1 represents a study that was initiated by Soumitra Tole and

completed by me. We contributed equally to this work.

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TABLE OF CONTENTS

LIST OF FIGURES vii LIST OF ABBREVIATIONS viii CHAPTER 1: INTRODUCTION

1.1 Inflammation 1 1.1.1 Leukocyte Trafficking and the Adhesion Cascade 1 1.1.2 Chemoattractants 4

1.2 The Monocyte 7 1.2.1 Chemotaxis 9 1.2.2 Adhesion 15 1.2.3 Phagocytosis 18 1.2.4 Monocytes and Vascular Inflammation 19

1.3 Slit2: A Guidance Cue for Cell Migration 21 1.3.1 Expression 24 1.3.2 Slit and Robo Structure 24 1.3.3 Slit2/Robo-1 Intracellular Signal Transduction 27 1.3.4 Slit/Robo in Leukocyte Trafficking 28

1.4 RHO GTPases: Rac and Cdc42 29 1.4.1 Structure and Regulation 30 1.4.2 Role of Rho GTPases in the regulation of the actin cytoskeleton 32

1.5 Rationale, Hypothesis & Objectives 33 1.5.1 Rationale 33 1.5.2 Hypothesis 35 1.5.3 Objectives 35

CHAPTER 2: MATERIALS & METHODS

2.1 Reagents and Antibodies 39 2.2 Isolation of Human Monocytes 39 2.3 Cell Culture 39 2.4 Slit2 Expression and Purification 40 2.5 Immunofluorescence 41 2.6 Transwell Migration Assay 41 2.7 Immunoblotting 42 2.8 Cdc42 & Rac2 Activation Assays 43 2.9 Adhesion 44 2.10 Murine Peritonitis 45 2.11 Phagocytosis 46 2.12 Statistical Analysis 46 CHAPTER 3: RESULTS

3.1 Monocytes express the Slit2 receptor, Robo-1. 48 3.2 Slit2 inhibits chemotaxis of human monocytic THP-1 cells. 48 3.3 Slit2 treatment inhibits activation of Rac2 and Cdc42. 49 3.4 Slit2 inhibits Akt and Erk, but not p38 MAPK pathways. 51

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3.5 Slit2 inhibits adhesion of monocytic THP-1 cells to activated human umbilical vein endothelial cell and human arterial endothelial cell monolayers.

52

3.6 Slit2 inhibits monocyte recruitment in vivo. 53 3.7 Slit2 does not alter monocyte phagocytosis. 55 CHAPTER 4: DISCUSSION & CONCLUSIONS

78

REFERENCES

89

APPENDIX1: The Axonal Repellent, Slit2, Inhibits Directional Migration of Circulating Neutrophils

A1.1 Abstract 123 A1.2 Introduction 124 A1.3 Materials and Methods 129 A1.4 Results 142 A1.5 Discussion 148 A1.6 Acknowledgments 155 A1.7 Authorship 156 A1.8 References 157 A1.9 Figure Legends 166

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LIST OF FIGURES Page

Figure 1.1 Leukocyte Adhesion Cascade. 2

Figure 1.2 Intracellular signal transduction upon chemokine GPCR activation. 13

Figure 1.3 Rho GTPases in the polarized monocyte. 16

Figure 1.4 Primary Structure of Mammalian Slit2 and Robo-1 Proteins. 25

Figure 3.1 Robo-1 is expressed by monocytes. 52

Figure 3.2 Slit2 inhibits monocyte chemotaxis. 54

Figure 3.3 Slit2 inhibits activation of Rho GTPases (Cdc42 and Rac2). 56

Figure 3.4 Slit2 inhibits Akt and Erk but not p38 MAPKs. 59

Figure 3.5 Slit2 inhibits adhesion of monocytic THP-1 cells to human umbilical vein endothelial cells.

62

Figure 3.6 Slit2 inhibits adhesion of monocytic THP-1 cells to human arterial endothelial cells.

64

Figure 3.7 Slit2 inhibits monocyte recruitment in vivo. 66

Figure 3.8 Slit2 dose-dependently inhibits monocyte recruitment in vivo. 68

Figure 3.9 Slit2 inhibits monocyte recruitment in vivo: time-course. 70

Figure 3.10 Slit2 does not affect RAW 264.7 macrophage phagocytosis. 72

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LIST OF ABBREVIATIONS AKT Protein Kinase B CCL5 RANTES CC-chemokine Ligand 5 CNS Central Nervous System CR1 Complement Receptor 1 CXCL4 CXC Chemokine Ligand 4 CXCR4 CXC chemokine Receptor 4 DAG Diacylglycerol DAPI 4’,6-diamidino-2-phenylindole DCs Dentritic Cells DH Dbl homology domain ECM Extracellular Matrix EGF Epidermal Growth Factor ERK Extracellular-signal Regulated Kinase Ena Enabled EPAC Exchange Factor Directly Activated by Cyclic AMP fMLP N-formyl-methionyl-leucyl-phenylalanine FKN Fractalkine GAP GTPase Activating Protein GDNF Glial Cell Line-derived Neurotrophic Factor GEF Guanine Nucleotide Exchange Factor GPCR G-Protein-Coupled Receptor GTPase Guanosine Triphosphatase HAEC Human Arterial Endothelial Cells HRP Horseradish Peroxidase HUVEC Human Umbilical Vein Endothelial Cells ICAM-1 Intercellular Adhesion Molecule 1 Ig Immunoglobulin IP3 Inositol (1,4,5)-triphosphate ITAM Immunoreceptor Tyrosine Activation Motif LFA-1 Lymphocyte Function Associated Antigen 1 LRR Leucine-Rich Repeat Mac-1 Macrophage Receptor 1, αMβ2-integrin MAPK Mitogen Activated Protein Kinase MCP-1 Monocyte Chemotactic Protein 1 Mena Mammalian Enabled MLK Myosin Light-chain Kinase OxLDL Oxidized Low Density Lipoprotein PAK1 P21-Activated Kinase PBD P21-Binding Domain PBS Phosphate Buffered Saline PDGF Platelet-Derived Growth Factor PFA Paraformaldehyde PH Pleckstrin Homology PI3K Phosphoinositide 3-kinase

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PI(4,5)P2 Phosphatidylinositol (4,5)-bisphosphate PI(3,4,5)P3 Phosphatidylinositol (3,4,5)-trisphosphate PLC Phospholipase C PKC Protein Kinase C PKC-ζ Protein Kinase C-ζ PSGL1 P-selectin Glycoprotein Ligand 1 SDF-1 Stromal Cell-derived Factor-1 SPA1 Signal-Induced Proliferation Associated Antigen 1 VCAM-1 Vascular Cell Adhesion Molecule VEGF Vascular Endothelial Growth Factor VLA-4 Very Late Antigen 4 VVO Vesiculo-Vacuolar organelles THP-1 Human Acute Monocytic Leukemia Cell Line TNF-α Tumor Necrosis Factor α

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CHAPTER 1

INTRODUCTION

1.1 INFLAMMATION

1.1.1 Leukocyte Trafficking and the Adhesion Cascade

An essential function of the inflammatory response is to selectively recruit the

appropriate subsets of leukocytes to specific sites of inflammation. Leukocytes are

recruited to sites of inflammation in a series of coordinated interactions with

endothelial cells lining the vessel wall. The classical leukocyte adhesion cascade

involves three main steps: leukocyte rolling, activation and arrest, and

transmigration (Fig. 1). In the first step, circulating leukocytes are captured by

selectin-mediated rolling. Rolling is mediated by L-selectin expressed on most

leukocytes, as well as P-selectin and E-selectin, which are expressed by endothelial

cells (Kansas, G., 1996). All of the selectins interact with P-selectin glycoprotein

ligand 1 (PSGL1), although other glycoprotein ligands exist (McEver et al., 1997).

The binding of leukocyte L-selectin to PSGL1 can facilitate secondary leukocyte

capture by adherent leukocytes (Eriksson et al., 2001). The interactions of selectins

with their ligands allows leukocytes to adhere to inflamed endothelium under flow

(Alon et al., 1995). In fact, shear stress is required to support L-selectin and P-

selectin adhesion, as the rolling cells detach if flow is stopped (Finger et al., 1996;

Lawrence et al., 1997). This selectin-mediated slow rolling allows the leukocyte to

sample the repertoire of chemokines and other activation signals presented on the

luminal surface of endothelial cells.

2

Figure 1.1 Leukocyte Adhesion Cascade.

Leukocytes are recruited to sites of inflammation in a series of coordinated

interactions with endothelial cells (ECs) lining the vessel wall. The classical

leukocyte adhesion cascade involves three main steps: leukocyte rolling, activation

and arrest, and transmigration. ECM Extracellular Matrix.

3

In addition to selectins, various integrins participate in rolling. Monocytes can

roll on immobilized vascular cell-adhesion molecule 1 (VCAM-1) by engaging

integrin receptor very late antigen 4 (VLA-4; α4β1-integrin). Members of the β2-

integrins can also support rolling. Resting mouse neutrophils roll on surfaces coated

with E-selectin ligand and intercellular adhesion molecule 1 (ICAM-1). Ligation of E-

selectin induces a conformational change in lymphocyte function-associate antigen

1 (LFA-1; αLβ2-integrin) which allows it to bind to its ligand ICAM-1 (Salas et al.,

2004). In addition, it has recently been demonstrated that the mechanochemical

design of LFA-1 allows shear stress to induce and maintain a state of high ligand-

binding affinity (Astrof et al., 2006). Rolling in vivo was shown to require E-selectin

(Kunkel et al., 1996) and engagement of β2-integrins (Jung et al., 1998), LFA-1 and

macrophage receptor 1 (MAC1; αMβ2-integrin) (Dunne et al., 2002).

Subsequently, leukocytes undergo integrin-dependent arrest. This is

mediated by the binding of leukocyte integrins to immunoglobulin superfamily

members ICAM-1 and VCAM-1 on endothelial cells and is rapidly triggered by the

binding of chemokines and other chemoattractants (Campbell et al., 1996;

Campbell et al., 1998). These chemokines are secreted by activated endothelial

cells, although platelets can also deposit chemokines, such as CC-chemokine

ligand 5 (CCL5; RANTES) and CXC-chemokine ligand 4 (CXCL4) and CXCL5, onto

the inflamed endothelial lumen to trigger monocyte arrest (von Hundelshausen et

al., 2001; Huo et al., 2003). Finally, following firm arrest, leukocytes transmigrate

across the endothelial cell barrier, its associated basement membrane, and the

pericyte sheath. Leukocytes can cross the endothelium either between adjacent

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endothelial cells (paracellular route) or directly through an endothelial cell

(transcellular route). Transcellular migration generally occurs in 'thin' parts of the

endothelium where there is less distance for the leukocyte to migrate. In addition,

caveolae containing ICAM-1 link together to form vesiculo-vacuolar organelles

(VVOs). This creates a channel inside the cell through which leukocytes can

migrate.

Although the leukocyte adhesion cascade has been divided into several

steps, these are not temporally exclusive, but instead work together to achieve the

desired effect of leukocyte arrest and diapedesis. Although leukocyte diapedesis

was described almost 200 years ago, its molecular mechanisms are only now

beginning to be fully understood (Imhof et al., 2004). In the past decade, new

insights have been gained into the signalling events that underlie integrin activation,

post-adhesion strengthening of leukocyte attachment, and the molecules involved in

diapedesis (Muller, W., 2003).

1.1.2 Chemoattractants

In vivo, there are several types of chemoattractant mediators that can recruit

leukocytes to inflammatory foci. These include bacterial components, leukotrienes,

complement factors and chemokines. C5a was the first chemoattractant to be

identified, and it is a cleaved product derived from complement component C5 (Shin

et al, 1968). Bacterial products such as fMLP (N-formyl-methionyl-leucyl-

phenylalanine) and other N-formylpeptides can also act as general

chemoattractants that non-specifically recruit leukocytes to inflammatory foci.

5

However, the main chemoattractants that specifically recruit leukocyte subsets to

inflammatory foci are a family of chemoattractant cytokines called chemokines.

Chemokines are a large family of small peptides that are structurally similar (Rossi

et al., 2000). Most are secreted, while only two, fractalkine (FKN;CX3CL1) and

CXCL16, are expressed on the cell surface. The physiological importance of FKN

expression can be highlighted in studies of cardiac allograft rejection where FKN

expression is negligible in non-rejecting cardiac isografts, but is significantly

enhanced in rejecting allografts (Robinson et al., 2000). In addition, FKN-/- mice

have reduced atherosclerosis compared to wild type littermates (Robinson et al.,

2000). Chemokine-induced signal transduction pathways are very similar. Thus, it is

the specific expression, regulation, and receptor binding patterns of each

chemokine that determine functional diversity. There are four families of

chemokines that are classified on the basis of the relative positions of their N-

terminal cysteine residues (Luster, A.,1998). Most chemokines contain four cysteine

residues and fit within the α (CXC) or the β (CC) chemokine families, although

another two families exist with lone members. FKN is a lone member in its family,

and its N-terminal cysteine residues are separated by three amino acids (CX3CL1).

The fourth family is composed only of lymphotactin (XCL1), which is a lymphocyte

specific chemokine (Kennedy et al., 1995). Most chemokines bind to

glycosaminoglycans (GAGs) on the luminal surface of endothelial cells. This binding

is required for leukocyte recruitment, since chemokines with mutations in their GAG

binding domains can induce in vitro chemotaxis, but are unable to recruit leukocytes

to the peritoneal cavity in vivo (Johnson et al., 2005).

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The binding of chemokines to their heterotrimeric G-protein coupled

receptors (GPCRs) activates leukocyte integrins instantaneously by inside-out

signalling mechanisms (Shamri et al., 2005). They rapidly regulate integrin avidity

by increasing both integrin affinity (by a conformational change that results in

increased ligand binding energy and a decreased ligand dissociation rate), and

valency (the density of integrins per area of plasma membrane involved in

adhesion, determined by expression levels and lateral mobility) (Laudanna et al.,

2002; Constantin et al., 2000). It is through these signaling mechanisms that

chemokines act as powerful activators of integrin-mediated adhesion and leukocyte

recruitment.

In monocytes/macrophages, chemokines interact with specific serpentine

(heptahelical) receptors on the plasma membrane, which transduce signals by

coupling to heterotrimeric G proteins. Heterotrimeric G proteins are composed of an

α, β, and γ subunit. The α subunit is the GDP/GTP binding element. When bound to

GDP, the α subunit interacts with the β and γ subunits to form an inactive

heterotrimer complex that binds to the serpentine receptor. Binding of the

chemokine to the serpentine receptor induces a conformational change that causes

the exchange of GDP for GTP on the α subunit. This results in the dissociation of

the α subunit from the receptor and the release of the Gβγ complex. The free Gα

and Gβγ subunits are then free to interact with and modulate the activity of target

enzymes. Thus, chemokine binding induces the dissociation of the G protein

complex into α and βγ subunits, which bind and activate target enzymes such as

phosphatidylinositol 3-kinase (PI 3K), phospholipase C (PLC), or adenyl cyclase.

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These enzymes play an important role in generating secondary intracellular

messengers that initiate a cascade of signaling events that culminate in cytoskeletal

mobilization required for the chemoattraction response.

1.2 The Monocyte

Mononuclear phagocytes arise in the bone marrow from dividing monoblasts

and are released into systemic circulation as nondividing monocytes (Wiktor-

Jedrzejczak et al., 1996). They circulate for several days before entering tissues

and replenishing resident macrophages and dendritic cells (DCs) (Akagawa et al.,

1996; Chapuis et al., 1997; Randolph et al., 1998). As half of the circulating

monocytes leave the bloodstream under steady-state conditions every day,

monocytes constitute a large systemic reservoir of myeloid precursors. Although

circulating monocytes give rise to tissue-resident macrophages, they also form

specialized cells such as DCs and osteoclasts, making up the mononuclear

phagocyte system (Hume et al., 2002). As precursors for microglia and osteoclasts,

monocytes play a role in the physiology of the central nervous system and in bone

remodeling (Servet-Delprat et al., 2002). Mononuclear phagocytes are important for

both innate and adaptive immunity. Their interactions with antigen-specific T

lymphocytes trigger the induction of adaptive immune responses (Geissmann et al.,

2003). Monocytes, defined as blood mononuclear cells, have "bean-shaped" nuclei

and express CD11b, CD11c, and CD14 in humans, and CD11b and F4/80 in mice

(Muller, W., 2001). Circulating monocytes are morphologically heterogeneous and

constitute approximately 5-10% of peripheral blood leukocytes (van Furth et al.,

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1968). In humans and mice, the two principal monocyte subpopulations are the

"inflammatory" and "resident" subsets (Geissmann et al., 2003). Human

inflammatory monocytes express high levels of L-selectin and several chemokine

receptors, including CCR2 the CCL2 receptor, but low levels of CX3CR1, the FKN

receptor. On the other hand, resident human macrophages express high levels of

CX3CR1 and low to non-detectable levels of L-selectin and most chemokine

receptors, such as CCR2 (Grage-Griebenow et al., 2001). Furthermore, in humans

CD14high CD16ˉ monocytes represent the inflammatory subtype, while CD14low

CD16+ monocytes represent the resident subtype (Ziegler-Heitbrock, H., 1996). In

mice, inflammatory monocytes are characterized by expression of high levels of Ly-

6C, a glycosylphosphatidylinositol-linked cell surface protein with unknown function.

Ly-6Chigh, or "inflammatory", murine monocytes are short lived under steady-state

conditions, and are preferentially recruited to inflammatory foci, such as

atherosclerotic lesions (Sunderktter et al., 2004; Tacke et al., 2007). Ly-6Chigh

monocytes express higher levels of PSGL-1 than Ly-6Clow, or "resident" monocytes,

and thus roll at slower velocities on P-selectin and E-selectin substratum. This

property allows Ly-6Chigh monocytes to interact preferentially with atherosclerotic

endothelium, compared with Ly-6Clow monocytes (An et al., 2008). Resident

monocytes in mice express low to non-detectable levels of Ly-6C, persist longer

and repopulate several tissues, such as the liver, lung, spleen and brain after

adoptive transfer (Geissmann et al., 2003). Interestingly, Ly-6Clow CX3CR1high

monocytes show unique patrolling behavior in mice that are deficient in natural killer

cells and T lymphocytes, and are readily recruited to sites of infection (Auffray et al.,

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2007). Although monocytes routinely emigrate from the blood to replenish tissue

macrophages, increased recruitment can be elicited by pro-inflammatory, metabolic,

and immune stimuli (van Furth et al., 1985). Following recruitment, monocytes

differentiate into macrophages and DCs, contributing to host defence and tissue

homeostasis through the clearance of senescent cells and tissue remodelling and

repair following inflammation (Gordon, S., 1986; Gordon, S., 1998). In addition to

host defense, monocytes have been implicated in atherosclerosis. The mechanisms

controlling monocyte functions will be outlined to better understand the role they

play in tissue homeostasis and host defense.

1.2.1 Chemotaxis

Chemotaxis, the directed cell migration towards external chemical gradients,

is a biological phenomenon of widespread occurrence. Chemotaxis can be

observed in many eukaryotic cells including: free-living organisms, leukocytes

(during inflammation), endothelial cells (angiogenesis), spermatocytes (fertilization)

and neurons (neurogenesis), highlighting the biological significance of this

phenomenon (Singer et al., 1986). Monocytes are amoeboid cells that move by

extending pseudopods. Chemotaxis is achieved by first polarizing or orienting the

cell in the direction of locomotion along a chemoattractant gradient. Polarization

results from preferential pseudopod extension towards areas of higher

chemoattractant concentration (Zigmond, S., 1974). Efficient chemotaxis requires

coordination of motile activities such as pseudopod formation at the leading edge of

the cell, and uropod retraction at the trailing edge.

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During chemotaxis, macrophages extend short surface protrusions called

filopodia, or microspikes, which are membrane extensions of approximately 0.1-0.2

µm in diameter and up to 20 µm in length. These structures act as cellular tentacles

and are supported by a core bundle of actin filaments called microfilaments (Mattila

et al., 2008). In macrophages, filopodia help to support thin sheets of membrane-

enclosed cytoplasm, called lamellipodia. Lamellipodia contain actin filaments and a

meshwork of myosin II-associated microfilaments. In macrophages, as well as in

other cell types, the actin network within the lamellipodia, in association with several

structural and regulatory proteins, comprises the molecular motor which drives cell

locomotion (Jones et al., 1998). This locomotory apparatus works against cell-to-

substratum adhesions called focal contacts or focal adhesions. These molecular

structures utilize members of the integrin family of proteins to link the myosin II-

containing bundles of cytoplasmic microfilaments, called stress fibers, to proteins in

the extracellular matrix (ECM) (Critchley et al., 1999). In macrophages, integrin-

mediated contacts to the ECM take two forms: focal complexes and podosomes.

Focal complexes are structurally similar to focal adhesions but lack stress fibers

(Allen et al., 1997), while podosomes are distinct circular structures that are only

observed in cells of the myeloid lineage (DeFife et al., 1999; Correia et al., 1999;

Linder et al., 2003).

Macrophage chemotaxis can be divided into several steps: actin-driven

protrusion of filopodia and lamellipodia at the leading edge, adhesion of the leading

edge to the ECM via integrin-mediated focal interactions, actomyosin-mediated cell

contraction, release of contacts at the trailing edge of the cell, and recycling of

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membrane receptors from the rear to the front of the cell (Allen et al., 1998; Sheetz

et al., 1999; Friedl, P., 2004; Friedl et al., 2009). The cell must integrate a number of

molecular events in order to efficiently move across a substratum. This coordination

is required for polarization and chemotaxis, and is largely mediated by the actin

cytoskeleton within the cell.

Exposure of circulating monocytes to chemoattractants leads to activation

and subsequent migration of monocytes across the endothelial barrier towards the

inflammatory foci. Monocytes/macrophages are responsive to even minuscule

gradients of extracellular signals and will undergo chemotaxis towards a variety of

stimuli. These signals include chemoattractants such as fMLP, vascular endothelial

growth factor (VEGF) and chemokines such as macrophage chemotactic protein 1

(MCP-1;CCL2) and stromal cell-derived factor-1α (SDF-1α;CXCL12) (Gyetko et al.,

1994; Barleon et al., 1996). Binding of the chemoattractant to its cell-surface

receptor activates intracellular signaling cascades, which mobilize the actin

cytoskeletal machinery. Subsequently, the cell polarizes forming actin-rich filopodia

and lamellipodia at the 'front' of the cell and a tail-like uropod at the cell rear. The

generation and maintenance of cell polarity and actin cytoskeleton remodelling are

necessary processes for efficient monocyte chemotaxis.

The SDF-1α receptor (CXCR4), like other chemokine receptors, is a

glycosylated seven-transmembrane domain GPCR. It has an associated

heterotrimeric GDP/GTP binding protein complex made up of α, β and γ subunits.

Several mechanisms for activation of GPCRs have been proposed (Gether, U.,

2000; Ulloa-Aguirre et al., 1999). Small molecule agonists can bind within the

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transmembrane helices and cause receptor activation, while larger molecules bind

to the extracellular surface leading to a conformational change that is transmitted to

an intracellular Gαβγ complex. This event is followed by exchange of GTP for GDP

in the Gα protein and its subsequent dissociation from the Gβγ complex, followed

by the activation of downstream signaling pathways. Both the Gα subunit and Gβγ

complex interact with several downstream effectors to generate cell polarity and

drive migration, although these signaling events also prime the cell for other

immune functions. Furthermore, the various immune functions of the leukocyte are

not temporally exclusive as they are dependent on complex interactions between

intracellular signaling events.

Ligation of GPCRs leads to the activation of four major signaling pathways

(Fig. 2): PLC, PI3K, mitogen-activated protein kinases (MAPKs) and Rho guanosine

triphosphatases (GTPases). Each of these pathways is involved in cell activation

and/or the generation of cell polarity. Once the Gα subunit dissociates, the Gβγ

complex activates PLC, which in turn cleaves phosphatidylinositol (4,5)-

bisphosphate (PI(4,5)P2) to generate inositol (1,4,5)-triphosphate (IP3) and

diacylglycerol (DAG). Generation of IP3 leads to the mobilization of intracellular

calcium stores from the endoplasmic reticulum and DAG activates protein kinase C

(PKC) (Li et al., 2000).

A convincing role for PI3Ks in GPCR signaling and chemotaxis has been

established (Li et al., 2000; Sasaki et al., 2000; Hirsch et al., 2000; Servant et al.,

2000; Jin et al., 2000). Although there are at least four Class I PI3K isoforms in

mammalian cells (Vanhaesebroeck et al., 1999), only a single Class IB variant,

13

Figure 1.2 Intracellular signal transduction upon chemokine GPCR activation.

Chemokine binding to GPCRs induced a conformational change that results in the

dissociation of Gα subunits from the Gβγ complex. This leads to rapid outside-in

signaling resulting in the activation of four major signaling pathways that influence

monocyte chemotaxis, adhesion and phagocytosis: PLC, PI3K, MAPKs and Rho

GTPases. Each of these pathways contributes to the generation of cell polarity

and/or modulation of integrin avidity.

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containing a p110γ catalytic subunit complexed with a 101 kDa regulatory protein,

has been shown to interact with G-proteins in leukocytes. Since chemokine

responses in leukocytes, such as phagocytosis, are sensitive to pertussis toxin, it

was believed that chemokine receptors are coupled to a Gαi subunit (Boulay et al.,

1997). However, there is also data implicating other families of G-protein α subunits

in chemokine-mediated signaling (Amatruda et al., 1993), in addition to Gβγ

subunits (Clapham et al., 1993). While there is evidence that Class IB PI3K is

responsive to activation by Gα subunits (Murga et al., 1998), it has also been shown

that p110γ is activated via interactions with the Gβγ subunits (Neptune et al., 1997).

Regardless of how PI3K is activated, the outcome leads to phosphorylation of

membrane PI(4,5)P2 by activated PI3K, resulting in the generation of PI(3,4,5)P3 at

the plasmalemma.

In addition, the Gβγ complex also activates PI3Kγ which activates Src-family

kinases and generates PI(3,4,5)P3 from membrane PI(4,5)P2 (Krugmann et al.,

1999). Src-family kinases phosphorylate adapter proteins such as Shc, resulting in

the recruitment of Ras GTPases and subsequent activation of MAPK pathways

(Kintscher et al., 2000). Although the p38 and Erk MAPK pathways are involved in

chemotaxis and adhesion, the most important biochemical events for cell

polarization are the production of PIP3 and activation of Rho GTPases at the

leading edge of the cell.

The PI3K dependent production of PIP3 at the cell membrane allows for the

recruitment of Rho GTPases, Rac, and Cdc42 to the cell membrane. The

localization of PIP3, Rac and Cdc42 then stimulate the polymerization of actin, a

15

process necessary for the formation of lamellipodia at the front of the cell (Fig 1.3).

On the other hand, at the back of the cell, Rho-kinase phosphorylation results in

inactivation of myosin light chain phosphatase, leading to increased myosin light-

chain kinase (MLK) dependent activation of myosin (Nguyen et al., 1999). These

biochemical conditions favour the formation of actomyosin bundles, contraction, de-

adhesion from the substratum and tail retraction (Ridley, A., 2001; Bokoch, G.,

2005). Interestingly, mutual inhibition of leading edge and trailing edge proteins

allows for the maintenance of cell polarity (Fenteany et al., 2004). To prevent the

accumulation of PIP3 at the trailing edge, PTEN dephosphorylates PI(3,4,5)P3 to

PI(4,5)P2. The lack of PIP3 in the back of the cell decreases the activation and

recruitment of Rho GTPases and subsequent actin polymerization, allowing the

formation of actomyosin bundles and tail retraction (Worthylake et al., 2001). Actin

polymerization at the leading edge coupled to tail retraction in the back allows for

directed leukocyte chemotaxis.

1.2.2 Adhesion

Firm adhesion of leukocytes to activated endothelium is required for

leukocyte transmigration, and is an integrin-dependent process. Integrins are large

transmembrane glycoproteins that anchor the cell's cytoskeleton to other cells or to

the ECM. These large protein complexes are heterodimers composed of α and β

subunits. The β2 family of integrins, only expressed on leukocytes, includes LFA-1

and Mac-1. These are responsible for firm adhesion and arrest on endothelial cells,

16

Figure 1.3 Rho GTPases in the polarized monocyte.

The PI3K dependent production of PIP3 at the cell membrane allows for the

recruitment of Rho GTPases, Rac, and Cdc42 to the cell membrane. The

localization of PIP3, Rac and Cdc42 stimulates the polymerization of actin, a

process necessary for the formation of lamellipodia at the front of the cell. At the

back of the cell, Rho-kinase phosphorylation results in inactivation of myosin light

chain phosphatase, leading to increased myosin light-chain kinase (MLK)

dependent activation of myosin.

17

and later attachment with ECM components. LFA-1 and Mac-1 can both bind

VCAM-2 and ICAM-1 (Ley et al., 2007).

Chemokines are powerful activators of integrin-mediated firm adhesion.

GPCR stimulation results in rapid inside-out signaling cascades that result in a net

increase in the average integrin affinity and valency (Chan et al., 2003; Carman et

al., 2003). Although the signaling cascades downstream of GPCRs are incompletely

understood, several pathways have been at least partially elucidated. For example,

PLC is known to be recruited after GPCR signaling. Recruitment and activation of

PLC results in the production of IP3 and DAG followed by an increase in intracellular

calcium. This calcium flux and the production of DAG activates guanine nucleotide

exchange factors (GEFs), such as Vav1 and CALDAG1, and results in the

recruitment and activation of Rho GTPases (Constantin et al., 2000; Vielkind et al.,

2005; Crittenden et al., 2004). Thus, Rho GTPases are also involved in the

signaling cascades linking GPCR activation and changes in integrin affinity. For

example, chemokine-mediated conformational changes in LFA-1 are induced by

GTPases RhoA (Giagulli et al., 2004) and Rap-1 (Shimonaka et al., 2003). Once

recruited and activated, Rho GTPases associate with actin binding proteins,

including talin-1 and α-actinin, to modulate integrin affinity (Sampath et al., 1998;

Jones et al., 1998; Brakebusch et al., 2003). Thus, chemokine-mediated changes in

integrin affinity are dependent on interactions with the actin cytoskeleton.

In addition to GPCR-dependent inside-out signaling, binding of ligands to

integrins also induces outside-in signaling cascades (Ley et al., 2007). Paxillin is a

scaffold protein for signaling molecules that can bind integrins. Ligand induced

18

integrin clustering activates Src-family kinases. These phosphorylate paxillin

allowing the recruitment of downstream effectors, such as ADP-ribosylating factor

GTPase activating protein (ArkGAP) and PAK interacting exchange factor (PIX).

ArfGAP inactivates Rac, while PIX activates Cdc42 (DeMali et al., 2003). Src-family

kinases also activate Vav1, a Cdc42 and Rac GEF (DeMali et al., 2003). Thus, Rho

GTPases are required for the mobilization of the actin cytoskeleton to form and

maintain adhesive contacts. Both GPCR-induced inside-out signaling and ligand-

induced outside-in signaling modulate the activity of Rho GTPases to mobilize the

actin cytoskeleton for adhesion.

1.2.3 Phagocytosis

Phagocytosis is a cellular process in which solid particles are engulfed by the

cell membrane to form internal phagosomes. Although this process can be used for

the acquisition of nutrients and to clear dead cells and debris, in leukocytes, this is a

major mechanism used to clear invading microorganisms. Although phagocytosis of

uncoated particles can occur, the efficiency of phagocytosis is greatly enhanced by

the binding of opsonins, such as complement factors or Igs, to the surface of a

particle. Opsonins can be recognized by receptors on the leukocyte surface,

including Fc (Ravetch et al., 1991) and complement receptors (Brown, E., 1991),

which mobilize the actin cytoskeleton and aid in the subsequent internalization.

Fc receptors facilitate the engulfment of Ig coated particles. These receptors

are members of the multichain immune recognition receptor family. Their signaling

parallels signaling via T and B cell antigen receptors, which signal via homologous

19

cytoplasmic sequences, called immunoreceptor tyrosine activation motif (ITAM)

(Strzelecka et al., 1997; Imboden et al., 1985). Fc receptor activation results in

recruitment of PI3K, which converts PI(4,5)P2 to PIP(3,4,5)P3 and DAG. DAG is also

produced by PLC and activates PKC (Botelho et al., 2000). Phagocytosis requires

normal signaling through phosphoinositide kinases and PLC (Greenberg et al.,

2003). Ultimately, Fc receptor ligation leads to actin mobilization and membrane

remodeling, which is mediated by Rho GTPases. GEFs, such as Vav-1, contain a

pleckstrin homology (PH) domain with a high affinity for PI(3,4,5)P3, promoting their

recruitment to membrane PI(3,4,5)P3 at sites of Fc receptor ligation (Bustelo, X.,

2002). Recruitment of GEFs results in localized activation of Rho GTPases Rac and

Cdc42, which mobilize the actin machinery to extend pseudopods to engulf the

particle (Greenberg et al., 2002). For example, Fcγ phagocytosis results in

activation of the Rho GTPases Cdc42 and Rac (Caron et al., 1998).

Complement-derived opsonins, such as C3b, can be generated by both the

classical and the alternative pathways. Complement receptor 1 (CR1) on the

surface of leukocytes recognizes and binds C3b. Unlike the active phagocytosis

seen in Fc receptor-mediated engulfment, complement-mediated phagocytosis is

slow and gentle (Greenberg et al., 2002). The phagocytic ability of monocytes is

important for host defence. In addition, monocyte phagocytosis of oxidized low

density lipoprotein (OxLDL) is implicated in the formation of foam cells in

atherosclerotic lesions.

1.2.4 Monocytes and Vascular Inflammation

20

Atherosclerosis is an inflammatory disease that can be characterized by

intense immunological activity in the vessel wall. It is the major cause of coronary

heart disease (CHD) the leading cause of death in North America (Castelli et al.,

1984; Pasternak et al., 2004). The process of atherogenesis is also involved in

cerebrovascular atherosclerotic disease, and in the aortic artery, renal artery and

peripheral vasculature. Thus, the pathogenesis of atherosclerosis is wide ranging

and threatens human health worldwide (Murray et al., 1997). Atherosclerosis

involves the formation of vessel lesions, called plaques, that are characterized by

lipid accumulation, inflammation, cell death and fibrosis. Over time, these plaques

can mature and grow in size. The main complications of atherosclerosis result from

plaques that occlude the vessel, causing stenoses by limiting blood flow and

starving the downstream tissues of oxygen and nutrients. In some instances, the

plaque may rupture, sending an embolus down the vessel and exposing the

prothrombotic material in the plaque to the blood. This may also result in an abrupt

formation of a thrombotic clot, which can occlude the vessel at the site of plaque

rupture (Libby et al., 2002). In the coronary vessels, this can lead to a myocardial

infarction. Atherosclerosis in the carotid vessels can result in ischaemic stroke and

transient ischaemic attacks.

Atherosclerotic plaques are composed of a mixture of immune cells (mainly

macrophages and T lymphocytes), extracellular matrix, lipids and lipid-rich debris.

This accumulation of immune cells and lipids in the intima is located between the

vascular endothelial cells and the smooth muscle cells of the vascular media.

Although the cellular compositions of plaques changes with time, lipid-laden

21

macrophages , or foam cells, tend to outnumber other cell types. The immune cells

in the plaque become activated and produce proinflammatory cytokines such as

tumor necrosis factor α (TNF-α) and interferon-γ (IFNγ). This proinflammatory

environment induces the vascular endothelial cells to express increased leukocyte

adhesion molecules, allowing monocyte and other immune cells to transmigrate into

the plaque. In addition, vascular endothelial cells increase adhesion molecule

expression in response to cholesterol accumulation in the intima (Cybulsky et al.,

1991). Macrophage colony-stimulating factor (M-CSF) produced by endothelial cells

and smooth muscle cells (Rajavashisth et al., 1990) allows monocytes to

differentiate into macrophages in the plaque (Smith et al., 1995). In addition,

atherosclerotic plaques produce the chemokine FKN, which can be shed by

proteolysis. Shed FKN can engage its receptor CX3CR1 on blood-borne monocytes

and macrophages, stimulating their recruitment to the atherosclerotic vessel wall

(Combadire et al., 2003; Lesnik et al., 2003). Thus, the recruitment of immune cells,

especially monocytes, initiates the formation of atherosclerotic plaques. Since the

recruitment of monocytes to the vessel wall is pivotal to atherogenesis, targeting the

recruitment of monocytes to the subintima may produce clinical benefits and slow

the atherogenetic process.

1.3 SLIT2: A GUIDANCE CUE FOR CELL MIGRATION

During the development of the central nervous system (CNS), neurons must

migrate and project axons over long distances. Most axons emanating from the

CNS must cross the midline and then project longitudinally towards their synaptic

22

targets. The molecular mechanisms that guide this pathfinding include: contact

attraction, chemoattraction, contact repulsion and chemorepulsion. These

mechanisms act simultaneously in a coordinated manner to direct axonal

pathfinding (Tessier-Lavigne et al., 1996). Generally, guidance cues can either

promote or repress migration of neurons and axonal projections. For example,

netrins are diffusible chemotropic factors that attract commissural axons to the

midline (Kennedy et al., 1994). The Slit family of secreted proteins, together with

their cell-surface receptor Roundabout (Robo), act to repel neurons during CNS

development. Once commissural axons have crossed the midline, midline glial cells

express Slit to prevent the axons from re-crossing the midline. Drosophila Slit

mutants exhibit midline defects, such as collapse of the regular scaffold of

commissural and longitudinal axon tracts in the embryonic CNS (Rothberg et al.,

1988; Rothberg et al., 1990). A similar defect is observed in Robo mutants, where

projecting axon tracts cross the midline repeatedly (Kidd et al., 1998).

Recent studies demonstrate a role for Slit and Robo as guidance cues

outside of the CNS. For example, in Drosophila mesoderm migration, myocyte

precursors migrate away from the midline towards peripheral target sites where they

fuse to form muscle fibers. In Slit and Robo mutants, these cells do not migrate

away from the midline and instead fuse across the midline (Rothberg et al., 1990).

Interestingly, this defect can be reversed by expressing Slit protein in midline cells

(Kramer et al., 2001). Slit and Robo signaling also plays a role in nephrogenesis.

The proper localization of the kidney is dependent on the formation of a structure

called the ureteric bud. This process requires secretion of glial cell line-derived

23

neurotrophic factor (GDNF) by nearby mesenchymal cells. Slit and Robo knockout

mice display abnormal patterns of GDNF secretion and develop multiple ureteric

buds and multiple urinary collecting systems, implicating Slit and Robo in

nephrogenesis (Ray, L., 2004). Furthermore, variations in the human Robo gene

have been associated with familial vesicoureteral reflux (Bertoli-Avella et al., 2008),

a condition with improper insertion of ureters into the bladder resulting in retrograde

flow of urine from the bladder to the kidney. Therefore, Slit and Robo appear to play

a role in normal human urinary tract formation.

In addition to its role in embryogenesis, Slit also plays a role in the mature

organism. A recent study demonstrated Slit2-mediated inhibition of aortic smooth

muscle cell migration toward a gradient of platelet-derived growth factor (PDGF)

(Liu et al., 2006). This inhibition was shown to be mediated by suppressing the

activation of small GTPase Rac1. Furthermore, Slit2 can prevent breast cancer cell

metastasis. Both Robo and the chemokine receptor CXCR4 are expressed by some

human breast cancer cells, allowing them to migrate towards gradients of SDF-1α.

The lungs may produce high levels of SDF-1α, promoting breast cancer metastasis

to this tissue. Slit2 inhibited the chemotaxis, adhesion and chemoinvasion of breast

cancer cells (Prasad et al., 2004). Several other studies have demonstrated a role

for Slit2 as a tumor suppressor. Slit2 was shown to inhibit colony formation in lung,

colorectal and breast cancer cell lines (Dallol et al., 2002). Furthermore, Slit2 was

often epigenetically silenced in more aggressive forms of these and other cancers

(Dallol et al., 2003; Dallol et al., 2003; Dickinson et al., 2004). Therefore, these

studies imply a role for Slit and Robo in the adult organism and in cancer biology.

24

1.3.1 Expression

The expression of the Slit genes has been demonstrated in many organisms,

including Drosophila (Battye et al., 1999), Caenorhabditis elegans (Hao et al.,

2001), Xenopus (Chen et al., 2000), Gallus gallus domesticus (Holmes et al., 2001;

Vargesson et al., 2001), mice (Holmes et al., 1998; Piper et al., 2000), rats (Marillat

et al., 2002) and humans (Itoh et al., 1998). In mammals, there are three members

of the Slit family. Although Slit1 is predominantly expressed in the developing CNS

(Yuan et al., 1999), Slit2 and Slit3 are expressed outside the CNS, in the lung,

kidney and heart tissues (Wu et al., 2001). Importantly, Slit expression persists in

the adult organisms, suggesting a role for the Slit family beyond embryogenesis.

Expression of Robo has been demonstrated in Drosophila (Kidd et al., 1998),

mice (Yuan et al., 1999) and humans (Kidd et al., 1998). There are four isoforms of

Robo in mammals. Robo-1 has been shown to be most highly expressed in tissues

outside the CNS, including human leukocytes (Wu et al., 2001). Interestingly, the

tissue expression of Slit and Robo is relatively complementary, suggesting a

functional relationship in the adult organism (Yuan et al., 1999).

1.3.2 Slit and Robo Structure

The Slit family of proteins contains an N-terminal signal peptide, four leucine-

rich repeats (LRRs), nine epidermal growth factor (EGF) repeats and a C-terminal

cysteine knot (Fig 1.4) (Rothberg et al., 1988; Rothberg et al., 1990; Rothberg et al.,

1992). The EGF repeats and LRR allow the Slit proteins to interact with ECM

25

Figure 1.4 Primary Structure of Mammalian Slit2 and Robo-1 Proteins..

Mammalian Slit2 contains four leucine rich repeats (LRRs), nine epidermal growth

factor (EGF) repeats, a laminin G (G) domain, and a cysteine rich C terminus. The

Robo-1 receptor contains five immunoglobulin (Ig) repeats, three fibronectin (FN)

type III, a transmembrane Domain (TM) and four conserved cytoplasmic (CC)

signaling motifs.

26

components, such as glypican-1 (Ronca et al., 2001), enabling them to act as

localized, non-diffusible, signaling molecules. Furthermore, Slit2 can be cleaved

after the fifth EGF repeat by proteases to form N-terminal (Slit-N) and C-terminal

(Slit-C) fragments (Brose et al., 1999; Wang et al., 1999). The N-terminal fragment

includes the first 1118 amino acids and contains the four LRRs and the first five

EGF repeats, while the C-terminal fragment contains the remaining residues (Brose

et al., 1999). Importantly, it is the four LRRs that are necessary and sufficient for

interaction with the Robo receptor and downstream signaling (Battye et al., 1999).

Therefore, both the full length protein and the N-terminal Slit2 fragment can bind

Robo receptors to repel migrating cells and projecting axons (Nguyen Ba-Charvet et

al., 2001). Although the cleavage of Slit2 does not eliminate its activity, it may play a

role in its diffusion, since N-Slit appears to be more tightly associated with the cell

membrane. In rat neural tissue both the N-terminal and C-terminal fragments of Slit

were shown to bind heparan sulfate proteoglycan glypican-1 (Liang et al., 1999),

although the C-terminal fragment bound with higher affinity, suggesting a possible

regulatory mechanism for its diffusion.

Robo is a single-pass type-1 receptor and signaling molecule for the Slit

family of proteins. The extracellular region of Robo-1 contains five immunoglobulin

(Ig) repeats and three fibronectin type III domains. The cytoplasmic region of Robo-

1 contains four conserved cytoplasmic signaling motifs, CC0, CC1, CC2 and CC3

(Kidd et al., 1998; Zallen et al., 1998). Only the Ig domains of Robo are required to

bind to the LRRs in full length and N-terminal Slit2 (Battye et al., 2001; Chen et al.,

27

2001; Nguyen Ba-Charvet et al., 2001). The cytoplasmic CC motifs of Robo are

required for its response to Slit (Bashaw et al., 2000).

1.3.3 Slit2/Robo-1 intracellular signal transduction

Studies of neuronal tissue have demonstrated that Robo-1 can signal

through two pathways that lead to mobilization and remodeling of the cytoskeleton:

Enabled (Ena) protein and Rho GTPases. Both of these pathways require the CC

motifs in the cytoplasmic domain of Robo to signal.

Ena and its mammalian homologue (Mena) are members of a family of

proteins that are believed to link signal transduction to localized remodeling of the

actin cytoskeleton by binding to profilin, an actin binding protein which regulates

actin polymerization (Lanier et al, 1999; Wills et al., 1999). The bacteria Listeria

monocytogenes utilizes the Ena proteins for actin-based motility (Laurent et al,

1999). Ena was demonstrated to be a substrate for the Abelson kinase (Gertler et

al., 1989). Ena and Abelson can both bind to Robo. Ena binds to the CC1 motifs

while Abelson binds to the CC3 motif (Bashaw et al., 2000). Impairing Ena binding

reduced Robo function while mutations in Abelson results in Robo hyperactivity

(Bashaw et al., 2000).

A second pathway through which Slit/Robo mediates cell repulsion is through

modulation of Rho GTPase activity. A family of GTPase activating proteins, Slit

Robo GTPase activating proteins (srGAPs), were shown to bind Robo (Wong et al.,

2001). The SH3 domain of these proteins is required to bind to the CC3 motif of

Robo, while the GAP domain has activity for the Rho GTPases Rac, Cdc42 and

28

Rho (Wong et al., 2001). This suggests a model where Slit ligation of Robo induces

the recruitment of srGAP followed by inactivation of Rho GTPases and inhibition of

actin remodeling and cell motility. The literature supporting the role of Rho GTPases

in cell motility is consistent with this model.

1.3.4 Slit and Robo in leukocyte trafficking

Both neuronal and leukocyte cell migration require the recognition of

guidance cues, polarization of the cell and mobilization of the actin cytoskeleton.

Thus, a conservation of cell migration guidance mechanisms was proposed when

Slit2 was found to inhibit leukocyte migration, in addition to its well established role

in neuronal guidance (Guan et al., 2003; Kanellis et al., 2004; Prasad et al., 2007).

The first study to demonstrate Slit-mediated inhibition of leukocyte migration

was published in Nature by Wu et al., 2003. This study utilized transwell migration

assays to demonstrate Slit-mediated inhibition of chemotaxis of rat lymph node cells

to gradients of MCP-1 and neutrophil-like HL-60 cells to fMLP gradients (Wu et al.,

2003). Subsequently, Kanellis et al. demonstrated Slit2-mediated inhibition of

chemotaxis towards MCP-1 and fMLP in rat-derived macrophages (Kanellis et al.,

2004). Another study showed that Slit2 inhibited migration of dendritic cells (DCs)

(Guan et al., 2003). Although these early studies implicated Slit2 in modulation of

leukocyte chemotaxis, clear evidence for this role was lacking in primary human

cells. However, in 2007, Prasad et al. were able to demonstrate that Slit2 can inhibit

chemotaxis and transendothelial migration of primary CD4+ T lymphocytes and the

human Jurkat T cell line (Prasad et al., 2007).

29

These results are indicative of a role for Slit2 in human leukocyte

chemotaxis, as Slit2 has been shown to inhibit migration of human DCs and

lymphocytes. Furthermore, we have shown that Slit2 inhibits the in vitro chemotaxis

of primary human neutrophils, and the in vivo recruitment of mouse neutrophils

(Tole et al., 2009). Importantly, we have demonstrated that Slit2 inhibits neutrophil

chemotaxis towards a range of chemokines, both in vivo and in vitro, including

fMLP, IL-8, C5a and FKN (Tole et al., 2009). Therefore, Slit2 may have a

therapeutic role as a universal inhibitor of leukocyte migration.

1.4 RHO GTPases: Rac and Cdc42

Small GTPases of the Rho family are a part of the Ras superfamily of small

GTP-binding proteins. They are pivotal regulators of many signaling networks that

are activated by a diverse variety of receptor types. To date, over 20 mammalian

Rho GTPases have been characterized, and these can be grouped into 6 different

classes: Rac (Rac1, Rac2, Rac3, RhoG), Rho (RhoA, RhoB, RhoC), Cdc42

(Cdc42Hs, G25K, TC10), Rnd (Rnd3/RhoE, Rnd1/Rho6, Rnd2/Rho7), RhoD, and

TFF (Aspenström, P., 1999; Kjoller et al., 1999). When activated, Rho GTPases

regulate many important processes in all eukaryotic cells, including actin

cytoskeleton dynamics, transcriptional regulation, cell cycle progression, and

membrane trafficking. The activation, and hence the activity of Rho GTPases is

regulated by outside-in signals from a variety of receptor types, including GPCRs,

tyrosine kinase receptors, cytokine receptors and adhesion receptors. Rho

GTPases play an critical role in leukocytes as regulators of signaling networks that

30

allow these cells to perform specialized functions, such as chemotaxis, adhesion

and phagocytosis.

1.4.1 Structure and Regulation

All Rho GTPases contain two main structural domains, the C-terminal 'CAAX'

motif and a catalytic GTP domain. The 'CAAX' motif undergoes post-translational

processing, involving carboxy-terminal proteolysis of the AAX residues followed by

carboxyl-methylation. The modified C-terminal domain can then attach to

membrane lipids and facilitates membrane association and subcellular localization

of Rho GTPases (Gutierrez et al., 1989; Casey et al., 1989; Fujiyama et al., 1990).

The catalytic domain contains two regions, switch I and switch II. These domains

correspond to different structural conformations in the GTP-bound and GDP-bound

forms. Rho GTPases function as molecular switches by cycling between GDP-

bound and GTP-bound forms. When bound to GDP, Rho GTPases are inactive.

Upstream signaling events leading to the exchange of GDP for GTP switches the

protein to an active state. The active form of the protein can transduce signals via

interactions with downstream targets or effector molecules to produce a cellular

response. The intrinsic GTPase activity of Rho GTPases completes the cycle, by

hydrolyzing GTP, returning the GTPase to its inactive GDP-bound state.

There are three classes of molecules that interact with Rho GTPases and are

capable of regulating their activation state: guanine nucleotide exchange factors

(GEFs), GTPase-activating proteins (GAPs), and guanine nucleotide dissociation

inhibitors (GDIs). GEFs catalyze the exchange of GDP for GTP, leading to the

31

activation of Rho GTPases. GEFs stimulate the release of GDP allowing GTP,

which is present at higher concentrations in cells than GDP, to bind and activate

GTPases. To date, over 69 mammalian GEFs for Rho GTPases have been

identified (Rossman et al., 2005). They are characterized by the presence of a Dbl

homology domain (DH), which is capable of interacting with both the switch I and

switch II regions and catalyses the exchange of GDP for GTP. In addition, many of

these DH-domain containing proteins, such as Vav, contain a PH domain. The PH

domain allows GEFs to bind phosphoinositides, such as PIP3. This allows GEFs to

be localized to the plasma membrane where they can interact with other Rho

GTPase interacting proteins. Thus, GEFs promote the activation of Rho GTPases

and also facilitate their interaction with downstream effector molecules. On the other

hand, GAPs enhance the intrinsic GTPase activity of Rho GTPases, resulting in the

suppression of their activity. Although GTPases posses intrinsic GTPase activity,

the actual rate of GTP hydrolysis is relatively slow. Therefore, the interaction with a

GAP is required for efficient GTP hydrolysis, as this accelerates the cleavage step

by several orders of magnitude (Vetter et al., 2001). To date, more than 70

eukaryotic RhoGAPs have been discovered, 35 of these can be found in humans

(Tcherkezian et al., 2007). There exists a large diversity in the primary sequences of

the various GAPs. However, each one contains a Rho GAP domain with a

conserved tertiary structure composed of α helices and a catalytically critical

'arginine finger' which stabilizes the formation of the transition state during GTP

hydrolysis (Nassar et al., 1998). In addition, the Rho GAP domain can interact with

both the switch I and switch II regions on the GTPase domain (Gamblin et al.,

32

1998). This interaction allows GAPs to facilitate the intrinsic hydrolysis of GTP,

resulting in the inactivation of Rho GTPases. Finally, GDIs associate with Rho

GTPases in their inactive GDP-bound state and inhibit their activation by GEFs. In

addition, GDIs have also been shown to bind to GTP-bound GTPases, such as Rho

GTPase, and suppress GTPase activity (Oloffson, B., 1999). Finally, there is

evidence that GDIs can bind to isoprenyl moieties on the C-terminus of GTPases in

order to sequester them in the cytosol (Keep et al., 1997). The role of GDIs in

partitioning GTPases between the membrane and cytosol may be physiologically

more important than the inhibition of their activation. It is possible that the GDI-

mediated partitioning of GTPases may provide a storage pool of Rho GTPases that

may be readily utilized upon cell activation. Ultimately, the function of GDIs is to

prevent the activation of Rho GTPases, prevent their interaction with membranes,

and inhibit their downstream signaling networks.

1.4.2 Role of Rho GTPases in the regulation of the actin cytoskeleton

Rho GTPases are pivotal regulators of signaling networks that are activated

by chemokine and cytokine receptors, along with other receptor types, and result in

the mobilization of the cytoskeleton (Machesky et al., 1997). Actin polymerization is

a common response of motile cells to chemoattractants, and occurs following the

activation of Rho GTPases (Carson et al., 1986; Hall et al., 1989; Howard et al.,

1984). The movement of eukaryotic cells relies on coordinated extension of actin-

rich lamellipodia in the leading edge and retraction of the uropod at the rear of the

cell. The extension of lamellae in the leading edge involves rapid turnover of actin

33

filaments (Symons et al., 1991; Wang, Y., 1985). More stable actin-myosin cables

can be found in more established protrusions and in the middle and rear of the cell

(DeBiasio et al., 1988). Thus, cell motility requires the coordinated polymerization of

actin in protrusions at the leading edge and contraction of actin-myosin cables at

the middle and rear of the cell. In addition, other factors such as recycling of the

plasma membrane and integrin-mediated adhesion are important for cell motility

(Bretscher, M., 1996; Martenson et al., 1993; Yamada et al., 1995; Mitra et al.,

2005). Furthermore, coordinated actin assembly is important for integrin-mediated

adhesion and phagosome formation (Defacque et al., 2000; Calderwood et al.,

2000). All of these processes are dependent on coordinated mobilization of the

actin cytoskeleton, and are regulated by deployment of actin-binding proteins by

activated Rho GTPases. Rho GTPases are in an ideal position to control cell

motility and morphological changes in response to extracellular stimuli, such as

chemokine gradients. For example activation of Rho in fibroblasts results in the

assembly of stress fibers and focal adhesions (Ridley et al., 1992). The activation of

Rac causes extension of lamellipodia and assembly of small focal complexes

(Nobes et al., 1995; Ridley et al., 1992). Finally, activation of the Cdc42 Rho

GTPase leads to the formation of filopodial extensions (Nobes et al., 1995).

1.5 Rationale, Hypothesis & Objectives

1.5.1 Rationale

34

Monocyte recruitment and proliferation in the subintima is a hallmark of

atherosclerosis and vascular inflammation. The trafficking signals that recruit

monocytes to sites of inflammation are provided by chemoattractants. Although we

can target certain individual chemoattractants and their receptors, redundancy

exists in the chemokine signaling pathways that allow other pathways to

compensate for the loss of one or more. Therefore, it would be more efficient to

knock-out chemoattractant-mediated cell recruitment with a universal inhibitor of

chemokine GPCR signaling pathways.

The Slit family of proteins have long been known to act as inhibitors of cell

migration and axon projection in the CNS. More recent studies have implicated a

role for Slit2/Robo-1 signaling in diverse cell types, including leukocytes, both in

vitro and in vivo (Dallol et al., 2002; Guan et al., 2003; Kanellis et al. 2004; Liu et al.,

2006; Prasad et al., 2004; Prasad et al., 2007). Furthermore, we have demonstrated

Slit2-mediated inhibition of circulating human and mouse neutrophils to several

chemoattractant gradients (Tole et al., 2009). These data suggest that Slit2 may

inhibit cellular migration outside of the CNS, implying that the guidance mechanisms

controlling cell migration may be conserved across cell types. Indeed, Slit2 was also

shown to inhibit the chemotaxis of circulating human neutrophils induced by several

classes of chemoattractants, including: fMLP, IL-8 and C5a (Tole et al., 2009). In

addition, Slit2 was shown to dramatically decrease neutrophil recruitment in an in

vivo model of murine peritonitis induced by sodium periodate or other

chemoattractants (C5a, mouse inflammatory protein 2). However, limited data is

available on the effect of Slit2 on monocyte migration and function. Studies in

35

neuronal cells have implicated the Rho GTPases in the Slit2-mediated inhibition of

migration. In human neutrophils, Slit2 was shown to inhibit the activation of Rho

GTPases, Cdc42 and Rac2, after fMLP stimulation (Tole et al., 2009). Since Rho

GTPases are important regulators of the cytoskeleton, the effects of Slit2 may go

beyond affecting the migration of a cell to include modulation of other functions such

as adhesion or phagocytosis, as these functions all involve actin cytoskeleton

remodelling.

1.5.2 Hypothesis

We hypothesize that Slit2/Robo-1 signaling can inhibit

monocyte/macrophage chemotaxis and modulate immune functions such as

adhesion to endothelial cells and phagocytosis of Ig-opsonized beads. We

hypothesize that Slit2 exerts its effects by suppressing the activity of Rho GTPases

Rac and Cdc42. However, we hypothesize that Slit2/Robo-1 signaling will have no

effect on the activation of MAPKs, as was observed in primary human neutrophils

(Tole et al., 2009). In addition, we hypothesize that Slit2, administered to mice

intraperitoneally or intravenously will inhibit monocyte/macrophage recruitment to

the peritoneal cavity, in vivo, in a murine model of sodium periodate-induced

inflammation.

1.5.3 Objectives

The first objective of this study is to determine if monocytes/macrophages

express the Slit2 receptor Robo-1. Monocytes/macrophages must express the

36

receptor in order to be responsive to the effects of Slit2. Next, the effect of Slit2 on

monocyte/macrophage chemotaxis will be characterized in vitro using transwell

chemotaxis assays and treatment with the monocyte/macrophage chemoattractant

SDF-1α. In addition, the intracellular signaling cascades that mediate Slit2/Robo-1

signaling in monocytes/macrophages will be investigated by observing the role of

Slit2 on Rho GTPases Cdc42 and Rac1, and on the Akt, Erk, and p38 MAPKs. Pull-

down assays for activated, or GTP-bound, Rho GTPases will be performed,

following incubation with Slit2 and activation with SDF-1α. To determine the effect

of Slit2 on MAPKs, western blots for phosphorylated and total MAPKs will be

performed, following incubation of monocytes/macrophages with Slit2 and treatment

with SDF-1α. The effect of Slit2 on monocyte/macrophage adhesion to activated

endothelial cells will also be investigated. Confluent endothelial cell monolayers will

be activated with a proinflammatory cytokine (TNF-α), and the adhesion of

monocytes/macrophages following incubation with Slit2 will be tested. Furthermore,

the effect of Slit2 on monocyte/macrophage recruitment in vivo using a murine

model of sodium periodate induced peritonitis will be conducted. Slit2 will be

administered intraperitoneally or intravenously an hour prior to the induction of

peritonitis, and peritoneal lavages will be performed to determine the number of

recruited monocytes/macrophages. In addition, the dose of Slit2 required to

optimally inhibit monocyte/macrophage recruitment in vivo will be determined by

performing a dose-response experiment using the murine peritonitis model and

intravenously administered Slit2. Furthermore, a time-course experiment will be

performed by administering Slit2 at 1 day, 4 days, and 10 days prior to inducing

37

peritonitis, to determine the biological half-life of intravenously administered Slit2.

Finally, the effect of Slit2 on other monocyte/macrophage functions involving Rho

GTPases, such as phagocytosis, will be investigated. To do this, phagocytosis

assays will be conducted with Ig-opsonized latex beads, and the

monocyte/macrophage phagocytosis following incubation with Slit2 will be

quantified.

38

CHAPTER 2

MATERIALS & METHODS

2.1 Reagents and antibodies

Unless otherwise stated, reagents were purchased from Sigma-Aldrich.

Monocyte isolation kit was purchased from StemCell Technologies. The following

primary antibodies were used: anti human Robo-1 (ab7279, Abcam, Cambridge,

MA), anti-myc 9E10 (Covance, QC, Canada), anti-human Cdc42 (Cell Signaling,

Danvers, MA), anti-human Rac1 (Upstate Biotechnology, Lake Placid, NY). The

following secondary antibodies were used: Cy-3 conjugated anti rabbit IgG, Cy-2

conjugated anti-human IgG, and horseradish peroxidase-conjugated anti rabbit IgG

(Jackson Immunoresearch Laboratories, Bar Harbor, ME). MAPK Antibodies were

purchased from Invitrogen Canada (Burlington, Ontario, Canada).

2.2 Isolation of human monocytes

Blood from healthy volunteers was obtained on each day of experimentation.

The monocytes were isolated using a Polymorphprep gradient separation solution

(Axis-Shield, Norway) and an EasySep® Negative Selection kit (StemCell

Technologies). A volume of blood was gently layered over an equal volume of

Polymorphprep solution, and centrifuged at 460 g for 35 minutes at an acceleration

of 2 units and deceleration of 0 units in order to prevent cell activation . The lower

layer containing peripheral blood mononuclear cells (PBMCs) was collected,

washed in cold PBS with 2% fetal bovine serum (FBS) and 1mM EDTA and

centrifuged at 260 g, room temperature for 5 min. When redness could still be seen

39

in the pellet, indicating the presence of red blood cells, an additional wash in cold

PBS with 2% FBS and 1 mM EDTA was performed. The EasySep® Negative

Selection procedure was performed according to the manufacturer‟s protocols.

Briefly, PBMCs (5x107 cells/mL) are labelled with EasySep® Human Monocyte

Enrichment Cocktail (50µg/mL cells)(StemCell Technologies) for 10 minutes at 4

°C. The Negative Selection Enrichment Cocktail contains a combination of

monoclonal antibodies that were purified from hybridoma culture supernatant by

affinity chromatography using Protein A or Protein G Sepharose. These antibodies

are bound in bispecific Tetrameric Antibody Complexes which are directed against

cell surface antigens on human leukocytes (CD2, CD3, CD16, CD19, CD20, CD56,

CD66b, CD123, glycophorin A) and dextran. These mouse monoclonal antibodies

are of the IgG1subclass. In addition, this cocktail also contains an FcR blocker to

prevent non-specific binding of monocytes. The antibody subclass of the FcR

blocker is IgG2b. The cells were then labelled with EasySep® Magnetic

Microparticles (50 µg/mL cells)(StemCell Technologies) for 5 minutes at 4 °C. The

Magnetic Microparticles contain a suspension of magnetic dextran iron particles in

TRIS buffer. Then, the EasySep® magnet was used to remove the magnetically

labelled cells, while the pure monocytes are poured off. The purified monocytes

were then resuspended in ice cold PBS with 2% FBS and 1 mM EDTA for

subsequent experiments. Experiments were performed within 1-2 hours of cell

isolation. Cell viability was determined to be >98% by Trypan blue staining

2.3 Cell culture

40

Human acute monocytic leukemia (THP-1) cells were cultured in RPMI-1640

(Sigma Chemical, St Louis, MO) supplemented with 5% FBS. Primary human

umbilical vein endothelial cells (HUVECs) and human arterial endothelial cells

(HAECs) were grown in endothelial basal medium 2 (EBM-2) supplemented with

Clonetics® EGM-2 SingleQuots® (Lonza, Walkersville, MD) These include: 10mL

FBS, 2mL of recombinant human fibroblast growth factor-B, 0.5mL of ascorbic acid,

0.5mL of recombinant human vascular endothelial growth factor, 0.5mL of

recombinant human epidermal growth factor, 0.5mL heparin, 0.2mL hydrocortisone,

0.5mL recombinant insulin-like growth factor-1, and 0.5mL gentamicin sulfate

amphotericin-B for 500mL of EBM-2. Only low passage cells (up to passage 11)

were used for adhesion experiments. Once cellular confluency was reached, cells

were passaged and/or seeded into 96-well clear bottom tissue culture plates for

adhesion experiments.

2.4 Slit2 expression and purification

Stable human embryonic kidney 293 cell line expressing full-length or N-

terminal human Slit2 with a His tag at its carboxyl terminus was used for Slit2

purification. Recombinant Slit2 was purified by Sylvie Perret and Dr. Yves Durocher

at the National Research Council of Canada. The presence of purified Slit2 was

confirmed by immunoblotting with poly anti-His antibody (Sigma A-7058). Following

purification, Slit2 was aliquotted, snap frozen and stored at -80 °C for future use.

Aliquots were never re-frozen or used after storage at 4 °C. In the experiments

described below, Slit2 was generally used at a concentration of 4.6 µg/ml diluted in

41

ice cold PBS. Endotoxin concentrations in our Slit2 preparation ranged from 0.2-0.8

ng/ml, yielding final experimental concentrations of 12-40 pg/ml which are well

below those thought to activate leukocytes (Moore et al., 2000). To verify this, we

added similar concentrations of endotoxin in neutrophil Transwell assays, and found

that such levels of endotoxin had no effect on neutrophil migration (Tole et al.,

2009).

2.5 Immunofluorescence

Primary human monocytes, murine RAW 264.7 macrophages and THP-1

cells were allowed to settle onto fibronectin-coated coverslips and allowed to adhere

for 3 minutes at room temperature (RT). The cells were then fixed with 4%

paraformaldehyde (PFA) for 10 minutes at RT. The cells were stained with rabbit

anti-Robo-1 (1µg/ml) for 2 hours at RT, washed with phosphate buffered saline

(PBS) and then incubated with Cy3-conjugated anti-rabbit secondary antibody for 1

hour at RT. A Leica DMIRE2 spinning disc confocal microscope (Leica

Microsystems, Toronto, Ontario, Canada) equipped with a Hamamatsu back-

thinned EM-CCD camera and Volocity software (Improvision Inc., Lexington, MA)

was used to capture images.

2.6 Transwell migration assay

Human THP-1 cells (1x106 cells/mL, 100 µL/condition) were incubated with

PBS vehicle or Slit2 at concentrations ranging from 46 ng/mL to 4.6 µg/ml for 10

minutes at 37°C and 5% CO2. The cells were then loaded into the top chamber of a

42

5 µm Transwell insert (Corning Life Sciences, Corning, NY) designed for a 24-well

plate. A glass coverslip was placed in the bottom well. The bottom chamber was

filled with 600 µL of serum-free RPMI-1640 alone or with SDF-1α (100ng/mL) in the

presence or absence of Slit2. Monocytes were allowed to migrate into the bottom

chamber for 3.5 hours at 37°C and 5% CO2. Following incubation, the monocytes

were rapidly spun down onto the coverslips (by centrifugation of the entire plate at

100 g, 1 min), fixed with 4% PFA, washed with PBS and labeled with DAPI dye for

visualization of cell nuclei. A Leica DMIRE microscope was used to take

representative high-power (40X) images and total number of cells was counted in at

least 10 random fields. The data represent the mean values ± SEM from at least 4

independent experiments.

2.7 Immunoblotting

THP-1 cells were serum starved overnight, resuspended in serum-free

RPMI-1640 (1x106 cells/mL) and incubated with either PBS vehicle or Slit2 (4.6

µg/mL) for 10 min at 37°C, 5% CO2. The cells were subsequently activated with

SDF-1α (100ng/mL) for 0, 2 and 5 minutes. The cells were then washed with 1 mL

of ice cold PBS. Next, the cells were lysed using ice-cold lysis buffer (50 mM Tris,

pH 7.5, 10% glycerol, 100 mM NaCl, 1% NP-40, 5 mM MgCl2, 1 mM DTT, 1 mM

PMSF, 1/100 protease inhibitor cocktail and 1 mM NaVO3). Protein samples were

added to 6x SDS gel loading buffer (1% ß-mercaptoethanol, 1% SDS, 30% glycerol,

0.0012% bromophenol blue, Tris HCl 0.28 M, pH 6.8). Samples were centrifuged

briefly at 10,000 rpm for 1 min. Protein gels were electrotransferred to poly-

43

vinyldene fluoride (PVDF) membranes (Millipore) in transfer buffer (25 mM Tris

base, 190 mM Glycine, 0.05% SDS, and 20% methanol) for 1.5 hr at 350 A at 4°C.

Membranes were probed for i) phosphorylated and total Akt, ii) phophorylated and

total Erk, and iii) phophorylated and total p38 MAP kinase. The membranes were

always probed with antibody detecting the phophorylated protein first, stripped, and

then reprobed with the antibody detecting total protein as a loading control.

Immunoreactive bands were visualized by enhanced chemiluminescence

(Amersham Biosciences, UK Ltd, Buckinghamshire, UK) and the signal captured

onto Kodak-Biomax film (Rochester, NY, USA). Image J software (NIH, Bethesda,

MA, USA) was used for densitometry analysis, and subsequent statistical analysis

was performed using Microsoft Excel.

2.8 Cdc42 and Rac2 activation assays

The pull-down assay for the Rho GTPases Cdc42 and Rac1 was performed

as previously described (Benard et al., 1999; Tole et al., 2009) with slight

modifications. The phosphate binding domain (PBD; 67-150 aa) of PAK1 in pGEX-

4T3 vector was expressed as a GST fusion protein in BL21 (DE3) E. coli cells. The

GST-PBD fusion protein was affinity purified using glutathione sepharose 4B beads

(GE Healthcare). THP-1 cells (1x107 cells/sample) diluted in 500 µL 37°C warmed

HEPES-HBSS were incubated with PBS vehicle or with 4.6 µg/ml Slit2 at 37°C and

5% CO2 for 10 minutes. Cells were then stimulated with SDF-1α (100ng/mL) for 0,

2, or 5 minutes at 37°C and the reactions were stopped by adding 500 µL ice-cold

lysis buffer. Samples were centrifuged at maximal speed in a bench-top centrifuge

44

for 5 min at 4°C and an aliquot of supernatant was used as a loading control. The

remaining supernatants were added to GST-PBD glutathione beads (20 mg

beads/sample). Samples were rotated at 4°C for 1 hour, washed 3 times with cold

wash buffer (50 mM Tris, pH 7.5, 40 mM NaCl, 0.5% NP-40, 30 mM MgCl2, 1 mM

DTT, 1 mM PMSF, 0.1 mM NaVO3 ) and added with 20 µLof 2 x Laemmli loading

buffer. The samples were then run on SDS-PAGE and transferred onto a 0.2 mm

PVDF membrane (Millipore). Cdc42 and Rac2 were detected using rabbit anti-

human Cdc42 (Cell Signaling, Danvers, MA) and rabbit anti-human Rac2 (Upstate

Biotechnology, Lake Placid, NY) primary antibodies and goat anti-rabbit HRP-

conjugated secondary antibodies. Densitometry analysis was performed on the

blots using Image J software (NIH, Bethesda, MA, USA). The data represent the

mean values ± SEM from 3 independent experiments.

2.9 Adhesion

Primary human endothelial cells (HUVECs and HAECs) were seeded

(~1x104 cells/well) in 96-well tissue culture plates and grown to confluence. Once

confluence was confirmed using a light microscope, the wells were aspirated and

replenished with endothelial basal medium 2 alone or with TNF-α (20 ng/mL). The

plates are then incubated at 37°C and 5% CO2 for 4 hours. THP-1 cells were

simultaneously labeled with Calcein AM at 37°C and 5% CO2 for 30 mins. After

labeling, monocytic THP-1 cells were washed once in 45 mL of PBS, pelleted (1500

rpm, 5 min) and resuspended in serum-free RPMI-1640 at 1x106 cells/mL. THP-1

cells were then incubated with PBS in the presence or absence of Slit2 (4.6 ug/mL)

45

at 37°C and 5% CO2 for 10 mins. The THP-1 cells were allowed to settle onto the

endothelial cell monolayers (1x105 cells/well) and incubated at 37°C and 5% CO2

for 30 mins. The plates were then centrifuged (100g, 1 min) upside down to remove

non-adherent cells. A fluorescent plate reader was used to measure the

fluorescence intensity of each well (494-517 nm for Calcein AM). Fluorescence

intensities are normalized to the unstimulated condition. The data represent the

mean values ± SEM from at least 4 independent experiments.

2.10 Murine peritonitis

Experimental murine peritonitis was carried out as previously described with

slight modification (Lotero et al., 2001; Jiang et al., 2005; Viriyakosol et al., 2005).

All procedures were performed in accordance with the Guide for the Humane Use

and Care of Laboratory Animals and were approved by The Hospital for Sick

Children Research Institute Animal Care Committee. For the experiments in Fig.

3.7, Slit2 (1.8 µg/mouse) was administered intraperitoneally to BALB/c mice

(Chares River Canada) an hour prior to sodium periodate induced peritonitis. For

the experiments in Fig. 3.8 and 3.9, CD1 mice were utilized. For the dose-titration

experiments presented in Fig. 3.8, Slit2 (4.6 µg, 460 ng or 46 ng) was administered

intravenously via tail-vein injection. For the time-course experiments presented in

Fig 3.9, we administered Slit2 intravenously (1.8 µg/mouse) at 1 day, 4 days, and

10 days prior to inducing peritonitis with sodium periodate (1mg/mouse) injected

intraperitoneally. For the experiments in Fig 3.7 and Fig. 3.8, peritonitis was induced

an hour after Slit2 or PBS treatment, with an intraperitoneal injection of sodium

46

periodate (1mg/mouse). Peritoneal lavages were performed after 24 hours with 5

mL of cold PBS containing 2% FBS. The cells were washed, red blood cells lysed

and hemocytometer counts performed. The data represent the mean values ± SEM

from at least 4 independent experiments.

2.11 Phagocytosis

Monocyte phagocytosis was performed as previously described (Yan et al.,

2007) with slight modifications. Human IgG (1 mg/ml) was coated onto 3.8 µm latex

beads for 2 hours at room temperature. RAW 264.7 macrophages were incubated

with Slit2 (600 ng/ml) or control medium (equal volume) for 10 minutes, exposed to

the latex beads, centrifuged (1000 rpm for 30s) to initiate phagocytosis, and plated

onto fibronectin-coated (20 µg/ml) coverslips. Phagocytosis was terminated after 30

min and external beads were labeled on ice using anti human Cy-2 conjugated

secondary antibody. Slit2 or control medium were present throughout the course of

phagocytosis. Images of at least 10 random fields were acquired using a Leica

deconvolution microscope. To determine the number of ingested particles, total

beads were counted using DIC and the number of external, fluorescently-labeled

beads were subtracted. The phagocytic index (number of ingested beads/ number

of cells) was used as an outcome measure.

2.12 Statistical analysis

Analysis of variance (ANOVA) followed by Bonferonni post-hoc tests were

performed using SPSS statistical software to analyze the data from adhesion

47

experiments. In all other cases, the Student‟s t-test was used. Significant difference

was considered for p<0.05. Graphic representation show mean ± SEM as variance

bars.

48

CHAPTER 3

RESULTS

3.1 Monocytes express the Slit2 receptor, Robo-1.

Immunoblotting confirmed Robo-1 protein expression in primary human

monocytes, mouse RAW 264.7 macrophages and human THP-1 monocytic cells.

Furthermore, immunofluorescence staining confirmed the presence of Robo-1 on

the surface of primary human monocytes and human monocytic THP-1 cells, co-

localizing with a membrane marker (Fig. 3.1). Collectively, these data demonstrate

that primary human monocytes, mouse RAW 264.7 macrophages and human

monocytic THP-1 cells express the Slit2 receptor, Robo-1.

3.2 Slit2 inhibits chemotaxis of human monocytic THP-1 cells.

The neuronal literature has clearly demonstrated the role of Slit2 as a

repellent of neuronal cells and projecting axons. More recent studies have shown

that Slit2 may act as a general chemorepellent, since it was shown to inhibit the

chemotaxis of diverse cell types, including: smooth muscle cells (Liu, et al., 2006),

DCs (Guan, et al. 2003), T lymphocytes (Prasad, et al., 2007), RAW 264.7

macrophages (Kanellis, et al., 2004) and primary human neutrophils (Tole et al.,

2009). Since human monocytes also expressed Robo-1, we hypothesized that Slit2

also inhibits monocyte chemotaxis.

Transwell migration assays were performed to determine the effect of Slit2

on monocyte chemotaxis. Human monocytic THP-1 cells were utilized for these

experiments. THP-1 cells failed to migrate in the absence of the chemokine SDF-1α

49

(Fig. 3.2A&C). When SDF-1α (100 ng/mL) was added to the bottom chamber,

monocytic THP-1 cell migration to the lower chamber was significantly increased

(Fig. 3.2B), p<0.01. To test the effect of Slit2 on monocytic THP-1 cell chemotaxis,

we incubated the cells with Slit2 (4.6 µg/ml) for 10 minutes and tested their ability to

migrate when Slit2 (4.6 µg/ml) alone (Fig. 3.2C) or Slit2 with SDF-1α (100 ng/mL)

(Fig. 3.2D) was added to the bottom chamber. In the absence of a chemokine

gradient, monocytic THP-1 cells pre-treated with Slit2 failed to migrate to the bottom

chamber (Fig. 3.2C). However, Slit2 treated cells exhibited decreased chemotaxis in

the presence of a chemokine gradient (Fig. 3.2D), p<0.01. In addition, we tested the

effect of N-Slit2, a cleaved N-terminal fragment containing all four LRR required for

signaling, on the chemotaxis of monocytic THP-1 cells. Monocytic THP-1 cells

treated with N-Slit2 also exhibited decreased chemotaxis in the presence of a

chemokine gradient (Fig. 3.2D), p<0.01. These data demonstrate that both the full

length Slit2 and N-Slit2 can inhibit SDF-1α-mediated chemotaxis of monocytic THP-

1 cells, but no effect on monocytic THP-1 cell chemotaxis is observed in the

absence of a chemokine gradient.

3.3. Slit2 treatment inhibits activation of Rac2 and Cdc42

Slit2 has been shown to inhibit migration of neuronal cells via recruitment to

the intracellular domain of Robo of a novel family of Slit Robo Rho GTPase

activating proteins (srGAPs). srGAPs convert the active GTP-bound forms of Rho

GTPases, Cdc42 and Rac1, to their inactive GDP-bound forms. (Wong et al., 2001).

In a study of vascular smooth muscle cell migration, Slit2 was shown to suppress

50

the activation of Rac1 (Liu et al., 2006). Rac1 and Cdc42 have been demonstrated

to play critical roles in leukocyte polarization and chemotaxis. HL-60 cells

transfected with dominant-negative construct of Cdc42 show impaired migration

(Srinivasan et al., 2003). Therefore, we hypothesized that the observed decrease in

monocyte chemotaxis may be due to Slit2-mediated inactivation of Rac1 and/or

Cdc42. We utilized the p21-binding domain (PBD) of PAK1, which only binds to

active GTP-bound forms of Rac1 and Cdc42 (Benard et al., 1999), conjugated to

GST beads (GST-PBD) in order to pull down activated forms of Rac1 and Cdc42.

Human monocytic THP-1 cells were incubated with PBS in the presence or absence

of Slit2 (4.6 µg/mL) for 10 minutes and then stimulated with SDF-1α (100 ng/mL) for

2 minutes. Activated Rho GTPases were pulled down using GST-PBD beads.

Subsequently, immunoblotting was performed for Cdc42 and Rac1. Unstimulated

monocytic THP-1 cells had low basal levels of activated Rac1 and Cdc42.

Stimulation with SDF-1α led to a 6.4 fold increase (†, p<0.05) in GTP-bound Cdc42

and a 21.6 fold (†, p<0.05) increase in GTP-bound Rac1, compared with

unstimulated cells (Fig. 3.3). Slit2 treatment alone had no effect on baseline

activation of Rac1 and Cdc42 (Fig. 3.3). However, Slit2 treatment significantly

reduced SDF-1α mediated activation of Cdc42 and Rac1 (Fig. 3.3). Monocytic THP-

1 cells incubated with Slit2 had a 3.8 fold (†, p<0.05) increase in GTP-bound Cdc42

and a 12.2 fold (†, p<0.05) increase in GTP-bound Rac1, compared to unstimulated

cells. These data suggest that a disruption in SDF-1α-mediated Rho GTPase

activation is involved in Slit2-mediated inhibition of monocytic THP-1 cell

51

chemotaxis towards SDF-1α. The data represent the mean values ± SEM from 4

independent experiments.

3.4. Akt and Erk, but not p38 MAPK pathways are affected by Slit2 treatment.

The signal transduction pathways leading from chemoattractant receptor

activation to chemotaxis are not fully understood. Generally, chemoattractant

receptor stimulation activates several MAPK pathways including: PI3K/Akt, Erk and

p38. In neutrophils, PI3K-dependent production of PIP3 and subsequent recruitment

and activation of Akt MAPK at the leading edge is important for migration (Heit et

al., 2002). Another study in human monocytes demonstrated that an inhibitor of

MEK inhibited MAPK activation and MCP-1-mediated chemotaxis (Yen et al., 1997).

In fact, MCP-1-mediated chemotaxis of monocytic THP-1 cells was shown to be Erk

MAPK dependent (Kintscher et al., 2000). Thus, MAPK inhibitors can arrest

chemotaxis. Therefore, we tested the effect of Slit2 on chemoattractant-induced

activation of Akt, Erk and p38 MAPK pathways in human monocytic THP-1 cells.

Stimulation with SDF-1α induced robust activation of the Akt and Erk MAPK

pathways (Fig. 3.4A). However, no activation of the p38 MAPK was observed (Fig

3.4A & D). Stimulation with SDF-1α for 5 minutes led to a 2.9 fold increase (†,

p<0.05) in phosphorylated Akt and a 3.2 fold (†, p<0.05) increase in phosphorylated

Erk, compared with unstimulated cells (Fig. 3.4A-C). Incubation with Slit2 alone had

no effect on baseline activation of Akt, Erk and p38 MAPKs (Fig. 3.3). However,

incubation with Slit2 significantly reduced SDF-1α-mediated activation of Akt and

Erk MAPKs at 5 minutes, although no effect was observed for p38 MARK (Fig. 3.3).

52

Monocytic THP-1 cells incubated with Slit2 had a 1.7 fold (†, p<0.05) increase in

phosphorylated Akt and a 1.9 fold (†, p<0.05) increase in phosphorylated Erk,

compared to unstimulated cells. Therefore, incubation with Slit2 decreased the

activation of Akt and Erk MAPKs (Fig. 3.4A-C) at 5 minutes after SDF-1α

stimulation. However, incubation with Slit2 had no effect on the p38 MAPK pathway

(Fig. 3.4A & D). These results suggest that Slit2 treated monocytes might have a

defect in the synthesis of PIP3 and subsequent recruitment and activation of Akt

MAPK.

3.5. Slit2 inhibits adhesion of monocytic THP-1 cells to activated human

umbilical vein endothelial cell and human arterial endothelial cell monolayers.

After their initial recruitment, monocytes must firmly arrest on the

endothelium and undergo diapedesis to reach inflammatory foci. Integrins on the

surface of leukocytes bind to Ig superfamily members such as ICAM-1 and VCAM-1

on the surface of endothelial cells (Ley et al., 2007). Rho GTPases participate in

many cellular processes that transmit signals from the cell surface to influence the

activity of the actin cytoskeleton (Sechi et al., 2000). Leukocyte adhesion to

endothelial cells requires outside-in signaling which can be initiated by integrin

ligation and clustering, which is partially dependent on Rho GTPases (Ley et al.,

2007). Since Slit2 inhibits the activation of Cdc42 and Rac2 Rho GTPases, we

tested the effect of Slit2 on adhesion of monocytic THP-1 cells to endothelial cells.

Endothelial cells were activated with TNF-α for 4 hours in order to simulate

inflammation and increase endothelial expression of adhesion molecules. Monocytic

53

THP-1 cells were allowed to adhere for 30 minutes and the plates centrifuged

upside down at 100xg for 1 min to remove non-adherent cells. For HUVECs (Fig.

3.5), Slit2 treatment alone had no effect on cell adhesion, whereas TNF-α

stimulation of endothelial monolayers increased baseline adhesion to almost 400%,

p<0.005 (Fig. 3.5). When monocytic THP-1 cells were incubated with Slit2,

adhesion was abolished to near baseline levels, P<0.05 (Fig. 3.5). Since adhesion

characteristics differ for different types of endothelial cells, we wanted to use cell

types similar to those affected in human cardiovascular disease. Therefore, we

utilized primary HAECs to confirm our findings from HUVECs. The same trend was

observed for HAECs (Fig. 3.6), Slit-treatment alone had no effect of cell adhesion,

while TNF-α stimulation of endothelial monolayers increased baseline adhesion by

over 200%, p<0.01 (Fig. 3.6). When monocytic THP-1 cells were incubated with

Slit2, adhesion was abolished to near baseline levels, P<0.005 (Fig. 3.6). These

data suggest that Slit2 may play a role in monocyte adhesion to vascular and

arterial endothelium under inflammatory conditions.

3.6 Slit2 inhibits monocyte recruitment in vivo.

We showed that Slit2 inhibits monocyte chemotaxis and adhesion to

activated endothelial cells. Thus, we wanted to investigate the functional relevance

of these observations. To study the effects of Slit2 on monocyte recruitment in vivo,

we used a previously described mouse model of chemical irritant peritonitis (Jiang

et al., 2005; Viriyakosol et al., 2005). Sodium periodate injection alone induced

peritonitis, with robust monocyte recruitment compared to control mice (Fig.3.7,

54

p<0.001). Intraperitoneal administration of Slit2 diminished monocyte recruitment

nearly four-fold (Fig. 3.7; p<0.001). Next, we wanted to test whether the effect of

Slit2 is dose-dependent. Again, injection of sodium periodate alone induced

vigorous peritonitis when compared to control (Fig. 3.8, p<0.05). Slit2 significantly

inhibited monocyte recruitment at doses of 4.6 and 0.46 µg (Fig. 3.8, p<0.05).

Although administration of 46 ng of Slit2 decreased monocyte recruitment by half,

this effect was not statistically significant (Fig. 3.8). Finally, we performed a time-

course experiment to determine the duration of Slit2 biological activity following

intravenous administration. Again, female CD1 mice were utilized for these

experiments. Slit2 (1.8 µg/mouse) was administered intravenously at 10 days, 4

days and 1 day prior to induction of experimental peritonitis. Sodium periodate

alone induced vigorous peritonitis when compared to control (Fig 3.9, p<0.001).

Intravenous administration of Slit2 one day before peritonitis completely abolished

cell recruitment to baseline (Fig 3.9, p<0.001). When Slit2 was administered

intravenously 4 days prior to the induction of peritonitis, monocyte recruitment was

again significantly inhibited (Fig 3.9, p<0.01). However, pre-treatment with Slit2 at

10 days prior to induction of peritonitis had no effect on monocyte recruitment.

Thus, these data indicate that Slit2 has very potent effects on in vivo monocyte

recruitment, with persistent biological activity even when administered 4 days prior

to an inflammatory insult. This suggests that local or systemic administration of Slit2

may be used to alleviate monocyte recruitment in inflammation and atherosclerosis.

Because Slit2 is heavily glycosylated, it is relatively 'sticky' and may achieve high

55

local concentrations by interacting with ECM components such as glypican-1

(Ronca et al., 2001).

3.7 Slit2 does not alter monocyte phagocytosis.

Monocytes/macrophages are professional antigen presenting cells and can

therefore internalize and destroy pathogens and cellular debris. They can also

internalize opsonized or non-opsonized targets. This is mediated by Fc receptors for

Igs and the integrin Mac-1 for complement components (Aderem et al., 1999). Rho

GTPases were shown to be required for calcium signaling and phagocytosis by Fcγ

receptors in macrophages (Hackam et al., 1997; Caron et al., 1998). Since Slit2

inhibits the activation of Rho GTPases, we tested the effect of Slit2 on monocyte

phagocytosis. RAW macrophages were centrifuged together with IgG-opsonized

latex beads to initiate phagocytosis for 10 minutes. External beads were

subsequently fluorescently labeled and images of at least 10 random fields were

captured. Slit2 treatment had no effect on the phagocytic index (number of ingested

particles/ number of cells) of RAW 264.7 macrophages (Fig 3.10A&B; Min Rui-Crow

performed these experiments).

56

57

Figure 3.1 Slit2 is expressed by monocytes.

Robo-1 expression has been confirmed in primary human monocytes and

human monocytic THP-1 cells. A, western blotting for Robo1 protein in murine

RAW 264.7 macrophages, human monocytic THP-1 cells and primary human

monocytes. B, Surface immunofluorescence staining showing the

co-localization of cell surface Robo-1 and a membrane marker. These results

confirm the presence of the Robo-1 receptor on the surface of monocytic

THP-1 cells, RAW 264.7 macrophages and primary human monocytes.

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59

Figure 3.2 Slit2 inhibits monocyte chemotaxis.

Monocyte chemotaxis was studied in vitro using a Transwell membrane inserts. A,

Human monocytic THP-1 cells failed to migrate in the absence of SDF-1α (Fig.

3.2A). When SDF-1α (100 ng/mL) was added to the bottom chamber, THP-1 cells

exhibited increased migration to the lower chamber (Fig. 3.2A) (†, p<0.001). To test

the effect of Slit2 on THP-1 chemotaxis, we incubated the cells with Slit2 (4.6 µg/ml)

for 10 minutes and tested their ability to migrate when Slit2 (4.6 µg/ml) alone (Fig.

3.2A) or Slit2 with SDF-1α (Fig. 3.2A) was added to the bottom chamber. In the

absence of a chemokine gradient, THP-1 cells incubated with Slit2 failed to migrate

to the bottom chamber (Fig. 3.2A). Slit2 treatment decreased chemotaxis towards

an SDF-1α gradient (Fig. 3.2A) (†, p<0.001). B, THP-1 cells were incubated with

full-length Slit2, N-terminal Slit2 or PBS vehicle for 10 minutes prior to migration.

Slit2 and the chemokine SDF-1α were added to the bottom well only. After 3.5

hours, the number of migrated cells was quantified microscopically. Both the full

length protein and the N-terminal fragment of Slit2 were able to inhibit THP-1 cell

chemotaxis towards an SDF-1α gradient (†, p<0.001).

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61

Figure 3.3 Slit2 inhibits activation of Rho GTPases (Cdc42 and Rac1).

Human monocytic THP-1 cells were incubated with PBS alone or containing

Slit2 (4.6 μg/mL) for 10 minutes and stimulated with SDF-1α (100ng/mL) for

2 minutes and lysates collected. GTP-bound or activated Cdc42 and Rac1 were

pulled down using GST-PBD beads. A, Western blots showing the activation of

Rho GTPases Cdc42 and Rac1 with SDF-1α stimulation. B&C, The graphs

depicts the band intensities normalized to the loading controls. Unstimulated

monocytic THP-1 cells had low basal levels of activated Rac1 and Cdc42.

Stimulation with SDF-1α led to a 6 fold increase (†, p<0.05) in GTP-bound Cdc42

and a 21 fold (†, p<0.05) increase in GTP-bound Rac1, compared with unstimulated

cells. Slit2 treatment alone had no effect on baseline activation of Rac1 and Cdc42.

However, Slit2 treatment significantly reduced SDF-1α mediated

activation of Cdc42 and Rac1. Monocytic THP-1 cells incubated with Slit2

had a 4 fold (†, p<0.05) increase in GTP-bound Cdc42 and a 12 fold

(†, p<0.05) increase in GTP-bound Rac1, compared to unstimulated cells.

The data represent the mean values ±SEM from 4 independent experiments.

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63

Figure 3.4 Slit2 inhibits Akt and Erk but not p38 MAPKs.

Human monocytic THP-1 cells were incubated with PBS vehicle or containing Slit2

(4.6 μg/mL) for 10 minutes and stimulated with SDF-1α (100ng/mL) for 5 minutes

and lysates collected. A, Western blots showing the activation of Erk, Akt and p38

MAPKs over time with SDF-1α stimulation. Blotting with phospho antibodies was

performed first, then the membranes were stripped and reprobed with antibodies for

the total protein to use for loading controls. B&C&D, Graphs depict the band

intensities normalized to the loading controls. Stimulation with SDF-1α induced

robust activation of the Akt and Erk MAPK pathways (Fig. 3.4A&B&C). However, no

activation of the p38 MAPK was observed (Fig 3.4A&D). Stimulation with SDF-1α

for 5 minutes led to a 2.9 fold increase (†, p<0.05) in phosphorylated Akt and a 3.2

fold (†, p<0.05) increase in phosphorylated Erk, compared with unstimulated cells

(Fig. 3.4A&B&C). Incubation with Slit2 alone had no effect on baseline activation of

Akt, Erk and p38 MAPKs (Fig. 3.4). However, incubation with Slit2 significantly

reduced SDF-1α-mediated activation of Akt and Erk MAPKs at 5 minutes, although

no effect was observed for p38 MARK (Fig. 3.4). Monocytic THP-1 cells incubated

with Slit2 had only a 1.7 fold (†, p<0.05) increase in phosphorylated Akt and a 1.9

fold (†, p<0.05) increase in phosphorylated Erk. Therefore, incubation with Slit2

decreased the activation of Akt and Erk MAPKs (Fig. 3.4A&B&C) at 5 minutes after

SDF-1α stimulation. However, incubation with Slit2 had no effect on the p38 MAPK

pathway (Fig. 3.4A&D). The data represent the mean values ±SEM from 8

independent experiments.

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Figure 3.5 Slit2 inhibits adhesion of monocytic THP-1 cells to human

umbilical vein endothelial cells.

HUVEC monolayers were stimulated with TNF-α for 4 hours and human

monocytic THP-1 cells were incubated with PBS vehicle alone or containing

Slit2 (4.6µg/mL) for 10 minutes. Slit-treatment alone had no effect on cell

adhesion, while adhesion to activated endothelial monolayers increased

from baseline to almost 400% (†, p<0.005). When THP-1 cells were

incubated with Slit2, adhesion was abolished to near baseline levels (‡, P<0.05).

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67

Figure 3.6 Slit2 inhibits adhesion of monocytic THP-1 cells to human

arterial endothelial cells.

HAEC monolayers were activated with TNF-α for 4 hours and human monocytic

THP-1 cells were incubated with PBS vehicle alone or containing Slit2 (4.6µg/mL).

Slit2 treatment alone had no effect of cell adhesion, whereas TNF-α stimulation of

the endothelial monolayers increased baseline adhesion by over 200% (†, p<0.01).

When monocytic THP-1 cells were incubated with Slit2, adhesion was

abolished to near baseline levels (‡, p<0.005).

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69

Figure 3.7 Slit2 inhibits monocyte recruitment in vivo.

Monocyte recruitment was determined in vivo using a model of murine peritonitis.

PBS alone or containing Slit2 (1.8µg/mouse) was administered intraperitoneally one

hour prior to sodium periodate induced peritonitis. Sodium periodate (1mg/mouse)

was injected intraperitoneally and peritoneal lavages were performed after 24 hours.

Sodium periodate alone induced vigorous peritonitis, reflected in the robust

monocyte recruitment (†, p<0.001). Intraperitoneal pretreatment with 1.8µg of Slit2

significantly inhibited monocyte recruitment (†, p<0.001) (n=5).

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71

Figure 3.8 Slit2 dose-dependently inhibits monocyte recruitment in vivo.

Monocyte recruitment was determine in vivo using a model of murine peritonitis.

Slit2 was administered intravenously via tail-vein injections at 4.6 µg/mouse, 460

ng/mouse and 46 ng/mouse prior to sodium periodate induced peritonitis. Sodium

periodate (1 mg/mouse) was injected intraperitoneally and peritoneal lavages were

performed after 24 hours. Sodium periodate alone induced vigorous peritonitis,

reflected in the robust monocyte recruitment (†, p<0.05). However, pre-treatment

with 4600 ng or 460 ng of Slit2 significantly diminished monocyte recruitment (†,

p<0.05).

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73

Figure 3.9 Slit2 inhibits monocyte/macrophage recruitment in vivo:

time-course.

Monocyte/macrophage recruitment was studied in an in vivo model of murine

peritonitis. Slit2 (1.8 µg/mouse) was administered intravenously via tail-vein

injections at 1, 4 and 10 days prior to sodium periodate induced peritonitis. Sodium

periodate (1 mg/mouse) was injected intraperitoneally and peritoneal lavages were

performed after 24 hours. Sodium periodate alone induced vigorous peritonitis,

reflected in the high number of recruited monocytes/macrophages (†, p<0.001).

Intravenous pre-treatment with Slit2 at 1 day and 4 days prior to induction of

peritonitis significantly diminished monocyte/macrophage recruitment († ,p<0.001)

at 1 day and (‡ ,p<0.01) at 4 days.

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75

Figure 3.10 Slit2 does not affect RAW 264.7 macrophage phagocytosis.

Murine RAW 264.7 macrophages were centrifuged together with IgG-

opsonized latex beads to initiate phagocytosis for 10 minutes. External

beads were then fluorescently labeled and images of at least 10 random

fields were captured. A, Representative images of control and Slit2

treated RAW macrophages performing IgG-mediated phagocytosis. B, Slit2

treatment had no effect on the phagocytic index (number of ingested

particles/ number of cells) of RAW 264.7 macrophages.

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CHAPTER 4

DISCUSSION & CONCLUSIONS

The aim of this project was to determine the effect of Slit2 on monocyte

chemotaxis, adhesion, and phagocytosis in vitro. Furthermore, we wanted to

determine whether Slit2 can inhibit monocyte recruitment in vivo. We have shown

that primary human monocytes and human monocytic THP-1 cells express the Slit2

receptor, Robo-1, and that Slit2 blocks monocyte migration in response to a SDF-1α

gradient. This finding is consistent with observations in the literature on the effect of

Slit2 on Robo-1 expressing cells. In fact, Slit2 has been shown to inhibit the

chemotaxis of a number of human hematopoetic cell types, including T-cells and

DCs (Guan et al., 2003; Kanellis et al., 2004; Prasad et al., 2007). In addition, we

have previously demonstrated that Slit2 inhibited the chemotaxis of circulating

human neutrophils (Tole et al., 2009).

Although Slit2 has been demonstrated to inhibit the chemotaxis of diverse

cell types, the mechanisms underlying its actions are not understood completely.

Chemotaxis is a complex process in which the cell polarizes to form a wide lamella

at the leading edge and a tail-like uropod in the trailing edge. Forward propulsion is

dependent on rapid turnover and polymerization of actin filaments. We have

previously shown that treatment of circulating human neutrophils with Slit2 reduced

chemokine-mediated generation of free barbed ends required for actin

polymerization at the leading edge (Tole et al., 2009; Glogauer et al., 2000). This

observation is consistent with previous findings in neuronal cells linking Slit2/Robo-1

signaling with proteins that are involved in the mobilization of the actin cytoskeleton,

77

such as Ena and srGAP (Bashaw et al., 2000; Wong et al., 2001). srGAP activates

Rho GTPases Rac and Cdc42, which are important for actin turnover in migrating

cells. Cdc42 is responsible for maintaining directionality, by driving the formation of

filopodia to sample extracellular cues, while Rac drives actin assembly in

lamellipodia required for forward propulsion during chemotaxis (Srinivasan et al.,

2003). We have found that Slit2 inhibits chemokine-mediated activation of Cdc42 in

human monocytic THP-1 cells. Our finding is consistent with studies in neuronal

cells and in our previous studies in primary human neutrophils, where Slit2 was

shown to inhibit activation of Cdc42, preventing these cells from undergoing

directional migration up a chemotactic gradient (Wong et al., 2001; Tole et al.,

2009). Furthermore, we have shown that Slit2 decreased chemokine-mediated

activation of Rac1 in monocytic THP-1 cells. Our observation is consistent with

Slit2-mediated suppression of Rac activation in studies of human vascular smooth

muscle cells and human T lymphocytes (Liu et al., 2006; Kanellis et al.,

2004;Prasad et al., 2007). Indeed, we have previously shown that Slit2 suppressed

the activation of Rac in circulating human neutrophils (Tole et al., 2009).

GPCR-mediated signaling in monocytes, as in other leukocytes, leads to

rapid phospholipid metabolism and the activation of MAPK pathways, including Akt,

Erk, and p38. Disruption of these pathways, using chemical inhibitors, has been

shown to inhibit chemotaxis (Heit et al., 2001). We have shown that Slit2 inhibited

chemokine-induced activation of Akt MAPK. This suggests that Slit2 may affect

phospholipid metabolism, specifically the generation of PIP3 at the plasma

membrane, which is required for the recruitment and activation of Akt MAPK. Our

78

observation is consistent with a previous report which found that in human breast

cancer cells, Slit2 inhibited chemokine-mediated activation of PI3K, and subsequent

activation of Akt MAPK (Prasad et al., 2004). Moreover, Slit2 was also shown to

inhibit SDF-1α-induced activation of Akt in Jurkat T cells (Prasad et al., 2007).

However, this trend in Slit2-mediated inhibition of SDF-1α-induced Akt MAPK

activation is inconsistent with our study in circulating human neutrophils, where Slit2

was shown to have no effect on the fMLP-induced activation of Akt, Erk and p38

MAPKs (Tole et al., 2009). We have also found that Slit2 inhibited SDF-1α-induced

activation of Erk MAPK. This finding is consistent with a study in human breast

cancer cells, where Slit2 inhibited SDF-1α-induced activation of Erk MAPK (Prasad

et al., 2004). However, our observation is inconsistent with studies in granulocytic

cells (Wu et al., 2001) and in our previous report in circulating human neutrophils,

where no Slit2-mediated inhibition in Erk MAPK activation was observed (Tole et al.,

2009). Finally, we have shown that Slit2 did not affect SDF-1α-induced p38 MAPK

activation, consistent with findings on Jurkat T lymphocytes and in our previous

report in circulating human neutrophils (Prasad et al, 2007; Tole et al., 2009). These

differential effects of Slit2 on MAPK activity may be attributable to differences in cell

types used or in the chemoattractants used for stimulation. For example, our

previous study on circulating human neutrophils utilized the bacterial product fMLP

to stimulate the MAPK pathways, while this study utilized the chemokine SDF-1α.

This is further supported by the consistency of our findings with those in human

breast cancer cells, where SDF-1α was also utilized for MAPK activation.

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In order to be recruited from circulation and extravasate, monocytes must

undergo a series of coordinated interactions with vascular endothelial cells. During

acute inflammation, or in chronic inflammatory conditions such as atherosclerosis,

the local cytokine microenvironment activates vascular endothelial cells to express

increased levels of adhesion molecules. These activated endothelial cells are able

to efficiently capture circulating leukocytes, including monocytes, facilitating their

arrest and diapedesis across the vessel wall. To determine if Slit2 affects monocyte

adhesion, we performed adhesion assays using endothelial monolayers activated

with the proinflammatory cytokine TNF-α. We have shown that Slit2 inhibited

adhesion of monocytic THP-1 cells to activated endothelial monolayers, specifically

HUVECs and HAECs. Our observations are consistent with a study of human

breast cancer cells which found that Slit2 inhibited CXCL12-mediated adhesion to

ligands such as fibronectin and collagen (Prasad et al., 2004). Furthermore, Slit2

has previously been shown to block Jurkat T cell adhesion to activated HUVEC

monolayers. Consistent with this line of evidence is a recent report showing that the

Slit/Robo pathway functions to antagonize E-cadherin-mediated cell adhesion of

Drosophila cardioblasts during development (Santiago-Martnez et al., 2008). Since

Rho GTPases are also involved in the actin mobilization required for cell adhesion,

it is likely that the signaling events downstream of Slit2/Robo-1 influence the ability

of the cell to form adhesive contacts. Further studies should explore the effect of

Slit2 on the adhesion of monocytes to Ig superfamily ligands, ICAM-1 and VCAM-1,

which are important in physiological cell adhesion. In addition, GPCR-mediated

activation of monocytes during rolling induces outside-in and inside-out signaling

80

pathways which lead to changes in integrin avidity on monocytes. Thus, future

studies should also explore the effect of Slit2 on transient upregulation of monocyte

integrin affinity induced by chemokines and other chemoattractants, using the

methods of Chan et al. (2003).

Signals elicited by chemokines and other chemoattractants activate

leukocyte β1- and β2-integrins, resulting in tight adhesion to the vascular

endothelium and induction of cytoskeleton-driven leukocyte migration. Many

chemokines can bind to transmembrane heparan sulphate proteoglycans on the

luminal surface of the endothelium in order to be presented to leukocytes

(Spillmann et al., 1998; Halden et al., 2004). When chemokines bind to these

proteoglycans, the chemokine receptor binding site remains exposed (Proudfoot et

al., 2000). This allows the chemokine to interact with its chemokine receptor

expressed on leukocytes in order to elicit a rapid integrin activation signal. Complex

signaling networks regulate the affinity of integrins, via spatial separation and

unfolding of the two integrin chains, and the avidity of integrins, by increasing lateral

mobility and clustering (Kim et al., 2003).

RAP1 and RAP2 are small GTPases of the RAS family that play an important

role in chemokine-mediated inside-out signaling, which activates the integrins LFA-1

and VLA-4 (Katagiri et al., 2000; McLeod et al., 2004). RAP1 is expressed by most

haematopoietic cells, cycling between an inactive GDP-bound form and an active

GTP-bound form. Like other GTPases, its activity is regulated by the GEF exchange

factor directly activated by cyclic AMP (EPAC) and the GAPs signal-induced

proliferation associated antigen 1 (SPA1) and RAPGAPII (Bos, L., 2003). Recent

81

reports have shown that CCL21 and SDF-1α rapidly activate RAP1 to its active,

GTP-bound, form (Bos, L., 2003, Shimonaka et al., 2003). The chemokine-mediated

activation of RAP1 induces LFA-1- and VLA-4-dependent adhesion and migration

(McLeod et al., 2004; Shimonaka et al., 2003). This is controlled by leukocyte

adhesion to ICAM-1 and VCAM-1 expressed by activated endothelial cells. The

importance of RAP1 is highlighted in studies of leukocytes transfected with

constitutively active RAP1, which are able to adhere and migrate independently of a

chemokine signal (Shimonaka et al., 2003; Tohyama et al., 2003). Furthermore,

transfection with RAP1 GAPs, SPA1 or RAPGAPII, blocks integrin-mediated cell

adhesion and migration. RAP1 modulates integrin affinity by binding to RAPL. This

complex then activates integrins by binding to a conserved GLY-PHE-PHE-LYS-

ARG motif on the integrin α-chain. Interestingly, overexpression of RAPL has been

shown to activate integrin-mediated cell adhesion, while overexpression of a RAPL

mutant that is incapable of binding RAP1 inhibits adhesion (Katagiri et al., 2003).

Therefore, the chemokine-induced activation of RAP1 is important for the signaling

networks that activate leukocyte integrins. In order to gain a better mechanistic

understanding of the Slit2-mediated inhibition of monocyte adhesion to activated

endothelium, future studies should explore the effect of Slit2/Robo-1 signaling on

the activation of RAP1 and on the activity and localization of its GEFs and GAPs.

In addition to RAP1 and RAPL, there are other signaling networks that

contribute to the rapid chemokine-induced integrin activation. Talin is a cytoskeletal

protein consisting of a globular head and a rod-like domain. The head domain can

bind to the ASN-PRO-XAA-TYR/PHE motif on the β-chain of integrins. This binding

82

activates the integrin by keeping the cytoplasmic domains of the α- and β-chains

separated, allowing the unfolding of the extracellular domain and exposure of the

ligand-binding pocket (Kim et al., 2003; Tadokoro et al., 2003). Although the

mechanism by which talin regulates chemokine-mediated integrin activation is

incompletely understood, it is believed to be required for integrin activation

downstream of several signaling pathways (Tadokoro et al., 2003). Furthermore, the

protease calpain can cleave talin between the head and rod domains. Once

cleaved, the head domain has a six fold higher affinity for the integrin β-chain than

does the intact molecule, allowing for more efficient integrin activation (Calderwood,

A., 2004). This cleavage may allow for further regulation of talin-mediated integrin

activation. In addition, the binding of phosphoinositol phosphate kinase type Iγ to

talin regulates talin-integrin interactions by enhancing the binding affinity of talin for

the integrin β-chain (Di Paulo et al., 2002; Ling et al., 2002; Martel et al., 2001).

Finally, phosphorylation of a tyrosine residue on the integrin talin-binding motif by

SRC-family kinases prevents talin-mediated integrin activation (Datta et al., 2002;

Sakai et al., 2001). Thus, talin may play a regulatory role in the chemokine-induced

integrin-mediated leukocyte adhesion. Future experiments to elucidate the

mechanism by which Slit2/Robo-1 signaling inhibits monocyte adhesion to activated

endothelial cells should include an investigation of talin. Specifically, the effect of

Slit2/Robo-1 signaling on talin phosphorylation and cleavage should be addressed.

Another important regulatory signaling pathway leading to integrin activation

involves RhoA. RhoA is a member of the RAS superfamily of GTPases, and is

involves in integrin activation, membrane ruffling, stress fiber formation and cell

83

migration (Alblas et al., 2001; Ridley et al., 1994; Laudanna et al., 2002). Several

studies have demonstrated that blocking RhoA or its downstream targets increases

monocyte adhesion to ICAM-1 ligand (Worthylake et al., 2003; Smith et al., 2003).

The effect of RhoA is complex, as it is activated by both chemokines and ligand-

bound integrins on adherent cells. Several RhoA-interacting adaptors are required

for β2-integrin-dependent adhesion to ICAM-1, and these provide another

mechanism for the tuning of integrin-mediated adhesion induced by chemokines.

Thus, RhoA regulates integrin-mediated adhesion via the activation of integrins, the

regulation of lateral integrin mobility in the plasma membrane and the effect on the

actin cytoskeleton. Future experiments to shed light on the mechanism by which

Slit2/Robo-1 signaling inhibits monocyte adhesion to activated endothelial cells

should include an investigation of RhoA activity. In addition, the effect of Slit2/Robo-

1 signaling on RhoA GAPs and GEFs should be addressed.

Another signaling network regulating integrin-mediated leukocyte adhesion

via the modulation of cell polarity involves atypical protein kinase C-δ (PKC-δ)

(Wang et al., 2003; Etienne-Manneville et al., 2003). Chemokines induce the kinase

activity of PKC-δ, via its interaction with PI3K and RhoA, resulting in its targeting to

the plasma membrane where it leads to increased integrin mobility (Giagulli et al.,

2004). This is required for the clustering of activated integrins, leading to high

integrin avidity, allowing leukocytes to rapidly induce firm adhesion during rolling.

Furthermore, it has been shown that further activation of adherent cells by

chemokines results in PKC-δ localization to the lamellipodium (Wang et al., 2003).

Therefore, PKC-δ plays a role in leukocyte polarity and in the reinforcement of

84

integrin-mediated cell adhesion. Future experiments into the mechanism by which

Slit2/Robo-1 signaling inhibits monocyte adhesion should include an investigation of

PKC-δ. Specifically, the effect of Slit2/Robo-1 signaling on PKC-δ activation and

membrane targetting should be addressed.

Since we found that Slit2 can inhibit monocyte chemotaxis and adhesion in

vitro, we next sought to investigate if it will work in vivo. To test the in vivo

recruitment of monocytes to inflammatory foci, we used a sodium periodate-induced

model of experimental murine peritonitis. When Slit2 is administered

intraperitoneally, it gets absorbed systemically (Kanellis et al., 2004). We have

shown that when Slit2 was administered intraperitoneally an hour prior to inducing

peritonitis, monocyte recruitment to the peritoneal cavity was significantly abolished.

This supports our previous observation which demonstrated that Slit2 administered

intraperitoneally was able to effectively diminish neutrophil recruitment in the same

murine model of experimental peritonitis induced by sodium periodate (Tole et al.,

2009). Thus, Slit2 may inhibit the recruitment of different subsets of leukocytes in

vivo. Moreover, we have previously shown that Slit2 can inhibit the recruitment of

neutrophils in vivo to diverse chemoattractants administered intraperitoneally,

including MIP-1 and C5a (Tole et al., 2009). To determine the dose of Slit2 required

to exert an optimal biological effect, we administered Slit2 intravenously at

decreasing doses (46 ng - 4.6 µg). We have found that Slit2 was able to exert a

significant effect on monocyte recruitment in vivo even when administered at 460

ng/mouse, although the effect wore off with further dilution. Furthermore, due to the

potent effect of Slit2 on leukocyte recruitment in vivo, we sought to determine the

85

timing of Slit2 administration required to induce an optimal biological effect. We

found that Slit2 significantly abolished monocyte recruitment when administered 1

day or 4 days prior to induction of peritonitis, although the effect of Slit2 was slightly

diminished at 4 days, compared to the effect at 1 day. Literature on the exogenous

application of Slit2 is scarce. These findings suggests that Slit2 can persist in the

circulation in order to exert a biological effect for up to 4 days. Due to its extensive

glycosylation and hence 'stickiness', Slit2 may associated with GAGs on the

endothelial lumen. This is consistent with findings that human full-length Slit2 and

N-terminal Slit2 are tightly associated with the cell membrane (Brose et al., 1999).

This property allows Slit2 to be concentrated on the endothelial lumen, where it can

signal and exert an effect on leukocytes as they interact with the vessel wall. In

addition, the extensive glycosylation on Slit2 may confer protection from

degradation by proteases, which may further increase its biological half life.

Phagocytosis is a vital monocyte/macrophage function required for innate

and adaptive immunity. Once monocytes are recruited to inflammatory foci, they

must engulf pathogens for immune clearance or antigen presentation. Since Rho

GTPases are involved in the actin mobilization required to form pseudopods and

engulf particles, we speculated that Slit2 might have an effect on monocyte

phagocytosis. To determine the effect of Slit2 on monocyte phagocytosis, we

performed phagocytosis assays with Ig-opsonized latex beads. We have found that

Slit2 had no effect on monocyte phagocytosis. Although this finding was surprising,

since Rho GTPases Rac and Cdc42 are involved in phagocytosis, this finding is

consistent with a recent report demonstrating that Slit2 treatment had no effect on

86

neutrophil phagocytosis of Ig-opsonized latex beads (unpublished observations).

This observations may be due to the fact that Slit2 only acts on polarized or

polarizing cells. Thus, further studies are needed to elucidate the precise effect of

Slit2 on leukocyte phagocytosis.

In our current study, we have shown that Slit2 can inhibit the chemotaxis of

monocytic THP-1 cells to gradients of SDF-1α. It is likely that this inhibition is

mediated by the ability of Slit2 to inhibit Rho GTPases Cdc42 and Rac1, and

therefore, the polymerization and turnover of actin, and mobilization of the actin

cytoskeleton. Consistent with this hypothesis is our observation that Slit2 inhibited

the adhesion of monocytic THP-1 cells to activated endothelial cells, since adhesion

is also dependent on Rho GTPase-mediated actin mobilization. However, it is

surprising that Slit2 had no effect on monocyte phagocytosis, as this process is also

dependent on Rho GTPases-mediated dynamic actin regulation. Although further

studies into the mechanism of Slit2 action is necessary, our data strongly support

the use of Slit2 as a novel anti-inflammatory agent. Currently, many anti-

inflammatory agents act via general suppression of immune activation and function,

and thus have serious side effects. Although Slit2 may also have

immunosuppressive effects, targeted local delivery of Slit2 could be utilized to

prevent localized inflammatory cell recruitment and the associated tissue damage,

while preserving the overall function of the immune system in the host. Our data

demonstrate that Slit2 can selectively inhibit monocyte recruitment and adhesion to

the vessel wall, while preserving vital immune functions such as phagocytosis.

87

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APPENDIX 1 The Axonal Repellent, Slit2, Inhibits Directional Migration of Circulating Neutrophils

A1.1 Abstract

In inflammatory diseases circulating neutrophils are recruited to sites of injury.

Attractant signals are provided by many different chemotactic molecules, such that

blockade of one may not effectively prevent neutrophil recruitment. The Slit family of

secreted proteins, and their transmembrane receptor, Roundabout (Robo), repel

axonal migration during central nervous system development. Emerging evidence

shows that by inhibiting the activation of Rho-family GTPases, Slit2/Robo also

inhibit migration of other cell types towards a variety of chemotactic factors, in vitro

and in vivo. The role of Slit2 in inflammation, however, has been largely unexplored.

We isolated primary neutrophils from human peripheral blood and mouse bone

marrow, and detected Robo-1 expression. Using video-microscopic live cell

tracking, we found that Slit2 selectively impaired directional migration, but not

random movement, of neutrophils towards formyl-methionyl-leucyl-phenylalanine

(fMLP). Slit2 also inhibited neutrophil migration towards other chemoattractants,

namely C5a and interleukin (IL)-8. Slit2 inhibited neutrophil chemotaxis by

preventing chemoattractant-induced actin barbed end formation and cell

polarization. Slit2 mediated these effects by suppressing inducible activation of

Cdc42 and Rac2, but did not impair activation of other major kinase pathways

involved in neutrophil migration. We further tested the effects of Slit2 in vivo using

mouse models of peritoneal inflammation induced by sodium periodate, C5a, and

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macrophage inflammatory protein-2 (MIP-2). In all instances, Slit2 effectively

reduced neutrophil recruitment (p < 0.01). Collectively, these data demonstrate that

Slit2 potently inhibits chemotaxis, but not random motion, of circulating neutrophils,

and point to Slit2 as a potential new therapeutic for preventing localized

inflammation.

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A1.2 Introduction

Neutrophils are a critical component of the innate immune system and

provide the first line of defense against bacterial and fungal pathogens. During an

inflammatory response, neutrophils are recruited to sites of inflammation in a series

of coordinated interactions with vascular endothelial cells. Traffic signals are

provided by diverse chemoattractant molecules, including chemokines such as IL-8,

and bacterial products such as formylated peptides. These chemoattractants recruit

circulating neutrophils to sites of inflammation, and activate recruited neutrophils to

adhere firmly to the endothelium. While their potent anti-microbial arsenal makes

neutrophils efficient at fighting microorganisms, it is also capable of causing injury to

the surrounding tissue. Indeed, neutrophils inflict significant tissue damage in

inflammatory conditions including ischemia-reperfusion injury of solid organs, acute

respiratory distress syndrome, and rheumatoid arthritis [1-4]. Once recruited to sites

of injury, infiltrating neutrophils release reactive oxygen species and degradative

enzymes, fuelling local tissue destruction.

Anti-inflammatory drugs such as aspirin and glucocorticoids are widely used,

and yet, have shown modest success in reducing neutrophil-mediated injury. These

drugs attenuate activation of transcription factors such as NF-κB, thereby reducing

expression of cytokines [5]. An alternative approach to prevent neutrophil-mediated

tissue damage would be blockade of chemotactic pathways that recruit neutrophils

to sites of inflammation. Indeed, some chemokine receptor antagonists or blocking

antibodies have shown success in animal models and are undergoing clinical trials

[6]. However, given the number of chemoattractant signals that recruit neutrophils, it

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is unlikely that targeting a single chemokine/chemokine receptor pathway would

achieve widespread clinical success. Thus, localized general blockade of

inflammatory chemoattractants could represent a clinically useful strategy to reduce

neutrophil-mediated tissue damage.

Clues as to how generalized blockade of neutrophil chemoattractant signals

might be realized are provided in the neurodevelopmental literature. The Slit family

of secreted proteins, together with their transmembrane receptor Roundabout

(Robo), repel migration of axons and neurons during development of the central

nervous system. Slit is expressed along the midline of the developing central

nervous system and its interaction with Robo prevents axons from repeatedly and

randomly crossing the midline [7, 8]. While the importance of Slit/Robo interactions

in development has been demonstrated, the intracellular signaling pathways that

lead to Slit-mediated inhibition of migration remain unclear. Data from Drosophila

suggests that Abelson kinase (Abl) and Enabled (Ena) proteins associate with the

intracellular domains of Robo-1 and may be involved in the repulsive response to

Slit2 [9]. Addition of extracellular Slit2 to neuronal cells results in the recruitment of

soluble Slit Robo guanosine triphosphatase (GTPase) activating protein 1 (srGAP1)

to the cytoplasmic tail of Robo-1 [10].

In addition to neuronal cells, Slit2 and Robo-1 also inhibit migration of other

cell types, including vascular smooth muscle cells, breast cancer cells, and brain

tumor cells [11-13]. Several studies have demonstrated that Slit2 inhibits migration

of haematopoietic cells, including murine macrophages, cultured cells of

granulocytic lineage, dendritic cells, and primary human T-lymphocytes, towards

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chemoattractant signals [14-17]. Importantly, Slit2 not only inhibits cell migration

towards one type of chemoattractant signal, but towards many diverse signals,

including platelet-derived growth factor (PDGF) and the chemokines, CXCL12 and

CCL2 [12,13, 16, 17]. In vivo, Slit2 inhibits neoangiogenesis by impairing pathologic

migration of endothelial cells to vascular endothelial growth factor [18]. Existing data

point to a role for Slit2 as a generalized “anti-migration” signal, which universally

inhibits cell migration. However, the potential use of Slit2 to prevent inflammation

has been largely unexplored. In particular, there is a paucity of data addressing the

effects of Slit2 on migration of human leukocytes, especially neutrophils. Moreover,

the mechanisms by which Slit2 mediates its anti-migratory effects are incompletely

understood.

The aim of this study was to assess, in real-time, the effect of Slit2 on

recruitment of primary neutrophils. We observed that primary human and murine

neutrophils express the Slit2 receptor, Robo-1, and that Slit2 inhibits directional

migration, but not random migration, of neutrophils towards a chemotactic stimulus.

Our studies demonstrate that Slit2 mediates these effects by preventing

chemoattractant-induced cell polarization and generation of actin free barbed ends,

a pre-requisite for directional migration of neutrophils. Our data further suggest that

Slit2 prevents chemoattractant-induced free barbed end formation by suppressing

inducible activation of the small GTPases, Cdc42 and Rac2, but does not affect

activation of other major kinase pathways involved in neutrophil migration. To

investigate whether Slit2 prevents neutrophil chemotaxis in vivo, we used mouse

models of peritoneal inflammation, and observed a significant reduction in the

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number of neutrophils recruited to the peritoneum in response to diverse

inflammatory stimuli, in the presence of Slit2 [19]. Taken together, these data

indicate a novel role for the axonal repellent, Slit2, as an anti-inflammatory agent

which specifically prevents chemotactic trafficking of circulating neutrophils.

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A1.3 Materials and Methods

Reagents and antibodies. Unless otherwise stated, reagents were purchased from

Sigma-Aldrich (St. Louis, MO). Polymorphprep neutrophil separation medium was

purchased from Axis- Shield, Norway. The following primary antibodies were used:

anti-Robo-1 (Abcam, Cambridge, MA, and Santa Cruz Biotechnology, Santa Cruz,

CA), anti-myc 9E10 (Covance, QC, Canada), anti-human Cdc42 (Cell Signaling,

Danvers, MA), anti-human Rac2 (Upstate Biotechnology, Lake Placid, NY), anti-

mouse CD3 (BD Biosciences, Mississauga, Ontario, Canada), anti-B220 (BD

Biosciences), anti-NK1.1 (BD Biosciences), anti-F4/80 (Serotec, Raleigh, NC), anti-

Erk, anti-phospho-Erk, anti-p38 MAPK, anti-phospho-p38 MAPK, anti-Akt, and anti-

phospho-Akt. Rhodamine-conjugated phalloidin was from Invitrogen Canada

(Burlington, Ontario, Canada). The following secondary antibodies were used: Cy3-

conjugated anti-rabbit IgG, Cy2- conjugated anti-human IgG, phycoerythrin (PE)-

conjugated anti-rat IgG and anti-mouse IgG (Jackson Immunoresearch

Laboratories, Bar Harbor, ME), and horseradish peroxidase-conjugated anti-rabbit

IgG and anti-mouse IgG (Jackson Immunoresearch Laboratories). C5a was

purchased from Biovision, Inc. (Mountain View, CA), interleukin-8 (IL-8) from

Invitrogen, and macrophage inflammatory protein-2 (MIP-2) from R&D Systems

(Minneapolis, MN).

Isolation of primary human and murine neutrophils. Human blood was obtained from

healthy volunteers and neutrophils were isolated using two methods. For

experiments testing the activation of Rac and Cdc42, neutrophils were isolated by

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dextran sedimentation as described with slight modifications [20]. Briefly, two

volumes of blood were mixed with one volume of 6% dextran T-500 in 0.9% NaCl

and set at room temperature until clear separation of layers was seen (about 30

min). The leukocyte-rich upper layer was collected and centrifuged at 260g at

room temperature for 5 min. The cell pellet was re-suspended in a volume of 0.9%

NaCl equal to the starting volume of blood, laid onto 10 ml of Ficoll-hypaque

solution, and centrifuged at 460g for 30 min. Red blood cells were lysed by adding

20 ml of ice-cold 0.2% NaCl for 30 s, resuspended in 20 ml of ice-cold 1.6% NaCl

and centrifuged at 250g at 4°C for 5 min. Neutrophils were re-suspended in ice-cold

PBS with 0.5% BSA. Cells were kept on ice for subsequent experimental use. The

purity of neutrophils isolated in this manner was assessed by modified Wright-

Giemsa stain (Hema-Tek Stain Pack; Bayer, Elkhart, IN) using an automated

stainer (Hema-Tek 2000; Bayer), and was consistently greater than 95%. For all

other experiments, the Polymorphprep gradient separation procedure was

performed according to the manufacturer‟s recommendations. Purified neutrophils

were suspended in PBS without calcium and kept at room temperature. Prior to use,

the neutrophils were re-suspended in HBSS with 1mM CaCl2 and 1mM MgCl2.

Experiments were performed within 1-2 h of isolation of neutrophils. Cell purity was

consistently >85-90%. Cell viability was >98% by Trypan blue exclusion. For

RTPCR experiments, a QIAmp RNA Blood Mini Kit (QIAGEN, Ontario, Canada)

was used to isolate total RNA from human leukocytes isolated from whole blood,

according to the manufacturer‟s specifications. Primary murine neutrophils were

isolated as previously described [19, 21]. Briefly, adult CD1 mice were killed by CO2

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inhalation. Femurs and tibias were removed and bone marrow was extracted. Bone

marrow cells were layered onto discontinuous Percoll gradients of 81%/65%/55%.

Mature neutrophils were isolated from the 81%/65% interface. More than 85% of

cells were neutrophils as assessed by Wright-Giemsa staining.

Slit2 expression and purification. Stable human embryonic kidney (HEK) 293 cell

line expressing full-length human Slit2 with a c-myc-tag at its carboxyl terminus was

a kind gift from Drs. Rolando del Maestro (McGill University, Montreal, Canada) and

Yi Rao (Washington University, St. Louis, MO) and grown as described [22].

Recombinant Slit2 was purified from the conditioned medium using two methods.

Conditioned medium was concentrated and Slit2 purified by affinity chromatography

using anti-c-myc Ab 9E10 (Covance, QC, Canada) and Size Primary

Immunoprecipitation kit (Thermo Scientific, Rockford, IL) following the

manufacturer's instructions. Slit2 was also obtained by Superdex-200 size exclusion

chromatography. Briefly, conditioned medium was concentrated using Centricon

Plus-20 (Millipore, Billerica, MA) and loaded onto the column [16]. The column was

then washed with PBS and fractions containing Slit2 were pooled, concentrated,

aliquoted and stored in –80°C before use. The presence of Slit2 was verifed using

silver staining and immunoblotting with anti-myc Ab (Supplementary Figure

1A & B). The above protocol was repeated with conditioned medium from control

HEK293 cells to obtain control medium. This preparation of Slit2 was titrated and

used at a concentration of 0.6 µg/ml. In parallel assays, control medium was used in

lieu of Slit2.

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Large scale preparation of Slit2 was performed by transfection of HEK293-EBNA1

cells. Briefly, human Slit2 cDNA (MGC: 177513; aa 26-1529 of NP_004778) was

amplified using forward (CTATCTAGACCTCAGGCGTGCCCGGCGCAGTGC) and

reverse (CTAGGATCCGGACACACACCTCGTACAGC) primers containing XbaI

and BamHI restriction sites. The amplified cDNA was cloned into the pTT28 vector

digested with NheI and BamHI. The pTT28 vector is a derivative of the pTT5 vector

[23, 24] and contains a synthetic and codon-optimized signal peptide

(MGELLLLLLLGLRLQLSLG) and a C-terminal (His)8G tag separated by NheI and

BamHI restriction sites. HEK293-EBNA1 cells (clone 6E) were transfected with 1

µg/ml cDNA as previously described [25]. Culture medium was harvested

120 h post-transfection, clarified by centrifugation (4,000 x g for 15 min), and filtered

through a 0.45 µm membrane. Slit2 secreted into the medium was purified by

immobilized metal-affinity chromatography using a Fractogel-cobalt column

equilibrated in PBS. Following washing steps with 5 column volumes (CV) of Wash1

Buffer (50 mM sodium phosphate pH 7.0 and 300 mM NaCl) followed by 5 CV of

Wash2 Buffer (50 mM sodium phosphate pH 7.0 , 300 mM NaCl and 25 mM

imidazole), bound Slit2 was eluted with Elution Buffer (50 mM sodium phosphate pH

7.0, 300 mM NaCl and 25 mM imidazole). The pooled eluted material was

immediately desalted on Econo-Pac™ 10 columns (Bio-Rad Laboratories,

Mississauga, ON) previously equilibrated with PBS according to the manufacturer‟s

specifications. Protein concentration was determined by absorbance at 280 nm

using a calculated Slit2 molar extinction coefficient of 114600

(http://ca.expasy.org/tools/protparam.html). For Western blots, proteins were

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resolved on reducing SDS-PAGE (4–12% Nu-PAGE Bis-Tris gradient gel,

Invitrogen) followed by transfer to a 0.2 mm Protran nitrocellulose membrane

(Schleicher & Schuell, Keene, NH) in Tris-glycine buffer for 1 h at 300 mA. Purity

was verified by Ponceau staining and immunoblotting (Supplementary Figure 1C &

D). The membrane was incubated in blocking reagent (Roche Diagnostics, Laval,

Canada), and then probed with anti-polyHis-HRP Ab (Sigma-Aldrich) for 1 h

(Supplementary Figure 1D). Detection was performed using BM

Chemiluminescence Blotting Substrate (Roche Diagnostics) with a Kodak Digital

Science Image Station 440cf equipped with Kodak Digital Science 1D image

analysis software version 3.0 (Eastman Kodak, New York, NY). We measured

endotoxin levels in purified Slit2 stock preparations using ToxinSensor

Chromogenic LAL Endotoxin Assay Kit (GenScript Corp., Piscataway, NJ).

Endotoxin concentrations ranged from 0.2-0.8 ng/ml, yielding final experimental

concentrations of 12-40 pg/ml which are well below those thought to activate

leukocytes [26]. To verify this point, we added similar concentrations of endotoxin in

neutrophil Transwell assays, and found that such levels of endotoxin had no effect

on neutrophil migration (Supplementary Figure 2).

RT-PCR. RNA isolation and RT-PCR were performed using the QIAamp RNA blood

mini kit and the QIAGEN one-step RT-PCR kit (QIAGEN, Missisauga, ON) as

described [13]. As previously described, the following primers specific for Robo-1

were used: GGCCCCACTCCCCCTGTTCG (forward primer) and

TCCTCTTCTGGCGCATCCGTATCC (reverse primer) [13]. Amplified products were

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analyzed by electrophoresis on 2% agarose gels containing ethidium bromide to

confirm primer specificity and PCR product size (278 bp).

Immunofluorescent labeling. Primary human and mouse neutrophils were allowed to

settle onto fibronectin-coated coverslips and to adhere for 3 minutes at room

temperature. The cells were fixed with 4% paraformaldehyde for 10 min at 4°C.

Neutrophils were stained with rabbit anti-Robo-1 Ab (1 µg/ml) for 2 h, washed and

then incubated with anti-rabbit-Cy3 secondary Ab for 1 h. In some experiments,

human or mouse neutrophils were incubated with fMLP (1 µM) for 3 min, following

incubation with purified Slit2 (4.5 µg/ml). Cells were fixed, permeabilized, and

incubated with rhodamine-conjugated phalloidin (1:500) for 30 min to visualize actin.

A Leica DMIRE2 spinning disc confocal microscope (Leica Microsystems, Toronto,

Ontario, Canada) equipped with a Hamamatsu back-thinned EM-CCD camera and

Volocity software (Improvision Inc., Lexington, MA) was used to capture images.

Flow cytometry. Cell surface expression of Robo-1 was verified by incubating

human and mouse neutrophils with anti-Robo-1 Ab, followed by PE-conjugated

secondary Ab. Analysis was performed using a FACScalibur flow cytometer (BD

Biosciences) and FlowJo software (Tree Star, Inc., Ashland, OR), as previously

described [27, 28].

Immunoblotting. Freshly isolated human or mouse neutrophils were pre-treated with

either control medium or Slit2 for 10 min and then activated with fMLP (1 µM). Cells

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were lysed using ice-cold 2x lysis buffer (1 x = 50 mM Tris, pH 7.5, 10% glycerol,

100 mM NaCl, 1% NP- 40, 5 mM MgCl2, 1 mM DTT, 1 mM PMSF, 1/100 protease

inhibitor cocktail and 1 mM NaVO3). Samples were run on SDS-PAGE, transferred

to 0.2 mm PVDF (Millipore) membrane and probed for Robo-1 or for both

phosphorylated and total Akt, Erk and p38 MAP kinase. Immunoreactive bands

were visualized by enhanced chemiluminescence (Amersham Biosciences, UK Ltd,

Buckinghamshire, UK) recorded on x-ray film. Prior to performing experiments, a

time-course study was performed to determine the optimal point at which to

measure phosphorylation of Akt, Erk, and p38 MAPK following exposure to fMLP.

Of samples harvested at 15 - 180 s, the maximum signal was observed at 30 s, and

therefore, a 30 s timepoint was used for all subsequent experiments.

Migration assay. Freshly isolated neutrophils (106 cells/ 100µl) were incubated with

medium alone, Slit2 (0.6 µg/ml), or control medium at 37°C for 10 minutes. Cells

were loaded into the top chamber of a 3 µm Transwell insert (Corning Life

Sciences, Corning, NY) in a 24-well plate. A coverslip was added to the bottom

chamber which was filled with 600 µl of HBSS alone, fMLP (1 µM), C5a (2 µg/ml), or

IL-8 (0.1 µg/ml) [29-32]. Into the bottom chamber Slit2, control medium, or HBSS

was dispensed. Transwell plates were incubated for 1 h at 37°C. To determine the

number of neutrophils which had migrated from the top to the bottom chamber, the

filter was removed and neutrophils in the lower chamber were rapidly spun down

onto the coverslips, fixed with 4% paraformaldehyde, washed, and labeled with

DAPI. A Leica DMIRE microscope was used to take representative 40x and 63x

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high-power images. A Nikon light microscope was used to count at least 10 random

fields from each coverslip. The data represent the mean value ± SEM from at least

4 independent experiments for each treatment condition.

Micropipette chemotaxis assays. To measure neutrophil migration, round glass

coverslips (25-mm diameter; Thomas Scientific, Swedesboro, NJ) coated with

fibronectin were mounted in Leiden chambers, overlaid with 0.5 ml of the indicated

solution, and placed on the heated stage of a Leica DM IRB microscope (Leica

Microsystem, Richmond Hill, Ontario, Canada). Next, a 100 µl aliquot of the

neutrophil suspension containing 106 cells was added, and cells allowed to

settle for 10 min. To induce chemotaxis, a point-source of chemoattractant was

delivered using a glass micropipette [33-36]. Micropipettes were prepared from

borosilicate capillaries with an outer diameter of 1.0 mm and an inner diameter of

0.78 mm (Sutter, Novato, CA) using a model P-97 micropipette puller (Sutter). The

tips of the micropipettes were 1.0 µm in diameter. Precise positioning of the

micropipette in the visual field was accomplished using a model 5171

micromanipulator (Eppendorf, Hamburg, Germany). Although the distance between

the pipette and the individual cells adherent to the coverslip varied, the initial

average distance of the cells under observation (i.e., those in the microscopic field

under observation) ranged between 40 and 50 µm. The pipette remained stationary,

and diffusion of the chemoattractant generated a standing gradient [33-36]. Images

were acquired every 10 s until completion of the experiment. Only cells which

started and remained in the field of view over the entire course of videocapture were

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analyzed. Using VolocityTM software (Improvision, Waltham, MA), the distance

traveled was measured by tracking the centroid of each cell over time. Four different

measures of chemotactic activity were assessed: total migration (distance), net

migration (displacement), speed (distance/time) and directionality

(displacement/distance). Total migration was defined as the sum of the absolute

distances traveled in all the individual time intervals. The net migration was

calculated as the difference between the initial distance of the cell with respect to

the pipette and that at the end of the experiment. Migration speed was calculated by

dividing the total distance travelled over the elapsed time. Directionality was

measured by obtaining a ratio of displacement over distance.

Actin free barbed end assay. To assess the effects of Slit2 on fMLP-induced actin

polymerization, actin nucleation activity was measured as enhancement of pyrene

actin fluorescence as previously described [21, 37, 38]. Briefly, human neutrophils

(5x106 /ml) were permeabilized for 10 s using 0.1 vol of OG buffer (PHEM buffer

containing 4% octyl glucoside, 10 µM phallacidin, 42 nM leupeptin, 10 mM

benzamidine, and 0.123 mM aprotinin) or NP-40 (final concentration of 1%).

Permeabilization was stopped by diluting the detergent with 3 vol of

buffer B (1 mM Tris, 1 mM EGTA, 2 mM MgCl2, 10 mM KCl, 5 mM β-

mercaptoethanol, 5 mM ATP; pH 7.4). We then assayed for nuclei by adding

pyrene-labeled rabbit skeletal muscle actin to a final concentration of 1 µM, and

followed the fluorescence increase with a microplate reader (FLUOstaroptima, BMG

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Labtech, Nepean, Ontario, Canada) at excitation and emission wavelengths of 366

and 386 nm, respectively [21, 37].

Cdc42 and Rac2 activation assays. Prior to performing these experiments, a time-

course study was performed to determine the optimal point at which to measure

activation of Rac2 and Cdc42 following exposure to fMLP. Of samples harvested at

15 - 180 s, the maximum signal was observed at 30 s, and therefore, a 30 s time-

point was used for subsequent experiments. To assess the effects of Slit2 on fMLP-

induced activation of Cdc42 and Rac2, pull-down assays were performed as

previously described with slight modifications [39]. The p21-binding domain (PBD;

aa 67-150) of PAK1 in pGEX-4T3 vector was expressed as a GST fusion protein in

BL21 (DE3) E. coli cells. The GST-PBD fusion protein was affinity purified using

glutathione sepharose 4B beads (GE Healthcare Bio-Sciences, Piscataway, NJ).

Protein bound beads were aliquoted and stored at –80°C for later use. Human

neutrophils purified by dextran sedimentation (~1 x 107/sample) were diluted in 0.5

ml 37°C warmed HEPES-HBSS and incubated with purified Slit2 (0.6 µg/ml) at

37°C for 10 min. Cells were stimulated with fMLP (1 µM) for 30 s at 37°C and the

reaction was stopped by adding 0.5 ml ice-cold 2x lysis buffer (1x = 50 mM Tris, pH

7.5, 10% glycerol, 100 mM NaCl, 1% NP-40, 5 mM MgCl2, 1 mM DTT, 1mM PMSF,

1/100 protease inhibitor cocktail, and 1 mM NaVO3). Samples were centrifuged at

maximal speed in a bench-top centrifuge for 5 min at 4°C and an aliquot of

supernatant was used as loading control. The remaining supernatants were added

to GST-PBD glutathione beads (20 µg GST-PBD/sample). Samples were rotated at

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4°C for 1 h, washed 3 times with cold wash buffer (50 mM Tris, pH 7.5, 40 mM

NaCl, 0.5% NP-40, 30 mM MgCl2, 1 mM DTT, 1 mM PMSF, 0.1 mM NaVO3) and 20

µl of 2x Laemmli loading buffer added. Samples were run on SDS-PAGE and

transferred onto a 0.2 mm PVDF (Millipore) membrane. Cdc42 and Rac2 were

detected using anti-human Cdc42 and anti-human Rac2 primary Ab and HRP-

conjugated secondary Ab. Densitometry analysis was performed on the blots using

Image J software. To examine the effects of Slit2 on spatial distribution of activated

Rac and Cdc42, assays were performed as previously described [33]. Briefly,

mouse bone marrow-derived neutrophils were isolated and 1x 106 cells were

suspended in Nucleofector solution supplemented with 6 µg cDNA expression

plasmids encoding each of yellow fluorescent protein-tagged p21-binding

domain of PAK (PAK-PBD-YFP), which selectively detects activated Rac and

Cdc42, together with red fluorescent protein-tagged H-Ras (H-Ras-RFP) to label the

plasma membrane [21, 33]. Cells were transfected using a Cell Line V

NucleofectorTM kit (Amaxa Biosystems, Amaxa, Inc.) and the NucleofectorTM

program Y-001 [21, 33]. Transfected cells were carefully recovered and transferred

to Iscove‟s Modified Dulbecco‟s Medium pre-warmed to 37°C and allowed to

recover for 2 h. Neutrophils were placed on coverslips coated with 1% BSA

mounted in an Attafluor cell chamber (Invitrogen) and exposed to a point source of

fMLP (1 µM) dispensed through a glass micropipette [21, 33]. In some experiments,

neutrophils were pre-incubated with purified Slit2 (4.5 µg/ml) for 10 min. Cells were

maintained on a microscope stage heated to 37°C, and digital images were

acquired every 3-5 s using a Leica DMIRE2 inverted fluorescence microscope

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equipped with a Hamamatsu backthinned EM-CCD camera and spinning disc

confocal scan head [21, 33]. Images were acquired and analyzed using VolocityTM

software. Following chemotactic stimulation with fMLP, the ratio of the fluorescence

intensity of PAKPBD-YFP: H-Ras-RFP was compared at the leading edge of the

cell and the trailing edge of the cell [21, 33]. The normalized mean fluorescence

intensity was calculated for 19 cells from three independent experiments [33].

Mouse peritonitis experiments. To determine the effects of Slit2 on neutrophil

chemotaxis in vivo, we used a mouse model of sodium periodate-induced peritonitis

as previously described [19]. All procedures were carried out in accordance with the

Guide for the Humane Use and Care of Laboratory Animals and were approved by

the Hospital for Sick Children Research Institute Animal Care Committee. Adult

CD1 mice were injected intraperitoneally with Slit2 (100 ng) or control medium, then

1 h later with 1 ml of 5 mM sodium periodate in PBS [15]. After 3 h, mice were

euthanized and the peritoneal exudate collected by lavage with chilled PBS (5

ml/mouse). Infiltrating neutrophils were counted using an electronic cell counter

(Becton Dickinson) and neutrophil influx was confirmed by analyzing cytospun

slides. To determine whether Slit2 administered systemically prevents neutrophil

recruitment, purified Slit2 (1.8 µg in 0.2 ml normal saline) was administered by

intravenous tail-vein injection. One hour later, 1 ml PBS containing sodium

periodate (5 mM), C5a (10 µg), or MIP-2 (2.5 µg) was injected intraperitoneally [40,

41]. After 3 h, mice were euthanized, peritoneal exudate collected, and infiltrating

neutrophils counted as described above. The number of infiltrating

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monocytes/macrophages, T lymphocytes, B lymphocytes, and natural killer cells

was determined by labeling cells with Ab directed to F4/80 (10 µg/ml), CD3 (5

µg/ml), B220 (2 µg/ml), or NK1.1 (2 µg/ml), respectively, and performing flow

cytometry as previously described [27, 28].

Statistical analysis. Analysis of variance (ANOVA) followed by Bonferonni post-hoc

testing was performed using SPSS statistical software to analyze the data from

Transwell experiments. In all other cases, the Student‟s t-test was used. p < 0.05

was considered significant.

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A1.4 Results

1) Primary human and mouse neutrophils express the Slit2 receptor, Robo-1. Robo-

1 mRNA and protein expression were detected in both human and mouse

neutrophils (Figure 1A & B) [19, 21]. Since Robo-1 expression has previously been

demonstrated in primary human lymphocytes, as a positive control, we verified

Robo-1 expression in human leukocytes isolated from whole blood (Figure 1A) [16].

We detected two distinct bands for Robo-1 protein in mouse neutrophils, consistent

with the splice variants previously reported (Figure 1B) [42]. Using

immunofluorescence microscopy and flow cytometry, we detected Robo-1

expression on the surface of human and murine neutrophils (Figure 1C-E).

2) Slit2 inhibits migration of human neutrophils towards fMLP. We studied the

effects of Slit2 on Transwell migration of human neutrophils. As expected, basal

migration was minimal (Figure 2A & E), but increased in the presence of an fMLP

chemotactic gradient (Figure 2B & E; p < 0.001). When no chemotactic gradient

was present, purified Slit2 did not stimulate neutrophil transmigration (Figure 2D).

However, Slit2 prevented neutrophil migration towards fMLP in the lower chamber,

in a dose-dependent fashion (compare Figure 2B & C; Figure 2E, p < 0.001 for

the two highest Slit2 concentrations tested). When fPLC-enriched Slit2 from

conditioned medium of Slit2-expressing HEK-293T cells was tested, very similar

results were obtained (Supplementary Figure 3). In this instance, control medium

from mock-transfected cells had no effect on neutrophil migration, verifying that the

Slit2 preparation did not contain any factors that could inadvertently affect neutrophil

141

migration (Supplementary Figure 3; p < 0.05 vs no fMLP). Together, these data

demonstrate that Slit2 inhibits fMLP-induced migration of primary human neutrophils

in a dose-dependent fashion.

3) Slit2 inhibits migration of human neutrophils towards other chemoattractants. To

determine whether Slit2 inhibits neutrophil migration towards different

chemoattractant signals, we performed Transwell assays in which C5a or IL-8 were

placed in the lower chamber. Slit2 resulted in a four-fold and six-fold decrease in

neutrophil migration towards IL-8 and C5a, respectively (Figure 2F; IL-8: 93 ± 23

cells/field; IL-8 + Slit2: 23 ± 5 cells/field; C5a: 51 ± 10 cells/field; C5a + Slit2: 8 ± 3

cells/field; p < 0.001 for C5a and IL-8). These data demonstrate that Slit2 is a potent

inhibitor of neutrophil migration towards diverse types of chemotactic cue.

4) Slit2 inhibits directional but not random migration of human neutrophils. We next

determined whether the observed effects of Slit2 on neutrophil migration were due

to inhibition of cell chemotaxis or chemokinesis. Chemokinesis is defined as random

movement in response to a stimulant. Unlike chemokinesis, chemotaxis includes a

vectoral assessment of migration and is defined as directional migration in response

to a chemotactic gradient. Therefore, defects in chemokinesis result in the failure of

a cell to move while defects in chemotaxis result in the failure of a cell to move in

the right direction. In the absence of Slit2, neutrophils migrated efficiently towards a

point-source of fMLP (Supplementary Video 1 and Supplementary Figure 4A-C). In

the presence of Slit2, neutrophils moved randomly but failed to move towards the

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micropipette (Supplementary Video 2 and Supplementary Figure 4D-F). These data

suggest that Slit2 does not inhibit generalized movement of neutrophils but rather,

their directionality. To refine the analysis, we tracked the centroid of each neutrophil

over time. Figure 3A depicts the migratory tracks of neutrophils exposed to an fMLP

gradient while Figure 3B represents the migratory tracks of neutrophils exposed to

fMLP in the presence of Slit2. The displacement, speed, and directionality were

determined for each cell. A neutrophil migrating efficiently (directly) up a

chemotactic gradient would have very similar displacement and distance

measurements. As such, its directionality value would be close to 1. Conversely, a

neutrophil moving randomly would have a smaller net displacement despite

traveling the same distance, thereby having a directionality value closer to 0.

Neutrophils incubated with fMLP alone had an average speed of 6.8 ± 0.6 µm/min,

no different from those incubated with fMLP together with Slit2 (6.5 ± 0.6 µm/min;

Figure 3C). In the presence of Slit2, the directionality ratio was significantly reduced

(Figure 3D; fMLP 0.61 ± 0.04; fMLP + Slit2 0.13 ± 0.04; p < 0.002). Taken together,

these data demonstrate that Slit2 does not inhibit the random movement and speed

of neutrophil migration but, rather, prevents directional migration towards a

chemotactic gradient.

5) Slit2 inhibits chemoattractant-stimulated actin free barbed end formation in

human neutrophils. We directly assayed the effects of Slit2 on actin free barbed end

formation, an event critical for formation of protruding lamellipodia and neutrophil

migration [21, 37, 43-46]. In pyrene-actin polymerization curves generated, the

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slope is proportional to the free barbed end numbers [21, 37]. As expected,

unstimulated neutrophils demonstrated low basal levels of free barbed end

generation, but fMLP promoted a rapid, six-fold increase (Figure 4A & B; p < 0.04).

Similarly, when neutrophils were treated with control medium prior to stimulation

with fMLP, we observed a five-fold increase in the rate of actin polymerization as

compared to unstimulated cells (Figure 4A & B; p < 0.01). In the presence of Slit2,

fMLP-induced actin polymerization was considerably more modest, resulting in less

than a three-fold increase compared to unstimulated cells (Figure 4A & B; p < 0.04).

Slit2 significantly reduced fMLP-stimulated generation of actin filaments (Figure 4A

& B; p < 0.05 vs control medium). Accordingly, Slit2 inhibited accumulation of actin

at the leading edge of neutrophils following exposure to fMLP (Figure 4C).

Collectively, these data suggest that Slit2 inhibits directional migration of

neutrophils by disrupting generation of high-affinity free barbed ends that drive actin

filament elongation. This in turn inhibits actin assembly at the leading edge of

migrating cells, thus preventing efficient chemotaxis.

6) Slit2 inhibits chemoattractant-induced polarization and activation of Rac2 and

Cdc42 in primary human neutrophils. Following chemotactic stimulation, activation

of the Rho GTPases, Rac and Cdc42, plays a key role in the re-organization of actin

filaments [19, 21, 34]. Since the predominant isoform of Rac in human neutrophils is

Rac2, not Rac1, we specifically studied activation of Rac2 [47, 48]. We used GST

beads conjugated to the p21-binding domain of p21-activated kinase-1 (PAK-PBD)

to detect the activated, GTP-bound species of Rac and Cdc42 [39]. Unstimulated

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neutrophils had low basal levels of activated Rac2 and Cdc42 (Figure 5A & B).

Exposure to fMLP increased levels of activated Cdc42 by five-fold, and of activated

Rac2 by three-fold (Figure 5A and B; p < 0.01 vs unstimulated for both Cdc42 and

Rac2). Slit2 did not affect basal levels of activated Rac2 and Cdc42, but significantly

inhibited fMLP-induced activation of these GTPases (Figure 5A & B; p < 0.05).

Upon stimulation with fMLP, levels of activated Cdc42 and Rac2 in the presence of

Slit2 were less than half those observed when Slit2 was not present (Figure 5B; p <

0.05). Moreover, Slit2 prevented spatial accumulation of activated Rac and Cdc42

at the leading edge of fMLP-stimulated neutrophils (Figure 5C & D; p < 0.001).

These data demonstrate that Slit2 inhibits neutrophil chemotaxis and actin

polymerization by preventing cell polarization and disrupting generation and

recruitment to the lamellipodium of activated Rac2 and Cdc42.

7) Slit2 does not inhibit chemoattractant-induced activation of other major kinase

pathways. We examined the effects of Slit2 on activation of a number of other

kinase pathways associated with neutrophil chemotaxis, namely, phosphoinositide

3-kinase (PI3K), Akt, Extracellular signal related kinase (Erk), and p38 mitogen-

activated protein kinase (MAPK) [49-52]. As expected, stimulation of neutrophils

with fMLP led to rapid phosphorylation of Akt, Erk and p38-MAPK (Figure 6A-D; p <

0.0005 for Akt; p < 0.05 for Erk; p < 0.05 for p-38 MAPK). Slit2 treatment had no

effect on the basal level of kinase activation (Figure 6A-D). Upon stimulation with

fMLP, resulting levels of activated Akt were comparable in the presence or absence

of Slit2, suggesting that Slit2 does not impair the ability of neutrophils to generate

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PI(3,4,5)P3 (Figure 6A & B). Similarly, Slit2 treatment had no effect on fMLP-induced

phosphorylation of Erk and p38 MAP kinase (Figure 6A, C, and D). Collectively,

these data suggest that Slit2 inhibits neutrophil chemotaxis by specifically

preventing activation of Cdc42 and Rac2, but not activation of Akt, Erk, or p38

MAPKs.

8) Slit2 inhibits leukocyte recruitment in peritoneal inflammation. To study the

effects of Slit2 on neutrophil recruitment in vivo, we used a well-described mouse

model of chemical irritant peritonitis [43]. In the presence of control medium, sodium

periodate administration resulted in influx of 1.90 x106 ± 0.50 x106

neutrophils

(Figure 7A). When Slit2 was pre-administered by intraperitoneal injection, neutrophil

recruitment to the peritoneal cavity decreased six-fold (Figure 7A; 0.30 x106 ±

0.11x106; p < 0.05). When purified Slit2 was pre-administered intravenously by tail

vein injection, neutrophil influx fell from 0.86 x106 ± 0.10 x106

to 0.05 x106 ± 0.02

x106 (Figure 7B; p < 0.001). Although the number of other leukocyte subsets

recruited to the peritoneal cavity was small, Slit2 also inhibited infiltration of several

of them, especially monocytes/macrophages (Supplementary Table 1; p < 0.01).

Slit2 prevented neutrophil recruitment to the peritoneum in response to other

chemoattractant factors, namely C5a and MIP-2 (Figure 7B; C5a: 1.50 x106 ±

0.60x106; C5a + Slit2: 0.30 x106 ± 0.08 x106; p < 0.001; MIP-2: 1.12 x x106

±

0.24x106; MIP-2 + Slit2: 0.65 x 106 ± 0.19 x106, p < 0.01). These data demonstrate

that Slit2 acts as a potent inhibitor of chemotaxis for circulating neutrophils, as well

as for other leukocytes, towards diverse inflammatory stimuli.

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A1.5 Discussion

The aim of this study was to assess the effect of Slit2 on the migration of

circulating neutrophils. We demonstrated that primary human neutrophils express

Robo-1 and that exogenous application of Slit2 blocks migration of neutrophils in

response to a chemotactic gradient. This observation is consistent with the effect of

Slit2 on other cells expressing Robo-1 on their surface. Indeed, Slit2/Robo-1 have

recently been shown to inhibit the migration of a number of different cell types,

including cells of hematopoetic lineage such as dendritic cells and T lymphocytes

[14-16]. A major finding of our study is that Slit2 did not inhibit all movement but

specifically the directed migration of neutrophils. This is a particularly important

distinction because neutrophil chemotaxis to sites of injury is an important

component of inflammatory tissue injury. Indeed, neutrophil-mediated tissue

damage is associated with a number of inflammatory conditions, including

rheumatoid arthritis and ischemia-reperfusion injury [2, 53]. The ability of Slit2 to

specifically disrupt neutrophil chemotaxis points to the potential use of this agent as

a novel therapeutic for inflammatory tissue injury.

While Slit2 has been shown to inhibit chemotactic migration of several cell

types, the mechanisms that mediate these effects remain poorly understood.

Neutrophil migration involves a complex series of events in which the cell, upon

sensing a chemotactic gradient, develops a polarized morphology with a wide

lamella at the front and a narrow tail-like uropod at the back. Critical to the

maintenance of this asymmetry and to forward propulsion is the rapid turnover of

actin filaments at the lamella. In this study, we demonstrated that treatment of

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neutrophils with Slit2 led to a significant reduction in fMLP-stimulated generation of

free barbed ends which are required for rapid actin polymerization at the leading

edge [37]. This observation is consistent with data from neuronal cells linking Robo-

1 to proteins associated with the actin cytoskeleton, including Enabled kinase (Ena)

and slit-robo GTPase activating protein-1 (srGAP1) [9, 10]. However, to the best of

our knowledge, this study provides the first evidence directly linking Slit2 treatment

to a reduction in chemoattractant-stimulated high affinity actin filament ends.

In neutrophils undergoing chemotaxis, the family of small GTPases mediate

turnover of actin. Indeed, treatment of cells with Clostridium difficile toxin, which

inhibits GTPases by monoglucosylation, results in severe defects in actin turnover

and migration [54]. Seminal work describing the effects of introducing dominant-

negative cDNA constructs into HL-60 granolucytic cells identified Rac as the key

determinant of actin assembly, and Cdc42 as being responsible for maintaining the

direction of migration [34]. We observed that exogenous application of Slit2

prevented chemoattractant-induced activation and recruitment of both Cdc42 and

Rac2. These data are consistent with data from neuronal cells where Slit2 treatment

has been shown to recruit the novel GTPase activating protein srGAP1, and to

subsequently inactivate Cdc42 and inhibit axonal migration [10]. In HL-60

neutrophil-like cells, inhibition of Cdc42 using a dominant negative allele prevents

cells from efficiently moving up a chemotactic gradient, and results in extension of

random lamellae in all directions [34].

We found that Slit2 also prevented chemoattractant-induced activation of

Rac2. Similarly, Slit2 has been shown to suppress Rac activation in human vascular

148

smooth muscle cells, human T lymphocytes, and murine RAW 264.7 macrophages

[13, 15, 16]. In murine neutrophils, Rac1 and Rac2 are expressed at similar levels,

and each isoform has distinct functions. Neutrophils deficient in Rac1 display

normal migratory velocity but reduced directionality towards chemotactic gradients

[19]. In contrast, Rac2-deficient neutrophils demonstrate reduced migration speed,

but normal chemotactic migration [19]. Rac1-deficient neutrophils show a partial

reduction in chemoattractant-induced actin polymerization, and the kinetics of actin

assembly are delayed, preferentially inhibiting early rather than later events [43].

Overall, the effects of Slit2 we observed on neutrophil migratory characteristics are

highly reminiscent of Rac1 deficiency. In our experiments, rather than evaluate

overall actin assembly, we focused on a key regulatory feature of this process,

namely, generation of free high-affinity actin filament ends. Measurement of free

barbed end formation specifically measures the initial burst of actin activity following

chemotactic stimulation. Indeed, free barbed end generation of actin is required for

efficient cell chemotaxis. We found that Slit2 inhibited chemoattractant-induced

generation of free barbed ends by over 50%. This falls in between values observed

in Rac1- and Rac2-deficient neutrophils, in which a 30% defect and a 70% defect in

free barbed end generation has been reported, respectively [21]. It is interesting to

note that following chemotactic stimulation of both Slit2-treated human neutrophils

and Rac1-deficient murine neutrophils, random migration of cells remains intact

despite a partial defect in generation of actin high-affinity free barbed ends.

Emerging data supports the concept that it is not the total amount of actin

polymerization that governs cell motility, but rather, the spatiotemporal

149

dynamics of actin assembly within the migrating cell. In support of this notion is the

recent discovery that hematopoietic protein 1 (Hem-1) constitutes part of an

organizational complex that localizes to propagating waves of actin nucleation

within migrating neutrophils [55, 56]. These waves interact reciprocally with actin to

define and organize the leading edge of neutrophils [56]. In this way, net cell

movement results from the collective actions of multiple self-organizing actin-based

waves.

At the molecular and cellular level, Slit2‟s effects on neutrophil migration

share features akin to those seen in both Rac1- and Rac2-null mice. This may be

explained by the differences in expression of Rac isoforms between murine and

human neutrophils. In murine neutrophils, Rac1 and Rac2 are expressed at

equivalent concentrations. In human neutrophils, Rac2 expression is 4 to 40 times

greater than that of Rac1 [47, 48, 57]. Thus, in human neutrophils it is likely that

Rac2 mediates functions assumed by Rac1 in murine neutrophils. In human

neutrophils it has proven very difficult to delineate the individual functions of Rac1

and Rac2. The two GTPases are 92% homologous and the guanine nucleotide

exchange factors that regulate them are the same, rendering expression of mutant

proteins in neutrophil-like cell lines an ineffective means of dissecting the individual

roles played by Rac1 and Rac2 in chemotaxis. Moreover, human neutrophils are

small, terminally differentiated cells which are difficult to transfect, further

complicating the ability to experimentally manipulate them. Together, our data

suggest a mechanism of action whereby Slit2 binding to Robo-1 in human

neutrophils prevents chemoattractant-induced activation of Rac2 and Cdc42, with

150

consequent disruption of actin free barbed end formation, and ultimately, inhibition

of directional neutrophil migration.

Stimulation of neutrophils by fMLP also leads to rapid phospholipid

metabolism and activation of major kinase pathways, including Akt, Erk, and p38-

MAPK, responsible for transcriptional changes. Studies using specific inhibitors

demonstrate that disrupting each of these pathways significantly disrupts neutrophil

chemotaxis. However, exogenous treatment with Slit2 had no effect on the

chemoattractant-induced activation of any of the above pathways. We observed

normal activation of the Akt pathway in response to chemotactic stimulation,

suggesting that Slit2 does not inhibit phospholipid metabolism and specifically,

generation of PI(3,4,5)P3. These results were somewhat surprising, given the

important role played by PI(3,4,5)P3 in chemotactic migration of neutrophils. In one

study, neutrophils from PI3Kγ-deficient mice displayed reduced directional migration

towards chemotactic gradients [50]. Our data is, however, consistent with

observations in human HL-60 granulocytic cells expressing a dominant negative

allele of Cdc42. In these studies, suppression of Cdc42 still led to normal

PI(3,4,5)P3 production and Akt activation [34]. In yet another study, Slit2 prevented

chemokine-induced activation of PI3K in human breast cancer cells [12]. We further

found that in human neutrophils, Slit2 did not inhibit chemoattractant-induced

activation of Erk, nor p38-MAPK. These data are in concordance with those of

others, demonstrating that neither activation of p38-MAPK in Jurkat T lymphocytes

nor activation of Erk in human granulocytic cells was affected by Slit2 [16, 17]. In

another study, Slit2 prevented chemotaxis and chemoinvasion of breast cancer

151

cells towards the chemokine, CXCL12, and inhibited CXCL12-induced activation of

Erk [12]. These differential effects of Slit2 on inducible kinase activity may be

attributable to the different cell types used and to the different chemotactic agents

used to stimulate them.

To determine whether Slit2 can prevent neutrophil recruitment in vivo, we

used mouse models of peritoneal inflammation induced by local instillation of

sodium periodate, C5a, or MIP-2. We found that administration of Slit2, either

intraperitoneally or intravenously, significantly reduced neutrophil recruitment. This

is the first direct demonstration of Slit2‟s potent anti-chemotactic actions on

neutrophils in vivo. These data confirm a universal “antimigratory” role for Slit2, and

are in keeping with recent work showing that Slit2 prevents pathologic

neovascularization within the eye by inhibiting chemotaxis of endothelial cells

towards vascular endothelial growth factor [18]. In another study, Slit2 ameliorated

glomerulonephritis-associated kidney injury by inhibiting chemotactic infiltration of

macrophages [15]. Our results would suggest that localized or systemic delivery of

Slit2 may reduce neutrophil recruitment and subsequent tissue damage associated

with inflammation. Soluble Slit2 is relatively “sticky” and could potentially be locally

maintained at high concentration by adhering to extracelleular matrix proteins such

as glypican-1 [58]. Thus, after regional administration, Slit2 could be retained at

sites of inflammation, such as joints and transplanted organs, thereby alleviating

neutrophil-inflicted tissue injury associated with rheumatoid arthritis and ischemia

reperfusion injury. Because Slit2 blocks migration of several types of inflammatory

cells, including neutrophils, T lymphocytes, macrophages, and dendritic cells,

152

towards diverse chemotactic stimuli, it could act as a highly effective anti-

inflammatory agent [14-17]. Further studies are needed to explore the clinical use of

Slit2, or a Slit-like agent, for prevention and treatment of localized inflammation.

153

A1.6 Acknowledgments

The authors wish to thank Dr. Mohabir Ramjeesingh for technical assistance,

and Drs. Gilles St-Laurent, and Sylvie Perret for reagents. We are grateful to Dr.

Sergio Grinstein for reagents and for helpful advice. This work was supported by the

Canadian Institute of Health Research (L.A.R.), the Kidney Foundation of Canada

(L.A.R.), and an Early Researcher Award from the Ministry of Research and

Innovation, Government of Ontario (L.A.R.). L.A.R. holds a Canada Research Chair,

Tier 2.

154

A1.7 Authorship

S.T. designed and performed experiments, analyzed results, and helped with

manuscript preparation. I.M.M., Y-W.H., M.A.O.M., and M.Y. designed and

performed experiments and analyzed results. M.R.C., G-Y.L., and C.X.S. designed

and performed experiments. Y.D. generated critical reagents and helped with

manuscript preparation. M.G. designed experiments and helped with manuscript

preparation. L.A.R. designed experiments, interpreted results, and prepared the

manuscript.

155

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A1.9 Figure Legends

Figure 1. Primary human and murine neutrophils express Robo-1. A, Primary

human neutrophils were isolated from venous blood of healthy volunteers, RNA was

extracted, and RTPCR was performed using specific primers for Robo-1. For

comparison, total RNA was isolated from human leukocytes from whole blood, and

RT-PCR similarly performed. B, Cell lysates from primary human neutrophils and

bone marrow-derived murine neutrophils were harvested and immunoblotting was

performed using anti-Robo-1 primary Ab and HRP-conjugated secondary Ab. C,

Human neutrophils were plated on fibronectin-coated coverslips and labeled

with anti-Robo-1 Ab followed by Cy3-conjugated secondary Ab. Cells were

examined using a Leica DMIRE2 spinning disc confocal microscope at 100x

magnification. Scale bar is 10 µm. Representative image from one of three separate

experiments. D, To detect cell surface expression of Robo-1, primary human

neutrophils were fixed, incubated with anti-Robo-1 Ab followed by PE-conjugated

secondary Ab or with secondary Ab alone, and analyzed using a FACScalibur flow

cytometer (BD Biosciences) and FlowJo software (Tree Star, Inc., Ashland,

OR). Representative image from one of three similar independent experiments.

Value indicates % of cells with positive labeling. E, Mouse bone marrow-derived

neutrophils were isolated and cell surface Robo-1 labeled as described in D.

Representative image from one of three similar independent experiments. Value

indicates % of cells with positive labeling.

165

Figure 2. Slit2 inhibits migration of human neutrophils towards diverse

chemoattractants. A-D, Primary human neutrophils were incubated with purified

Slit2 (4.5 µg/ml) for 10 min at 37 °C, then migration assays performed across 3 µm

Transwell inserts. The lower chamber contained HBSS or Slit2-containing HBSS in

the presence or absence of fMLP (1 µM). Neutrophils were placed in the upper

chamber and Transwell plates incubated for 1 h at 37 °C. The insert was removed,

and cells which had migrated from the upper to the lower chamber were gently

centrifuged onto coverslips and cell nuclei labeled with DAPI to facilitate

visualization. Representative high-power (63x) images of migrated cells from four

independent experiments were taken using a Leica deconvolution microscope: A,

HBSS. B, HBSS with fMLP. C, Slit2 with fMLP. D, Slit2. E, Transwell assays were

performed as described above, in the presence of the indicated concentrations of

Slit2. Random fields were counted using a Nikon light microscope. Mean number of

cells counted per 63x field ± SEM. *, p < 0.001; n=10. F, Transwell migration assays

were performed as described above. In the lower chamber was placed either C5a (2

µg/ml) or IL-8 (0.13 µg/ml), in the presence or absence of purified Slit2 (4.5 µg/ml).

*, p < 0.001; n = 4.

Figure 3: Slit2 inhibits neutrophil chemotaxis. Primary human neutrophils were

allowed to settle onto fibronectin-coated coverslips. A micropipette containing fMLP

(1 µM) was used to dispense a point-source and gradient of chemoattractant, and

neutrophil migration was monitored using time-lapse video microscopy. The cells

were maintained on the 37 °C-heated stage of a Leica DMIRE2 inverted microscope

166

equipped with a Hamamatsu back-thinned EM-CCD camera and spinning disc

confocal scan head. Digital pictures were acquired every 3 s. In some experiments,

neutrophils were also exposed to anti-myc Ab affinity-purified Slit2 (0.6 µg/ml).

VolocityTM (Improvision) software was used to track the centroid of migrating

neutrophils and thus calculate the total distance, net distance and speed of

migration. Directionality (displacement/distance) was used as a measure of

chemotaxis. A minimum of 8-10 cells for each condition were examined from each

of three separate experiments. Two to 3 cells from each quadrant were randomly

selected prior to initiating tracking. Only cells which started and remained in the field

of view over the entire course of video capture were analyzed. A, Migratory tracks

from one experiment where neutrophils were exposed to fMLP. „X‟ marks the

position of the micropipette. B, Migratory tracks from one experiment where

neutrophils were exposed to fMLP together with Slit2. „X‟ marks the position of the

micropipette. Panel inset depicts an enlarged view of the tracks made by a single

neutrophil. C, Graph depicting the mean migratory speed of neutrophils exposed to

fMLP alone or to fMLP in conjunction with Slit2. Mean values ± SEM for 3 separate

experiments. D, Graph depicting the mean directionality of neutrophils exposed to

fMLP alone or to fMLP together with Slit2. Mean values ± SEM for 3 separate

experiments. *, p < 0.002.

Figure 4: Slit2 inhibits chemoattractant-stimulated formation of actin free

barbed ends in human neutrophils. A, Time series analysis of the fluorescence

increase associated with actin polymerization. Briefly, 1x106 freshly isolated human

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neutrophils were permeabilized for 10 s with 0.2% OG buffer, and the

permeabilization process was stopped by diluting the detergent with 3 vol of buffer

B, as described in „Materials and Methods‟. Cells were stimulated with fMLP (1 µM)

for 120 s in the presence of fPLC-enriched Slit2 (0.6 µg/ml) from conditioned

medium or control medium. Free barbed end generation was assayed by adding

pyrene-labeled rabbit skeletal muscle actin to a final concentration of 1 µM and

following the fluorescence increase using a microplate reader (FLUOstaroptima)

with fluorescence excitation and emission wavelengths of 355 and 405 nm,

respectively. Representative results of four separate experiments are shown. B,

Pyrene-actin incorporation was monitored as in (A) for 150s and the change in

slope of the curve was used as a measure of the rate of actin polymerization. Mean

rate of actin polymerization normalized to the unstimulated control ± SEM. *, p <

0.05; **, p < 0.04; ***, p < 0.01. C, Freshly isolated human and mouse neutrophils

were incubated with Slit2 and plated on fibronectin-coated coverslips in a 6 well

tissue culture plate. Cells were incubated with fMLP (1 µM) for 3 min, then fixed,

permeabilized with 0.1% Triton, and incubated with rhodamine-conjugated

phalloidin for 30 min to visualize actin. Cells were examined using a Leica DMIRE2

spinning disc confocal microscope at 100x magnification.

Figure 5: Slit2 prevents chemoattractant-induced activation and redistribution

of Rac2 and Cdc42. A, Neutrophils were activated with PBS or fMLP (1 µM for 30

s) in the presence or absence of anti-myc Ab affinity-purified Slit2 (0.6 µg/ml), and

cell lysates collected. GST beads conjugated to the p21-binding domain of PAK1

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were used to pull down activated Cdc42 and Rac and immunoblotting was

performed using specific Ab directed against Cdc42 or Rac2. Blots shown are

representative of five independent experiments. B, Mean values ± SEM of

normalized band intensities from five independent experiments (*p < 0.01; ** p <

0.05). C, Neutrophils were isolated from murine bone marrow as described in

„Materials and Methods‟. One million cells were suspended in 100 µl NucleofectorTM

solution (Amaxa, Inc.) supplemented with 6 µg cDNA for PAK-PBD-YFP and H-Ras-

RFP. Cells were transfected using a Cell Line V NucleofectorTM kit and the

NucleofectorTM program Y-001. Transfected cells were carefully recovered with 500

µl Iscove‟s Modified Dulbecco‟s Medium (IMDM) pre-warmed to 37 °C, and

transferred to 1.5 ml pre-warmed IMDM supplemented with 10% FBS in six-well

plates for 2 h. After the recovery period, cells were incubated with purified Slit2 (4.5

µg/ml) for 10 min. Cells were mounted on a 1% BSA-coated coverslip in an Attafluor

cell chamber mounted on the 37 °C heated stage of a Leica DMIRE2 inverted

fluorescence microscope quipped with a Hamamatsu back-thinned EM-CCD

camera and spinning disc confocal scan head. Cells were exposed to a point-

source of chemoattractant using a glass micropipette containing fMLP (1 µM).

Digital pictures were taken every 3 s for 5 min, and images were acquired and

analyzed using Volocity software (Improvision Ltd). Images showing the distribution

of PAK-PBD-YFP, H-Ras-RFP, and the resulting GFP-RFP ratio at the leading edge

compared to the trailing edge of cells exposed to fMLP alone or fMLP in the

presence of Slit2. Arrow indicates the direction of the chemotactic gradient. Images

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are representative of at least 19 cells analyzed from 3 separate experiments. D,

Experiments were performed as described in (C). Mean values ± SEM for the

normalized mean fluorescence intensity (MFI), calculated as the GFP:RFP ratio at

the leading edge compared to the trailing edge of the cell. A minimum of 19 cells

were analyzed from 3 separate experiments. *, p < 0.001.

Figure 6: Slit2 does not inhibit chemoattractant-induced activation of Akt, Erk,

or p38-MAPK. A, Neutrophils were incubated with fMLP and/or Slit2, as described

for Figure 5A. Cell lysates were collected, and immunoblotting was performed using

specific Ab detecting phospho-Akt, phospho-Erk, and phospho-p38 MAPK. Blots

were stripped and re-probed using Ab detecting total Akt, total Erk, and total p38

MAPK, respectively. Blots are representative of 3 independent experiments. B,

Band intensities for phospho-Akt (p-Akt) normalized to total Akt. Mean values ±

SEM for 3 independent experiments (*, p < 0.0005; **, p < 0.05). C, Band

intensities for phospho-Erk (p-Erk) normalized to total Erk. Mean values ± SEM for 3

independent experiments (**, p < 0.05). D, Band intensities for phospho-p38-MAPK

(p-p38) normalized to total p38-MAPK. Mean values ± SEM for 3 independent

experiments. (**, p < 0.05; ***, p < 0.005).

Figure 7: Slit2 inhibits neutrophil chemotaxis in vivo towards diverse

attractant stimuli. A, Adult CD1 mice were injected intraperitoneally with Slit2 (0.1

µg/mouse) or control medium, and 1 h later, with 1 ml of 5 mM sodium periodate

(NaIO4). After 3 h, mice were euthanized and the peritoneal exudates collected by

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lavage with chilled PBS (5 ml/mouse). Infiltrating leukocytes were counted using an

electronic cell counter and the number of neutrophils quantified using Wright-

Giemsa stain. Mean values ± SEM from 5 separate experiments. *, p < 0.05; **, p <

0.01. B, Adult CD1 mice received an intravenous dose of purified Slit2 (1.8 µg in

0.2 ml normal saline) by tail-vein injection. One hour later, mice were given 1 ml of

NaIO4 (5 mM), C5a (10 µg), or MIP-2 (2.5 µg) by intraperitoneal injection. After 3 h,

mice were euthanized and the peritoneal exudates collected by lavage with chilled

PBS (5 ml/mouse). Infiltrating leukocytes were counted using an electronic cell

counter and the number of neutrophils quantified using Wright-Giemsa stain. Mean

values ± SEM from 4 to 6 separate experiments per treatment condition. *, p <

0.001; **, p < 0.01.

Supplementary Figure 1: Recombinant hSlit2 purified by size-exclusion

chromatography and cobalt-affinity chromatography. A-B, Conditioned medium

was harvested from HEK293- hSlit2-myc cells and control HEK-293 cells as

described in „Materials and Methods‟. Using size-exclusion chromatography,

fractionated samples were collected and were run in 8% SDSPAGE. A,

Representative gel for a sample from pooled fractions was silver stained. B,

Representative gel, transferred to a PVDF membrane and immunoblotting

performed using monoclonal anti-myc Ab. C-D, For larger-scale preparation of Slit2,

conditioned medium was harvested from HEK293-EBNA1 cells transfected with

pTT28-Slit2 expression plasmid, as described in „Materials and Methods‟. Slit2

secreted into the medium was purified by immobilized metal-affinity chromatography

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using Fractogel-cobalt columns. Samples were desalted and immunoblotting

performed. Proteins were resolved on reducing NuPAGE 4-12% Bis-Tris gradient

gels, and transferred to nitrocellulose membranes. C, Representative membrane,

stained with Ponceau red solution. D, Representative membrane, probed with

antipolyHis- HRP Ab. For C and D, lanes are marked as follows: 1) harvested

medium 5 days posttransfection; 2) IMAC flow-through; 3) Wash1; 4) Wash 2; 5)

pooled eluted fractions from Fractogel-cobalt column.

Supplementary Figure 2: Measurement and verification of endotoxin levels

and activity present in Slit2 preparations. From each separate Slit2 preparation,

endotoxin levels were measured using ToxinSensor Chromogenic LAL Endotoxin

Assay Kit (GenScript Corp., Piscataway, NJ), according to the manufacturer‟s

specifications. Endotoxin concentrations ranged from 2.5-8.0 EU/ml, corresponding

to 0.2-0.8 ng/ml endotoxin, and yielding final experimental concentrations of 12-40

pg/ml, which are well below those thought to activate leukocytes. To verify this,

endotoxin (40 pg/ml) was added to Transwell assays, and effects on neutrophil

transmigration examined as described in „Materials and Methods‟ and in Figure 2.

n=2.

Supplementary Figure 3: Slit2 inhibits migration of primary human

neutrophils. Primary human neutrophils were incubated with FPLC-enriched Slit2

from conditioned medium (0.6 µg/ml) or with similar fractions from control medium

for 10 min at 37 °C, then migration assays performed across 3 µm Transwell

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inserts. The lower chamber contained HBSS, control medium or Slit2 in the

presence or absence of fMLP (1 µM). Neutrophils were placed in the upper

chamber and Transwell plates incubated for 1 h at 37 °C. The insert was removed,

and cells which had migrated from the upper to the lower chamber were gently

centrifuged onto coverslips, fixed and random fields were counted using a Nikon

light microscope. Representative high-power (63x) images of migrated cells from

four independent experiments were taken using a Leica deconvolution microscope.

Mean number of cells counted per 63x field ± SEM for 4 independent experiments.

(*, p < 0.05).

Supplementary Figure 4: Slit2 inhibits directional migration of human

neutrophils. Primary human neutrophils were allowed to settle onto fibronectin-

coated coverslips. A micropipette containing fMLP (1 µM) was used to dispense a

point-source and gradient of chemoattractant, and neutrophil migration was

monitored using time-lapse video microscopy at 37 °C. A-C, Migration of neutrophils

exposed to a gradient of fMLP over the course of 5 minutes. D-F, Migration of

neutrophils exposed to a gradient of fMLP together with Slit2 (0.6 µg/ml) over 5

minutes. Representative images from one of five separate experiments. For 4

independent experiments. (*, p < 0.05).

Supplementary Video 1. Human neutrophils migrate effectively towards a

point source of fMLP. Glass coverslips were coated with fibronectin, mounted in a

Leiden chamber, and placed on the heated stage of a microscope. A suspension of

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human neutrophils containing 106 cells/ 100 µl was added and allowed to settle for

10 min. To induce chemotaxis, a point-source of fMLP (1 µM) was delivered using a

borosilicate capillary micropipette. The pipette was held stationary and diffusion of

fMLP generated a standing gradient. Images were acquired using MetaMorph

software (Universal Imaging, West Chester, PA) running on a Dell Optiplex DGX

590 computer interfaced with a Photometrics camera via a 12-bit GPIB/IIA board

(National Instruments, Foster City, CA). Image acquisition was started upon the

pipette entering the field and images were obtained every 10 s until completion of

the experiment. Representative video from one of five separate experiments.

Supplementary Video 2. Slit2 inhibits directional migration of human

neutrophils towards a point source of fMLP. Experiments were performed as

described in „Supplementary Video 1‟. Neutrophils were also exposed to anti-myc

Ab affinity-purified Slit2 (0.6 µg/ml) and cell migration was monitored by time-lapse

videomicroscopy. Representative video from one of five separate experiments.

Supplementary Table 1. Leukocyte subsets recovered from peritoneal lavage

fluid following sodium periodate-induced peritonitis. Adult CD1 mice received

an intravenous dose of purified Slit2 (1.8 µg in 0.2 ml normal saline) by tail-vein

injection. One hour later, mice were given 1 ml of NaIO4 (5 mM) by intraperitoneal

injection. After 3 h, mice were euthanized and the peritoneal exudates collected by

lavage with chilled PBS (5 ml/mouse). The total number of cells was counted, and

the numbers of monocytes/macrophages, T lymphocytes, B lymphocytes, and

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natural killer cells determined by labeling with Ab directed to F4/80, CD3, B220, and

NK1.1, respectively, followed by PE-conjugated secondary Ab. Flow cytometry was

performed using a FACScalibur flow cytometer and FlowJo software. Mean values ±

SEM from 4 separate experiments.

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