MOLECULAR MECHANISMS OF SIGNAL TRANSDUCTION

IN NEUTROPHIL CHEMOTAXIS

BY

YUAN HE

DISSERTATION

Submitted in partial fulfillment of the requirements

for the degree of Doctor of Philosophy in Cell and Developmental Biology

in the Graduate College of

the University of Illinois at Urbana-Champaign, 2011

Urbana, Illinois

Doctoral Committee:

Professor Jie Chen, Chair

Assistant Professor Fei Wang, Director of Research

Associate Professor Phillip A Newmark

Assistant Professor William M Brieher

Assistant Professor Yang ‘Kevin’ Xiang

ii

ABSTRACT

Neutrophils play a critical role in host defense against invading pathogens.

Chemotaxis, the directed migration of cells, allows neutrophil to seek out the sites of

inflammation and infection. Neutrophil chemotaxis as well as other type of cell migration

are considered as cycles composed of highly orchestrated steps. Recently the underlying

signaling mechanisms of neutrophil chemotaxis are better understood with the studies in

knockout mice and neutrophil-like cell lines: a number of signaling molecules in

neutrophil chemotaxis have been identified, and a feedback loop-based model of

“frontness” and “backness” pathways has been proposed to explain the establishment of

neutrophil polarity and chemotaxis. However, the signaling mechanisms that control actin

cytoskeleton reorganization and interaction between the cells and the substratum on

which cells migrate are still not fully understood.

In my first research project, we have identified a signaling pathway, mediated by

non-receptor tyrosine kinase Lyn that is essential for localized integrin activation, leading

edge attachment, and persistent migration during neutrophil chemotaxis. This pathway

depends upon Gi protein-mediated activation and leading edge recruitment of Lyn. We

documented the small GTPase Rap1 as a major downstream effector of Lyn to regulate

neutrophil adhesion during chemotaxis. Depletion of Lyn abolished chemoattractant-

induced Rap1 activation at the cell's leading edge. Furthermore, Lyn controls spatial

activation of Rap1 by recruiting the CrkL/C3G protein complex to the leading edge.

Together, these results provide novel mechanistic insights into the poorly understood

signaling network that controls leading edge adhesion during neutrophil chemotaxis.

iii

In my second research project, we have explored the role of mammalian Target of

Rapamycin complex 2 (mTORC2) in neutrophil chemotaxis. Lines of evidence indicate

that mTORC2 is essential for the dynamics of actin cytoskeleton: depletion of mTORC2

impairs the organization of actin cytoskeleton in mammalian cells, and knockout of

mTORC2 inhibit chemotaxis in D. discoideum through PKB mediated signaling pathway.

In our studies, we found that mTORC2 is essential for neutrophil chemotaxis. Depletion

of mTORC2, not mTORC1, abolishes fMLP-induced actin polymerization and

chemotaxis in dHL-60 cells. Pharmacological inhibition of mTOR kinase activity and

AKT phosphorylation fails to affect actin polymerization and chemotaxis in both human

neutrophils and dHL-60 cells. We further demonstrated that mTORC2 is required for

fMLP-induced Rac activation. After chemoattractant stimulation, mTORC2 translocates

to and accumulates at the leading edge of chemotactic cells. Taken together, our findings

indicate that mTORC2 is essential for neutrophil chemotaxis by controlling actin

polymerization-mediated leading edge protrusion during neutrophil chemotaxis.

iv

To my wife, children and parents

v

ACKNOWLEDGEMENTS

First of all, I would like to thank my advisor, Dr. Fei Wang, for his guidance and

support over the past four and half years. I highly appreciate that Dr. Fei Wang gave me

the precious opportunity to continue the long journey of Ph.D. study when I was thinking

about quitting the scientific career path forever. My thanks also go to Fei lab members

who have given me a lot of help. I have really enjoyed working in this friendly and

supportive lab.

Second, I also would like to appreciate my committee members Drs. Jie Chen,

Phillip Newmark, William Brieher and Yang „Kevin‟ Xiang for their support and

insightful suggestions in the road of my Ph.D. study.

In the end, I would like to thank my wife for her unwavering support when the

days were bad for me and for her consideration when I have been a graduate student for

many years. Without her standing behind me, all of this would never happen. I also

would like to thank my parents for their continuous support and encouragement. I would

also like to thank my son and daughter even if they are always making trouble. Every

time when I come back home and see them growing up little by little, I tell myself that I

should keep moving forward.

vi

TABLE OF CONTENTS

CHAPTER 1 BACKGROUND……………………………………………………………………1

1.1 Cell migration………………………………………………………………………………….1

1.2 Neutrophil chemotaxis………………………………………………………………………...7

1.3 Figures………………………………………………………………………………………...28

CHAPTER 2 NON-RECEPTOR TYROSINE KINASE LYN REGULATES

LEADING EDGE ADHESION IN NEUTROPHIL CHEMOTAXIS…………………………..31

2.1 Summary……………………………………………………………………………………...31

2.2 Introduction…………………………………………………………………………………...32

2.3 Materials and methods………………………………………………………………………..34

2.4 Results………………………………………………………………………………………...42

2.5 Discussion…………………………………………………………………………………….55

2.6 Figures……………………………………………………………………………...................60

CHAPTER 3 MAMMALIAN TARGET OF RAPAMYCIN COMPLEX 2 (mTORC2)

REGULATES ACTIN POLYMERIZATION AND NEUTROPHIL CHEMOTAXIS…………76

3.1 Summary……………………………………………………………………………………...76

3.2 Introduction…………………………………………………………………………………...77

3.3 Materials and methods………………………………………………………………………..79

3.4 Results………………………………………………………………………………………...85

3.5 Discussion…………………………………………………………………………………….95

3.6 Figures and table……………………………………………………………………………...99

CHAPTER 4 CONCLUSION……………………………………………………………...........116

REFERENCES………………………………………………………………………….............119

1

CHAPTER 1 BACKGROUND

1.1 Cell migration

Both prokaryotic and eukaryotic cells are able to move the cell body from one

place to another place in certain conditions. This phenomenon is called cell migration.

Why do cells bother to do that? How can cells achieve this body translocation? What are

the counterparts of the wheels, engine and steering in these tiny entities when they decide

to do the business trip? These questions have fascinated biologists for many decades.

With the help from new tools and methodologies in modern biology, we are beginning to

unravel the details of cell migration at the molecular level.

The importance of cell migration

Cell migration plays a pivotal role in normal development and homeostasis of

multicellular organisms, and the abnormities of cell migration contribute to the

progression of many diseases (Lauffenburger and Horwitz, 1996; Ridley et al., 2003).

During the process of early embryonic development, the formation of three germ layers,

including endoderm, ectoderm and mesoderm, results from the collective migration of

cells within the blastocyst. Subsequently, cells migrate toward target locations to form

various tissues and organs. Errors occurring in this process can cause developmental

defects or even early embryonic failure. In adult organisms, cell migration is crucial for

homeostasis, such as mounting an effective immune response and wound healing. Many

diseases, such as vascular disease, chronic inflammatory diseases, and tumor formation

and metastasis, are related to aberrant cell migration. The understanding of cell

2

migration mechanisms will help to develop novel therapeutic strategies to treat cell

migration-related diseases.

Embryonic Development

Cell migration is a robust process that occurs throughout the embryonic

development (Gilbert, 2010). The life of human and other vertebrate animals starts from a

zygote, a fertilized egg. The zygote, which contains the entire genetic blueprint for the

life of a new individual, quickly divides and becomes a blastocyst. During the process of

gastrulation, cells within the blastocyst collectively migrate to form three germ layers-

endoderm, ectoderm and mesoderm. Cells within each layer further migrate to target

locations and develop into different tissues and organs. In the developing brain, for

example, neuronal precursors migrate to distinct layers where they begin to send

projections (axons and dendrites) to find their synaptic partners. The guidance of axons

and dendrites shares many similarities with a migrating cell. The precise guidance to their

final targets is essential for the formation of neuronal networks and development of

cognitive functions.

Homeostasis

Homeostasis, including effective immune response and wound healing, also relies

on cell migration. During the immune response, immune cells, such as neutrophils and

macrophages, migrate out from the circulatory system to the infection sites of

surrounding stromal tissues, where they destroy invading microorganisms. When cuts in

the body occur, a complex biological process called wound healing is activated to repair

3

the damage. During this process, immune cells are quickly recruited to prevent invading

foreigners for entering the wound. Fibroblast cells migrate to the cut sites and coordinate

with other biological activities to repair the wounds (Grinnell, 1994; Martin and

Leibovich, 2005).

Pathology

Increasing number of migration-related proteins has been implicated in the

process of embryonic and fetal development. Defects in these proteins can cause the

failure of early embryonic development, resulting in early loss of pregnancy. Defects in

the migration related proteins which participate in the later stage of embryonic

development can result in defective embryos with disorganized tissues. Although some

defects of migration-related proteins do not affect early fetus, they can cause congenital

abnormalities such as epilepsy, focal neurological deficits and mental retardation.

Migration-related proteins are also critical for normal homeostasis. Alteration in

functions of these proteins contributes to many pathologies. During the immune response,

the continuous recruiting of immune cells can cause chronic inflammation. In asthma, for

example, the constant recruitment and activation of white blood cells in the airways

(lungs) of asthmatics causes tissue damage and results in hyper-reactivity of the airways.

Similarly, in rheumatoid arthritis inflammatory cells are constantly recruited to the joint

tissue and cause constant destruction, which results in compromised limb function and

crippling pain.

4

The migratory cycle

How is cell migration achieved? Cell migration has been considered a cyclic

process, comprised of a set of orchestrated steps: protrusion, adhesion, contraction and

de-adhesion, and translocation (Lauffenburger and Horwitz, 1996; Ridley et al., 2003)

(Figure 1.1). Cell migration is initiated when cells detect external signals (chemotactic

molecules). Cells, such as neutrophils, respond to the chemotactic signals by polarizing

actin and extending a protrusion in the side of membrane encountering the highest

concentration of signal (Zigmond, 1977). The new protrusion in the front is then attached

to the substratum which they migrate on, and stabilized by the de novo adhesion

complexes which are composed of transmembrane receptor integrin. Integrin-mediated

adhesion complexes are not only providing the anchor sites for the cell moving forward

but also initiating and transmitting signals into the inside of cells to regulate cell

migration (Beningo et al., 2001; Burridge and Chrzanowska-Wodnicka, 1996). When

cell polarity is established, signals trigger actin-myosin contraction at the rear and side of

the cell, which provide the essential forces for cell body translocation. The adhesion

complexes attenuate and break up in the rear of the cell, and the tail retracts back from

the rear (Huttenlocher et al., 1997; Lawson and Maxfield, 1995; Worthylake et al., 2001).

A migratory cycle is completed and continuous cycles allow cell to move toward the

sources of external signals.

The modes of cell migration

Based on the cell type and the context in which the cell is migrating, two modes

of cell migration have been proposed: single-cell migration and monolayer cell migration.

5

For single-cell migration, cells move as single entities, such as immune cells. As seen in

monolayer cell migration, cells move in groups, including cells and sheet-like layers. The

monolayer cell migration is often observed during embryogenesis when sheets of cells or

loosely associated clusters migrate. However, the mode of cell migration can be changed

according to the environment. Primary melanoma explants, for example, migrate in

collagen gels as multicellular clusters and switch to a single cell mode of migration when

β1 integrin function was blocked (Hegerfeldt et al., 2002). The mechanism of single cell

migration has been intensively studied over the past decades. Many factors, such as

adhesion strength and the type of substratum, external migratory signals and the

organization of the cellular cytoskeleton, have been shown to affect its properties. More

importantly, based on the nature of interaction between cells and underlying substrate,

mesenchymal migration and amoeboid migration have been proposed for two prototypes

of cell migration with distinct characters.

In slow-moving mesenchymal cells such as fibroblasts, myoblasts, single

endothelial cells or sarcoma cells, the cyclic steps in migration are most apparent. During

movement, mesenchymal cells often adopt a spindle-shaped fibroblast-like morphology

(Grinnell, 1994; Tamariz and Grinnell, 2002). This elongated morphology results from

integrin-mediated focal adhesions and the presence of high traction forces on both cell

poles (Ballestrem et al., 2001; Tamariz and Grinnell, 2002). In this process, focal

adhesions play a central role in motility, by providing strong anchors to the ECM,

allowing the actin-myosin-based contractile system, known as stress fibers, to pull the

cell body and trailing edge forward (Huttenlocher et al., 1995; Lauffenburger and

Horwitz, 1996; Small et al., 1996). Concomitant with integrin and actin localization at

6

substrate-binding sites, mesenchymal cells also recruit surface proteases to digest and

remodel ECM. Focal adhesion formation and turnover occur on the timescale of 10–120

min, resulting in relatively slow migration velocities (0.1–2μm/min) on ECM substrate

(Ballestrem et al., 2001).

In contrast to mesenchymal cells, fast-moving cells like neutrophils use amoeboid

movement, exhibit highly polarized morphology and much more rapid membrane

turnover rates, and seem to glide over the substrate (Haston and Shields, 1985; King et

al., 1980; Zigmond et al., 1981). Amoeboid movement, which mimics features of the

single-cell behavior of the amoeba Dictyostelium discoideum, is arguably the most

primitive and in some ways the most effective form of cell migration. Dictyostelium is an

ellipsoid cell that has fast deformability (within seconds) and translocates via rapidly

alternating responses of morphological expansion of the leading edge and contraction of

the trailing edge (Firtel and Meili, 2000). In higher eukaryotes, amoeboid movement is

often carried out by hematopoietic stem cells, leukocytes and certain tumor cells (Francis

et al., 2002; Friedl et al., 2001; Wang et al., 2002b). These cells use a fast crawling type

of movement that is driven by actin assembly and short-lived interactions with the

substrate. Like Dictyostelium discoideum, leukocytes including neutrophils and T

lymphocytes are highly deformable and move at high velocities (2–30 μm/min) (Friedl et

al., 1998; Smith et al., 2005). In contrast to the mesenchymal migration paradigm, large

focal adhesions seen in mesenchymal cells and stress fibers were not observed in these

cells, which may help cells crawl more rapidly (Friedl, 2004; King et al., 1980).

7

1.2 Neutrophil chemotaxis

Neutrophils, also called polymorphonuclear neutrophils (or PMNs), are the most

abundant type of white blood cells in mammals. In response to an injury, Neutrophils act

as the first line of immune defense and are recruited to fight against invading microbes.

Chemotaxis, the directed cell migration process by which neutrophils move in response

to chemical gradients, plays critical role in this defense process. A number of chemicals,

called chemoattractants, are produced at or proximal to sites of infection and

inflammation and then diffuse into the surrounding tissue. Neutrophils sense these

chemoattractants and move in the direction where their concentration is greatest, thereby

locating the source of the attractants and the associated targets. Neutrophils normally

circulate in the blood stream and, upon activation, adhere to and squeeze through the

vascular endothelium, crawling to the sites of infection and inflammation. There they

phagocytose bacteria and release a number of proteases with antimicrobial activity

(Schiffmann, 1982).

The experimental models of neutrophil chemotaxis

Neutrophils, the fastest moving cells in mammals, are an interesting model system

for studying amoeboid migration. Furthermore, aberrant neutrophil chemotaxis

contributes to many pathological conditions of immune diseases, including rheumatoid

arthritis, ischemia-reperfusion syndrome, acute respiratory distress, and systemic

inflammatory response syndromes. After stimulation by a variety of chemotactic

attractants, Neutrophils undergo complex sequential events: reorganization of actin

cytoskeleton, morphological changes, development of polarity and chemotaxis. The

signaling mechanisms underlying these events start to be unraveled recently with the

8

experimentation in transgenic mice and human neutrophil cell lines. The dissection of

signaling pathways underlying neutrophil chemotaxis would help to develop novel

therapeutic strategies to counteract the progression of diseases caused by aberrant

neutrophil chemotaxis.

However, neutrophils are terminally differentiated cells and can not be genetically

manipulated, which pose a major challenge to study the signaling pathways underlying

chemotaxis. Most studies of signaling pathways in isolated human primary neutreophils

solely rely on pharmacological treatments. Although pharmacological approaches have

the advantage of rapid action on the activity of target enzymes, the specificity and

efficiency of these pharmacological reagents have been always problematic and should

be validated in the vitro experiments (Davies et al., 2000). Another approach is to use

cultured cell lines which can be differentiated into neutrophil-like cells. One of these cell

lines, human promyelocytic leukemia cell (HL-60), has been validated as neutrophil-like

cell after DMSO differentiation and has been used in many reports (Hauert et al., 2002;

Servant et al., 2000b; Srinivasan et al., 2003; Wang et al., 2002a; Xu et al., 2003).

However, this cell line is very difficult to transfect and variation has been observed in

different studies. The third strategy is to use transgenic mice which lack specific

signaling components (Hirsch et al., 2000; Li et al., 2000; Sasaki et al., 2000). In this

case, the bone marrow neutrophils can be isolated and used for cell signaling studies and

chemotaxis analysis. However, functional differences of signaling components in

chemotaxis may exist between human neutrophils and murine neutrophils.

Until recently a lot of progress has been made to understand the signaling

pathways controlling neutrophil chemotaxis. The early studies on signal transduction of

9

neutrophil chemotaxis, which almost exclusively relied on experiments with

pharmacological inhibitors, have proposed a linear model of neutrophil polarity and

chemotaxis to connect chemoattractant receptor to the cytoskeleton: attractant receptors

activate Gi, which then triggers actin polymerization at the leading edge via PI3Ps and

one or more Rho GTPase. Using a culture model system, differentiated HL-60 cells, and

combining findings in human primary neutrophils and knockout mice, Henry Bourne’s

lab at UCSF proposed a model of integrated signaling network to explain the

establishment of neutrophil polarity and chemotaxis. This model is comprised of two

divergent signaling pathways which act downstream of different Gi proteins: frontness

pathways and backness pathways (Xu et al., 2003). In the following sections, I will begin

with the discussion on functions of individual signaling components and then provide the

current model of integrated signaling pathways in neutrophil chemotaxis.

The signaling components of neutrophil chemotaxis

Chemoattractants and receptors

Neutrophil chemotaxis is induced by the binding of chemoattractants to surface

receptors. When injury occurs within the body, chemoattractants are released and

activated neutrophils crawl to the sites of infection or inflammation. Although they have

the similar function to induce chemotaxis, the structure of chemoattractants shares little

similarity. Neutrophil chemoattractants vary from lipid molecules to small short peptides

and polypeptides. Neutrophils have receptors for bacterial peptides to sense the presence

of bacterial infection. During the process of bacterial protein synthesis, a formyl group is

added on the nitrogen of an N-terminal methionyl residue, which becomes a signature for

10

bacterial peptides. The tripeptide N- formyl-met-leu-phe (fMLP) is commonly used for

studies of neutrophil activation and chemotaxis (Marasco et al., 1984). Neutrophil

chemotaxis can also occur in the condition of inflammation without bacterial infection.

When inflammation occurs in the body, mitochondria in the cells of damaged tissues,

which are evolutionarily of bacterial origin in protein synthesis machinery, also release

N-formyl peptide sequences (Carp, 1982). Additional chemoattractants, including the

lipids leukotriene B4 and platelet-activating factor, are also released by tissues and

recruited immune cells at the inflammatory sites (Baggiolini et al., 1994; Funk, 2001).

Furthermore, activated complement system releases the polypeptide chemoattractant C5a.

Although these chemoattractants differ dramatically in structure, their receptors belong to

the family of G protein coupled seven–transmembrane-domain receptors (Murphy, 1994;

Murphy, 1996), GPCRs. These receptors include N-formyl peptide receptor, leukotriene

B4 receptor, C5a receptor, interleukin-8 receptor, and platelet-activating factor receptor,

all of which mediate the migratory function of neutrophils to the sites of inflammation. In

addition to chemotaxis, these chemoattractant receptors also initiate other functions of

neutrophils, including cytokine release, oxidant production by NADPH oxidase complex,

production of lipid inflammatory mediators and degranulation.

Neutrophils respond to stimuli by developing polarized morphology. How

chemoattractant receptors distribute along the membrane in polarized neutrophils is an

interesting question, the answer to which may shed light on the mechanism of spatial

organization of other signal molecules in chemotactic neutrophils. Early studies on

chemoattractant receptors have conflicting results: a few reports indicate that receptors

accumulate at the F-actin enriched region (leading edge) (McKay et al., 1991; Walter and

11

Marasco, 1987); the majority of the reports demonstrate the receptors predominantly

localize in the mid-region of the cell body (Model and Omann, 1996; Schmitt and

Bultmann, 1990; Servant et al., 1999; Sullivan et al., 1984). In live neutrophil-like cells,

Servant et al. failed to detect the accumulation of C5a in the leading edge of migrating

cells by using GFP fused C5a receptor, suggesting that chemotaxis can occur in the

absence of receptor gradient (Servant et al., 1999). Furthermore, uniform distribution of

chemoattractant receptor (e.g., cAMP receptor) along the membrane was also observed in

chemotactic D. discoiddeum (Xiao et al., 1997). Other chemoattractant receptors (e.g.,

CCR5) are found to accumulate in the leading edge of neutrophils (Gomez-Mouton et al.,

2004). Taken together, localization of chemoattractant receptors is not the essential

initiator for the asymmetric accumulation of spatial signals during neutrophil chemotaxis.

Gi proteins

The receptors for fMLP, C5a and IL-8 are coupled to pertussis-toxin sensitive Gi

proteins (Cockcroft, 1992). Chemoattractant stimulation activates Gi proteins by

releasing βγ subunits from Gα binding (Gerard and Gerard, 1994). Inhibition studies

demonstrated that neutrophil chemotaxis depends on Gi protein: cells treated with

pertussis toxin (PTX), which catalyses the ADP-ribosylation of Gαi and thereby uncouple

Gi from GPCR stimulation, show complete loss of actin polymerization and other

leading-edge activities (Molski and Sha'afi, 1987; Spangrude et al., 1985). However,

Gα12/13 may participate in the signaling pathway to the activation of RhoA (Xu et al.,

2003). Chemotaxis does not depend on the Gαi subunit, which instead is required for

terminating Gβγ activities (Neptune et al., 1999); Gβγ appears to mediate most of the

known signaling pathways activated by chemoattractants in leukocytes (Rickert et al.,

12

2000; Suire et al., 2006; Weiner, 2002). inactivation of Gβγ or inhibition of their

downstream effectors impair neutrophil motility in response to chemoattractants; In D.

discoideum, genetic deletion of the only Gβγ renders cells non-chemotactic, further

supporting the importance of Gβγ regulating chemotaxis. The Gβγ downstream effectors

include: phosphatidylinositol 3-kinases (PI3Ks), guanosine triphosphatases of the Rho

family (Rho GTPases), protein kinase C isoforms (PKCs), cytosolic tyrosine kinases,

mitogen-activated protein kinases (MAPKs) and protein phosphatases (Figure 2.2.).

Phosphatidylinositol 3-kinase (PI3Ks)

Activated phosphatidylinositol 3-kinases (PI3Ks) phosphorylate inositol

phospholipids on the 3’-position of the inositol ring (Toker and Cantley, 1997;

Vanhaesebroeck and Waterfield, 1999; Wymann and Pirola, 1998). Three classes of

PI3Ks have been indentified, of which class I isoforms selectively use

phosphatidylinositol 4,5-bisphosphate (PIP2) as substrate to generate

phosphatidylinositol 3,4,5-triphosphate [PI(3,4,5)P3], which is a second messenger in

signaling pathways that control cell adhesion, cytoskeletal reorganization, cell growth

and survival. The PI3K class I is thought to be only class playing roles in chemotaxis for

both neutrophils and Dictyostelium. PI3Kγ belongs to class I and consists of p110γ

catalytic subunit and a 101 KDa regulatory protein. In contrast, other class I family

members including α, β, δ isoforms consist of a 110 kDa catalytic subunit (p110) and a

regulatory subunit of differing molecular weights.

In neutrophils, chemoattractants such as fMLP activate PI3Ks and result in the

production of the signaling lipid PI(3,4,5)P3. PI3Kγ is directly activated in vivo by Gβγ

13

through the binding of Gβγ-sensitive regulatory subunits (Stephens et al., 1994; Stephens

et al., 1997; Suire et al., 2005). Other class I isoforms exist more abundantly than PI3Kγ

and are activated by tyrosine kinase Lyn (Ptasznik et al., 1996; Ptasznik et al., 1995). The

role of each PI3K class I enzyme in production of PI(3,4,5)P3 is controversial (Naccache

et al., 2000; Ptasznik et al., 1996; Vlahos and Matter, 1992): some reports suggest that

Lyn kinase-regulated PI3K predominantly controls the production of PIP3 (as much as

70% ); other reports indicate that majority of PIP3 (90% or more) is produced through

PI3Kγ.

PI3K activation by chemoattractant stimulation can be inhibited by fungal toxin

wortmannin and LY294002. Some studies have shown that PI3K inhibition impairs some

responses of neutrophils, including degranulation and oxidant production but fails to

affect phospholipase C activity and calcium release response (Arcaro and Wymann,

1993; Nijhuis et al., 2002; Vlahos et al., 1995). Inhibition of PI3Ks also attenuates N-

formyl peptide-induced neutrophil polarization and incorporation of F-actin into

cytoskeletal actin (Niggli and Keller, 1997; Nijhuis et al., 2002; Vlahos et al., 1995).

However, the total cellular F-actin content increases normally in these inhibition studies.

The exact roles of PI3Ks in neutrophil chemotaxis are still not fully understood. In some

reports, PI3K inhibitors inhibit neutrophil chemotaxis, where neutrophil chemotaxis is

not altered in other studies (Coffer et al., 1998; Niggli and Keller, 1997; Nijhuis et al.,

2002; Thelen et al., 1995). This discrepancy suggests that PI3K-mediated downstream

pathways are not essential for netrophil chemotaxis, but are required for optimal

chemotaxis. Furthermore, mouse neutrophils with PI3Kγ deficiency show dramatically

reduced production of PI(3,4,5)P3, which indicates PI3Kγ is the major isoform for

14

PI(3,4,5)P3 production in mouse neutrophils (Hirsch et al., 2000; Li et al., 2000; Sasaki et

al., 2000). Interestingly, chemotaxis is significantly but not completely inhibited in these

neutrophils. In conclusion, studies using PI3Ks inhibitors and PI3Kγ deficiency

neutrophils indicate that PI3K is not regulating the initiation of actin polymerization but

may be involved in the optimal organization and orientation of actin cytoskeleton for

chemotaxis.

The major functions of PI3Ks are mediated by their product PI(3,4,5)P3, which

has been implicated in regulating cell adhesion, cytoskeletal reorganization, growth and

survival (Toker and Cantley, 1997). PI(3,4,5)P3 is a membrane molecule and functions

by recruiting proteins containing pleckstrin homology (PH) domains to the membrane,

where these proteins are activated and assemble signaling complexes. In neutrophils,

around 20 PI(3,4,5)P3-binding proteins have been identified by a tartgeted proteomic

screen (Krugmann et al., 2002). Chemoattractants including N-formyl peptides and IL-8

induce transient activation of AKT/PKB (Roberts et al., 1999; Sasaki et al., 2000; Tilton

et al., 1997). This activation of AKT/PKB is inhibited by treatment with PI3K inhibitors,

suggesting that AKT/PKB activation is the downstream event of PI3K signaling pathway.

Furthermore, neutrophils from PI3Kγ null mice fail to activate AKT/PKB after

chemoattractant stimulation (Sasaki et al., 2000). Experimentally the PH domain from

AKT has been fused with GFP (PH-Akt-GFP) and transfected into cells as a marker for

PI3P localization during migration.

A body of research work indicates that PI(3,4,5)P3 plays a role in neutrophil

polarity and chemotaxis. In neutrophils and HL-60 cells, application of exogenous

membrane-permeable PI(3,4,5)P3 induces cell polarization, accumulation of F-actin in

15

lamellipodia, and random motility, which are similar to the responses with uniform

stimulation of chemoattractants (Niggli, 2000; Weiner et al., 2002). Unexpectedly, these

responses are dependent on PI3K activity by showing sensitivity to PI3K inhibitors.

These results suggest that a positive feedback loop exists between PI3K activation and

PI(3,4,5)P3. Transfection studies with PH-Akt-GFP have revealed that PIP3 localizes in

the lamellipodia of migrating dHL-60 cells (Servant et al., 2000b; Wang et al., 2002a;

Weiner et al., 2002). In a polarized HL-60 cell, a much deeper gradient of PI(3,4,5)P3,

compared to the external gradient of chemoattractant to which it responds, suggests that

an internal mechanism exists to amplify the effect of the chemoattractant gradient.

Furthermore, exogenously added PI(3,4,5)P3 induces lamellipodia accumulation of

PI(3,4,5)P3, which is also inhibited by Rho GTPase inhibitors (Weiner et al., 2002).

Disruption of the actin cytoskeleton markedly impairs the amplification of the internal

PI(3,4,5)P3 gradient (Wang et al., 2002a). Taken together, all these results imply the

existence of a positive feedback loop though which PI(3,4,5)P3 regulates PI3K activity

via Rho GTPase and actin cytoskeleton. Perturbation of this feedback loop causes cells to

migrate more slowly and wander more than untreated cells. Thus PI(3,4,5)P3 has been

called the internal cell “compass” that navigates moving cells. The downstream effectors

of PI3K include the GEFs for small GTP-binding proteins, and the MAPK cascade,

which have been implicated in the signaling pathway of neutrophil chemotaxis.

Rho family small GTP-binding proteins

The Rho family members have emerged as the key regulators for actin

cytoskeletal reorganization, actin-myosin contractility and cell adhesion during

neutrophil chemotaxis. Small GTP-binding proteins act as a molecular switch for many

16

signaling pathways by alternating in GDP bound form (inactive) and GTP bound form

(active). The exchange of GTP and GDP at the binding site is regulated by guanine

nucleotide exchange factors (GEFs) and guanine nucleotide dissociation inhibitors

(GDIs). The intrinsic GTPase activity of small GTP-binding proteins can be modulated

by GTPase-activating proteins (GAPs). In neutrophils, many small GTP-binding proteins

have been identified, including Ras, Rac1, Rac2, Cdc42 and Rho. Also the regulators for

small GTP-binding proteins, such as DOCK2, PIX, Vav1/3, Sos, p120-GAP and

RhoGDI, have been documented in neutrophils (Bokoch et al., 1994; Dusi et al., 1996;

Stephens et al., 2008; Zheng et al., 1997). Among the indentified small GTP-binding

proteins, Rac, Cdc42 and Rho have been intensively investigated and implicated in the

regulation of neutrophil chemotaxis.

Rac and Cdc42 have been shown to regulate neutrophil actin polymerization.

Upon N-formyl peptide stimulation, Rac and Cdc42 are translocated to the plasma

membrane, where both become activated by resident GEFs (Philips et al., 1995; Quinn et

al., 1993). The activation of Rac and Cdc42 is dependent on Gi-protein activation (Li et

al., 2000). PI3K inhibitors markedly but not completely inhibit Rac and Cdc42 activation,

suggesting that both PI3K dependent and independent activation of Rac and Cdc42 exist

in neutrophils (Benard et al., 1999). PI3K-independent pathway of Rac activation is

further suggested by the results that normal Rac2 activation was observed in PI3Kγ null

mouse neutrophils (Li et al., 2000).

Cdc42 has been investigated for its role in regulating actin polymerization of

stimulated neutrophils. It has been hypothesized that active Cdc42 regulates actin

polymerization via the actin-nucleating protein Arp2/3. In this hypothesis, Cdc42 can link

17

the activated receptor to the activation of Wiskott-Aldrich syndrome protein (WASP),

which in turn stimulates Arp2/3-mediated actin nucleation reaction (Machesky and Insall,

1998; Machesky et al., 1999). In vitro experiments with neutrophils lysate, activated

Cdc42 induce actin nucleation and F-actin polymerization (Zigmond et al., 1997). Two

mechanisms, creation of free barbed ends and enhancement of actin elongation, have

been proposed for active Cdc42-induced actin polymerization.

Two isoforms of Rac, Rac1 and Rac2, have been identified in neutrophils, of

which Rac2 is more abundant (> 96%) in human neutrophils (Quinn et al., 1993).

Chemotaxis is significantly inhibited in neutrophils from Rac2-deficient mice (Roberts et

al., 1999). Furthermore, neutrophils from a patient with a Rac2 dominant-negative

mutation show dramatically impaired chemotaxis to N-formyl peptide and IL-8

(Ambruso et al., 2000; Williams et al., 2000). Actin polymerization was dramatically

inhibited in Rac2-defiecient neutrophils after chemoattractant stimulation (Roberts et al.,

1999). Rac has been suggested to function downstream of active Cdc42 and regulate

PI(3,4,5)P3 synthesis, by uncapping the barbed end and initiating actin polymerization

(Glogauer et al., 2000). In addition, another downstream effector p21-activated kinases

(PAKs) has been implicated in mediating the effects of activated Rac and Cdc42 in

neutrophil chemotaxis (Knaus and Bokoch, 1998).

In contrast to the role of Rac and Cdc42 in actin polymerization, Rho is important

for uropod formation and release from the substratum. Inhibition of Rho by C3

exoenzyme impairs neutrophil chemotaxis to N-formyl peptide, although the initiation of

actin polymerization is not altered (Ehrengruber et al., 1995b; Niggli, 1999; Stasia et al.,

1991; Yoshinaga-Ohara et al., 2002). Neutrophils with Rho inhibition normally polarize

18

and extend the protrusion, but the tails are stuck on the substratum and the cell bodies are

largely elongated. Several downstream effectors have been identified for activated Rho,

including ROCK and atypical δ PKC. Inhibition of ROCK by Y-27632 caused the similar

defect in neutrophils as Rho inhibition, suggesting that ROCK is the major mediator of

activated Rho modulating myosin II for actin-myosin contractility during neutrophil

chemotaxis (Kawaguchi et al., 2000; Niggli, 1999). Atypical δ PKC, one of the protein

kinase C family members (PKCs), has also been implicated acting downstream of

activated Rho. PKCs include three subfamilies, classical, novel, and atypical, which are

regulated in different fashions. Neutrophils express classical isozymes α, βI, and βII,

novel isozyme δ, and atypical isozyme, δ(Kent et al., 1996; Laudanna et al., 1998). PKC δ

has been shown to be activated and translocated to the plasma membrane after N-formyl

peptide stimulation. Inhibition of this kinase almost abolishes most of neutrophil

responses induced by N-formyl peptide. Rho appears to be the upstream regulator of δ

PKC, suggested by the evidence that Rho inhibition by C3 exoenzyme impairs the

membrane translocation of δ PKC. However, it is interesting to note that Rho inhibition

does not affect actin polymerization (Ehrengruber et al., 1995a). The relationship

between Rho and δ PKC in neutrophil chemotaxis needs to be further investigated.

It is clear that Rac/Cdc42 and Rho regulate neutrophil chemotaxis in different

ways. Rac and Cdc42 control actin polymerization and lamellipodium formation. Rho

modulates the actomyosin contractility machinery and uropod release from the

substratum. These two divergent pathways are linked by the downstream effector of

Rac/Cdc42, PAK, which inactivates MLCK and promotes relaxation of myosin II-based

19

contraction. These two pathways have been suggested to oscillate out of phase such that

the lamellipodium extension is followed by uropod retraction.

MAPKs

The small GTP-binding protein, Ras, is activated in neutrophils upon N-formyl

peptide stimulation by Gi-protein dependent pathways (Knall et al., 1996; Worthen et al.,

1994). The results that Ras can be activated in the presence of inhibitors of tyrosine

kinases, PI3K, PKC, rule out these molecules as acting upstream of Ras activation. In

other cell systems, Ras functions as the initiator of the mitogen-activated protein kinase

(MAPK) cascade, which has been assumed to be the same in neutrophils. Thus, the

interest in Ras-mediated signaling is focused on the investigation of MAPK signaling

pathways in neutrophil chemotaxis. Three modules of MAPKs have been characterized,

including ERK-1/2 module, p38MAPK module, and JNK/SAPK module, which also has

been demonstrated to exist in neutrophils. ERK-1/2 module and p38MAPK module have

been shown to be the major activated modules of MAPK cascade in neutrophils

stimulated by N-formyl peptide (Avdi et al., 1996; Thompson et al., 1993; Worthen et al.,

1994). Inhibition of MEK1/2, the ERK-1/2 module, fails to inhibit neutrophil chemotaxis

to N-formyl peptide and IL-8, suggesting that this pathway is not essential for the

migratory function of neutrophils (Coffer et al., 1998; Zu et al., 1998). In contrast,

inhibitors of p38 MAP kinase impair neutrophil chemotaxis to N-formyl peptide but fail

to inhibit IL-8 response of neutrophils (Knall et al., 1997; Nick et al., 1997; Zu et al.,

1998). One mechanism has been proposed that p38MAP kinase activated by N-formyl

peptide relieves the Hsp27-mediated inhibition of actin polymerization via MK2,

20

although the details of this pathway need to be further delineated (Landry and Huot,

1995; Landry and Huot, 1999).

Tyrosine kinases

Neutrophils express Src family kinases, Syk- and FAK-related tyrosine kinase

PYK2, which can be activated by several cell surface receptors including the

chemoattractant receptors, receptor tyrosine kinases and integrins (Berton and Lowell,

1999; Gaudry et al., 1992; Nijhuis et al., 2002). Studies with inhibitors and mouse

neutrophils lacking Src family kinase or Syk reveal that these kinases are important for

integrin-dependent events in neutroplils, such as cell spreading and adhesion (Berton and

Lowell, 1999; Gaudry et al., 1992; Mocsai et al., 2002). However, inhibition or depletion

of Src or Syk kinases does not affect N-formyl peptide induced neutrophil migration

through transwell filters, suggesting that Src family is not essential for neutrophil motility

(Nijhuis et al., 2002). Interestingly, in under-agarose migration assay, inhibitors of these

kinases attenuate N-formyl peptide-induced migration. Taken together, tyrosine kinases

may be required for optimal migration by regulating the interaction between neutrophils

and the underlying substratum (Yasui et al., 1994).

Serine/threonine protein phosphatase

Human neutrophils express both phosphatases 1 and 2A. Inhibition studies in

neutrophils by using calyculin A and okadaic acid suggest that phosphatase 1 and/ or 2A

are required for shape maintenance, sustained increase of cytoskeletal actin, N-formyl

peptide induced chemotaxis (Kreienbuhl et al., 1992; Yokota et al., 1993). However, due

to the other possible targets of these two inhibitors, these experimental results may be due

21

to off-target effects on neutrophil chemotaxis. More specific approaches, such as

knockdown in cultured cell lines and conditional knockout mice, are needed for

investigating the role of these protein phosphatases in neutrophil chemotaxis.

Integration of the signaling pathways in neutrophil chemotaxis

The compass mechanism

When encountering chemoattractants, neutrophils and neutrophil-like dHL-60

polymerizes actin, and thrust membrane projection toward the source of chemoattractants

or random directions in the uniform concentration of chemoattractants. During the

sensing process of extracellular chemoattractants, a signal is transmitted across the

membrane to the cell inside. Cells respond to the signal by comparing the signal strength

throughout all the membrane, determine which side of the membrane has the highest

stimulation of chemoattractants, and thus develop the protrusion on that side. The

comparing mechanism in neutrophils as well as other cell systems has been called

“compass” (Bourne and Weiner, 2002; Rickert et al., 2000). The chemoattractant gradient

difference across the cell body of neutrophils is only about 2% (Devreotes and Zigmond,

1988). However the internal gradients of signal molecules, such as PI(3,4,5)P3, are much

deeper than outside chemoattractants. Therefore, amplification mechanisms of signaling

molecules must associate with the intracellular compass. Although more than a billion

years of eukaryotic evolution have conserved most components of the compass, it is only

recently that the molecular components and underlying mechanisms of the compass are

beginning to emerge as positive and negative regulatory feedback loop for the

organization of leading edge and trailing edge in chemotactic neutrophils (Xu et al.,

22

2003). A “front” signaling pathway and a “back” signaling pathway have been proposed

to explain neutrophil polarity and migration (Figure 1.3).

The frontness signaling pathway: positive feedback loop of PI(3,4,5)P3, Rac and F-actin

How neutrophils assemle polarity signal cascades after exposure of

chemoattractants has been a challenging puzzle for many years. The development of

neutrophil culture models allow researchers to begin to dissect the complex positive

feedback mechanisms which neutrophils use to establish and reinforce the leading edge

(hence, the frontness response).

The first step toward understanding signals involved in the positive feedback loop

was the demonstration, first in Dictyostelium (Meili et al., 1999; Parent et al., 1998) and

later in the neutrophil-like cell line, HL-60 (Servant et al., 2000b; Wang et al., 2002a),

that phosphatidylinositol-3,4,5-tris-phosphate (PI(3,4,5)P3) and other products of

phosphoinositide-3′-kinases (PI3Ks) accumulate selectively in membranes at the moving

cell's leading edge. These 3′-phosphoinositol lipids (PI3Ps) play an essential role in

organizing the neutrophil's leading edge, as indicated by experiments in which

pharmacologic inhibitors of PI3Ks block attractant-stimulated formation of actin

polymers and pseudopods (Servant et al., 2000b; Wang et al., 2002a), apparently by

abrogating activation of small GTPases of the Rho family, including Rac and Cdc42

(Benard et al., 1999), which promote formation of actin polymers (Benard et al., 1999;

Ridley, 2001; Tapon and Hall, 1997).

lines of evidence have demonstrated that PI(3,4,5)P3 and Rac serve as signals in

a positive feedback loop that organizes the leading edge of differentiated HL-60 cells

23

(Srinivasan et al., 2003; Wang et al., 2002a; Weiner et al., 2002). While PI(3,4,5)P3 are

required for activation of Rac, the reverse is also true: inhibiting Rac activation (with

toxins or a dominant interfering Rac mutant) prevents PI3P accumulation and

morphological polarity (Servant et al., 2000b; Srinivasan et al., 2003; Wang et al.,

2002a). Moreover, addition of exogenous PI(3,4,5)P3 can trigger the positive feedback

loop directly, initiating accumulation of endogenous PI(3,4,5)P3 (Weiner et al., 2002),

morphologic polarity, and motility in the absence of added chemoattractant (Niggli,

2000; Weiner et al., 2002); these effects are completely blocked by PI3K inhibitors and

toxins that inhibit activation of Rho GTPases, indicating that they depend on activation of

both endogenous PI(3,4,5)P3 synthesis and Rac by the exogenous lipid. Finally, Rac

stimulates PI(3,4,5)P3 accumulation, at least in part, via the actin polymers that are

generated in response to activated Rac: either attractant or expression of constitutively

active Rac can trigger persistent PI(3,4,5)P3 accumulation, but both effects are prevented

when actin polymerization is blocked by latrunculin B, which sequesters actin monomers

(Srinivasan et al., 2003; Wang et al., 2002a). Taken together, these observations provide

a mechanistic explanation for the persistence of robust actin, and PI(3,4,5)P3-containing

lamellipodium during neutrophil chemotaxis.

A backness signaling pathway: a model for neutrophil self-organizing polarity

While the frontness pathway provides a mechanistic explanation for the protrusive

behavior of neutrophils, it cannot account for the actin-myosin contractility at the back of

the cells. Studies in the Bourne lab (Xu et al., 2003) suggested that in addition to the

frontness signals, attractant receptors also trigger a backness pathway that contributes to

polarity. In contrast to the frontness pathway, which is triggered by Gi, the backness

24

signaling pathway is initiated by G12 and G13. Its downstream components include Rho,

a Rho-dependent protein kinase (ROCK), and activated myosin, which result in

contraction of actin-myosin complexes parallel to the cell membranes at the trailing edge.

At each step of the pathway the corresponding constitutively activating mutants induced a

global backness response: round cells failed to form actin polymers or translocate PH-

AKT-GFP (the PH domain of protein kinase Akt fused to GFP, used as a spatial probe for

3′-phosphoinositol lipids (Servant et al., 2000b)) in response to fMLP, but instead

accumulated activated myosin around the entire cell periphery. Conversely, inhibition of

these steps impaired cell contractility at the back and caused cells to form a strikingly

long tail. Myosin II-mediated contractility is also required for reducing the sensitivity of

the back of the cells, because abrogating the backness pathway renders the entire cell

sensitive to fMLP stimulation (Xu et al., 2003).

Divergent frontness and backness signals begin to explain fMLP-induced

neutrophil polarity. The two sets of signals now allow us to specify roles for proteins that

mediate neutrophil polarity. While these proteins are well characterized in other cells, the

model makes a novel contribution, by casting actin polymers and actin-myosin

assemblies in symmetry-breaking roles, similar to those observed in fish keratocytes

(Verkhovsky et al., 1999). Polarity is self-organizing because mutual opposition between

these assemblies induces them to separate from each other in the membrane. Mutual

incompatibility and inhibition between frontness and backness also provide a mechanistic

explanation for the ability of neutrophils to polarize in uniform concentrations of

chemoattractant and to perform U-turns rather than simply reverse polarity in response to

changes in the direction of the attractant gradient. The neutrophil follows its sensitive

25

“nose” because positive feedback loops at the front and Rho/myosin-mediated

contraction (inhibition) at the back render the pseudopod much more sensitive to PI3P-

and Rac-activating actions of the attractants (Xu et al., 2003).

Conclusion

The signal transduction underlying neutrophil chemotaxis is revealed as a

complex interrelated network with multiple signaling branches. Neutrophil chemotaxis

can be induced by many chemoattractants, including N-formyl peptide, IL-8, C5a and

platelet-activating factor. Although many signaling pathways have been identified in the

studies with N-formyl peptide stimulation, many of these signaling pathways are also

activated in response to other chemoattractants. In addition, some of the signaling

pathways are triggered by integrin ligation or priming factors but fail to induce neutrophil

chemotaxis.

Redundancy is observed in many aspects of the signaling pathways underlying

neutrophil chemotaxis, which also may contribute to the discrepancy of experimental

results. PI3K activity can be regulated by Gi-protein interaction, tyrosine kinase -

dependent pathway or cross-talk from other signaling pathways; Rac activity is controlled

by PI3K –dependent and –independent pathways, actin polymerization can be initiated by

Rac-dependent and Cdc42-dependent pathways. The activation of either pathway in the

redundant pathways may be dependent on the context of cells and experimental

conditions. These redundant and complex signaling pathways allow neutrophils to adapt

to complicated surrounding environment and migrate toward the site of injury. The nature

of reduntant signaling pathways reflects the evolution of signaling networks that are

26

required to be robust and refract to the perturbation of outside environments. No matter

which chemoattratctant is presented to them, neutrophils must temporally and spatially

coordinate protrusive forces, traction forces, and contractile forces for the cell to move.

The existence of multiple signaling pathways and interconnections among these

pathways poses challenges to fully understand the signaling mechanism of neutrophil

chemotaxis. The signaling model of frontness and backness pathways expands our

understanding on the molecular property of neutrophil polarity and chemotaxis. However,

most of the molecular details for both pathways are still not resolved. The formation of

the leading edge is the hallmark of the neutrophil polarity, of which the actin

polymerization has been thought to be the driving force. The small GTP-binding proteins

such as Rac and Cdc42 have been postulated as the essential regulators for the process of

actin polymerization. However, the upstream molecules as well the downstream

molecules of Rac /Cdc42 activation are still elusive in neutrophil chemotaxis. Also

similar to other signaling arms there might exist other signaling pathways to regulate the

actin polymerization and thus front organization.

Furthermore, as an important feature in the cyclic process of neutrophil

chemotaxis, the interactions between neutrophils and the underlying substrate during

chemotaxis have been largly unexplored, although the importance of cell adhesion in

neutrophil functions has been shown by many researchers. The signaling mechanisms

regulating the adhesion formation at the leading edge and the adhesion disassembly of the

trailing edge are very interesting questions for future studies. When neutrophils encounter

chemoattractants, multiple signaling pathways have been activated, as observed in the

stimulation with fMLP. Which signaling pathways are dedicated to control the adhesion

27

and de-adhesion? How are these signaling pathways integrated to regulate these two

seemingly opposite processes? How are the signaling pathways regulating adhesion and

de-adhesion temporally and spatially integrated to the other pathways regulating actin

polymerization and myosin contraction?

28

1.3 Figures

1

2

4

3

1

2

4

3

Figure 1.1 Migration cycles. Cell migration is an integrated process that requires

the continuous, coordinated protrusion (2) formation (3) and disassembly of

adhesions and retraction of the trailing edge (4).

29

αi

GPCR

βγ

PI3Kγ

Rac/Cdc42Tyr-

kinaseMAPK

phosphatase

fMLP

Neutrophil chemotaxis

Figure 1.2 Overview of signaling molecules activated downstream of GPCRs.

Chemoattractants, such as fMLP, bind to and activate the seven-transmembrane-domain

receptors on the cell surface, which trigger the dissociation of coupled trimeric Gi protein

and release the βγ subunits. Subunits βγ directly or indirectly activate numerous

signaling molecules which play a role in neutrophil chemotaxis

30

“fro

ntn

ess”

“ba

ckn

ess”

Figure 1.3 Frontness and backness signals modulate neutrophil polarity and

sensitivity to attractant. Abbreviations: R, G protein-coupled receptor; Gi and

G12/13, trimeric G proteins; Rac and Rho, small Rho GTPases; PI3P, 3′

phosphoinositides. The more or less symmetrically distributed actin ruffles and

PI3P accumulation seen at early times (e.g., 30 s) after application of a

uniform stimulus presumably mask a fine-textured mosaic of interspersed

backness and frontness signals, some triggering activation of PI3Ks, Rac, and

actin polymerization, others promoting activation of Rho and myosin. Localized

mechanochemical incompatibility of the two cytoskeletal responses, combined

with the ability of each to damp signals that promote the other (dashed inhibitor

lines), then gradually drive them to separate into distinct domains of the

membrane. As a result, within 2 min, a morphologically distinct pseudopod,

which is highly sensitive to attractant, cleanly demarcates itself from relatively

insensitive membrane, enriched with myosin, at the back and sides (Xu et al

2003) .

Actin-myosin

(contraction)

F-actin

(protrusion)

Gi

Gα13

GβγPI3K Rac

Rho

ROCK

Actin/myosin

F-actin

GPCR

Gi

Gβγ

GPCR

Gα13

31

CHAPTER 2 NON-RECEPTOR TYROSINE KINASE LYN REGULATES

LEADING EDGE ADHESION IN NEUTROPHIL CHEMOTAXIS

(Data presented in this chapter are in press at Journal of Cell Science, 2011)

2.1 Summary

Establishing new adhesions at the extended leading edges of motile cells is

essential for stable polarity and persistent motility. Despite recent identification of

signaling pathways that mediate polarity and chemotaxis in neutrophils, little is known

about molecular mechanisms governing cell-extracellular matrix (ECM) adhesion in

these highly polarized and rapidly migrating cells. Here we documented in neutrophils a

signaling pathway essential for localized integrin activation, leading edge attachment and

persistent migration during chemotaxis. This pathway depends upon Gi protein-mediated

activation and leading edge recruitment of Lyn, a non-receptor tyrosine kinase belonging

to the Src kinase family. We identified the small GTPase Rap1 as a major downstream

effector of Lyn to regulate neutrophil adhesion during chemotaxis. Depletion of Lyn in

neutrophil-like HL-60 cells prevented chemoattractant-induced Rap1 activation at the

cell's leading edge, while ectopic expression of Rap1 largely rescued the defects induced

by Lyn depletion. Furthermore, Lyn controls spatial activation of Rap1 by recruiting the

CrkL/C3G protein complex to the leading edge. Together, these results provide novel

mechanistic insights into the poorly understood signaling network that controls leading

edge adhesion during chemotaxis of neutrophils, and possibly other amoeboid cells.

32

2.2 Introduction

Cell migration has been conceptualized as a cycle (Ridley et al., 2003). After a

stimulus, cells polymerize actin at the front to initiate protrusions in the direction of

migration. The protrusions are stabilized by adhering to ECM molecules (adhesion).

Adhesions serve as traction sites as the cell moves forward and are disassembled at the

rear, allowing it to detach and retract. In general, strategies for single-cell migration can

be classified into two patterns: amoeboid movement and mesenchymal migration (Friedl,

2004). Slow-moving mesenchymal cells (e.g., fibroblasts) often adopt a spindle-shaped

morphology and adhere via large integrin-mediated plaque-like structures called focal

adhesions. In contrast, fast-moving cells (e.g., neutrophils) use amoeboid movement

associated with fast deformability and highly polarized morphology, and move much

more rapidly. Unlike the mesenchymal migration paradigm, large focal adhesions seen in

slow moving fibroblasts and stress fibers are not detected in these cells (Friedl, 2004),

suggesting fundamental differences between amoeboid and mesenchymal cell adhesions,

but integrin-mediated adhesion is necessary for persistent cell migration (Ley et al.,

2007).

When exposed to chemoattractants, neutrophils polarize and initiate

polymerization of actin at the leading edge (pseudopod) to provide the protrusive force.

Primary neutrophils and neutrophil-like cells have complex signaling pathways that

control formation of their front and back. A frontness pathway consisting of Rac, the lipid

products of phosphoinositide-3 kinases (PI3Ks) and polymerized actin organizes the

leading edge of neutrophils, while the backness signals, including Rho, p160-ROCK and

33

myosin II, lead to contraction of actin-myosin complexes at the trailing edge, causing it to

de-adhere (Srinivasan et al., 2003; Wang et al., 2002a; Xu et al., 2003).

Despite these advances, little is known about the regulation of cell-ECM substrate

adhesion during chemotaxis. Frontness and backness responses per se do not entirely

account for the persistent polarity and motility of neutrophils when they are exposed to

chemoattractants. Indeed, chemoattractant stimulation of neutrophils in suspension (i.e.,

without cell adhesion to the substrate) only causes transient membrane extensions, in

sharp contrast to cells stimulated on substrates, which polarize and migrate persistently

(Srinivasan et al., 2003; Wang et al., 2002a; Xu et al., 2003). In circulating neutrophils

β2-integrins (CD18) are highly expressed and mediate the interactions of neutrophils with

the endothelial cells. The importance of β2 integrin-mediated adhesion in neutrophil

chemotaxis is highlighted by patients with leukocyte adhesion deficiency (LAD), who

have decreased number of neutrophils accumulated at the sites of infection and

inflammation (Bunting et al., 2002). These findings imply a critical role of cell-substrate

adhesion in regulating neutrophil polarity and motility.

To dissect the molecular program that governs adhesion of the neutrophil’s

leading edge during chemotaxis, we assessed the role of proteins known to be involved in

signaling from chemoattractant receptors and integrins. One such protein is the non –

receptor tyrosine kinase Lyn, which belongs to the Src kinase family. Emerging evidence

suggests that Lyn plays a role in modulating integrin signaling and cell migration in

hematopoietic cells (Baruzzi et al., 2008; Maxwell et al., 2004; O'Laughlin-Bunner et al.,

2001). For instance, neutrophils adhesion to fibrinogen leads to Lyn activation and

redistribution to the cytoskeletal fractions (Yan et al., 1996). Furthermore, Lyn can also

34

be activated by multiple chemoattractants in human neutrophils and is required for

phosphatidylinositol 3,4.5-triphosphate formation (Gaudry et al., 1995; Ptasznik et al.,

1996). Interestingly, Lyn deletion in mouse neutrophils led to enhanced integrin-mediated

cell responses (Pereira and Lowell, 2003), suggesting that Lyn plays a negative role in

integrin signaling events. Despite these findings, how Lyn is involved in the regulation of

chemoattractant-stimulated neutrophil migration has not been well defined. Here we

uncovered a Lyn-dependent mechanism for promoting localized integrin activation and

establishing adhesion sites at the leading edge to allow neutrophils to move forward in

the chemoattractant gradient.

2.3 Materials and methods

Reagents and plasmids

Human fibrinogen, human serum albumin (HSA) and fMLP were from Sigma-

Aldrich. Y-27632, LY 294002 and pertussis toxin were from Calbiochem. Alexa fluor

488 phalloidin, and Alexa fluor 647 phalloidin were from Invitrogen. ON-TARGETplus

SMARTpool siRNAs, which contain four siRNAs specifically targeting human Lyn, non-

specific siRNAs and siGLO were from Dharmacon. Mouse or rabbit anti-Lyn antibodies,

Rabbit anti-CRKL and Rabbit anti-C3G were from Santa Cruz Biotechnology. Rabbit

anti-Src (pY418

) antibodies were from Invitrogen. Rabbit anti-CD11/18 antibody was

from Abcam (ab53009). Mouse anti-human CD11/18 antibody (IB4) (Johansson et al.,

1997) was from Ancell Corporation. Monoclonal antibody m24 for the activation epitope

of human CD11/18 was kindly provided by Dr. Nancy Hogg (London Research Institute,

London, UK) and purchased from Abcam (ab13219). Rabbit anti-phospho [Ser19]

myosin light chain antibody, rabbit anti-Akt antibody, and rabbit anti-phospho [Thr308]-

35

Akt antibody were from Cell Signaling Technology. Goat anti-GAPDH polyclonal

antibody was from GenScript. Other reagents if not mentioned specifically were from

Sigma.

The DNA sequences for Lyn 53/56 were amplified from dHL-60 genomic DNA

by PCR and cloned into pcDNA3 with the EGFP gene at the C-terminus. The dominant

negative mutant of Lyn 56 (K275D) (Corey et al., 1998) and both shRNA-resistant

mutants of wild-type Lyn56 and Lyn 56 (K275D) were generated by site-directed

mutagenesis. Mammalian expression vector encoding pEGFP-C3/Rap1A and pcDNA3-

EGFP-RalGDS were provided by Dr. Mark Philips (New York University, School of

Medicine, New York, NY) (Bivona et al., 2004) and Dr. Philip Stork (The Vollum

Institute, Portland, OR) (Wang et al., 2006), respectively. Two target sequences of Lyn

(shRNA1: forward,

TGTATCAGCGACATGATTAATTCAAGAGATTAATCATGTCGCTGATACTTTTT

TC, reverse,

TCGAGAAAAAAGTATCAGCGACATGATTAATCTCTTGAATTAATCATGTCGCT

GATACA; shRNA2; forward,

TGGTGCTAAGTTCCCTATTATTCAAGAGATAATAGGGAACTTAGCACCTTTTT

TC, reverse,

TCGAGAAAAAAGGTGCTAAGTTCCCTATTATCTCTTGAATAATAGGGAACTT

AGCACCA) were annealed and cloned into pSicoR vectors for Lyn knockdown. For Lyn

knockdown with PLKO.1 vectors, two shRNA constructs from Sigma were used

[SHCLNG-NM_002350: shRNA1 (GCCAAACTCAACACCTTAGAA) and shRNA2

(GGAAGAAGCCAACCTCATGAA)]. Two shRNA constructs targeting human Rap1A

36

and CrkL, respectively, were from Sigma [SHCLNG-NM_002884: shRNA1

(GCCAGCATTCCAGACTTCAAA) and shRNA2 (GCTCTGACAGTTCAGTTTGTT)]

and [SHCLNG-NM_005207: shRNA1 (GTGACTAGGGAGACCATTAAC) and

shRNA2 (TGTACGGACTCTGTATGATTT)]

Cell culture, transient transfection, lentiviral production and infection

Cultivation and differentiation of HL-60 cells were described previously (Wang et

al., 2002a). For transient transfections of dHL-60 cells, the AMAXA nucleofection

system was used. Differentiated HL-60 cells (2 x 107, on day 5-6 after DMSO addition)

were spun down and suspended in nucleofector solution V. DNA (5 µg) or siRNA (3 µg)

was added to the cells, and the cell-DNA mixture was subjected to nucleofection

(program T-19). Nucleofected cells were transferred to 20 ml of complete medium.

Subsequent assays were performed 12–48 hours after transfection.

For lentiviral production, shRNA-containing plasmids were co-transfected with

three packaging plasmids pPL1, pPL2 and pPL/VSVG (Invitrogen) into actively growing

HEK293T cells. After 3 days of transfection, supernatant containing the lentiviral

particles was concentrated by Lenti-X concentrator (Clontech) and added to growing HL-

60 cells in the presence of 10 μg/ml polybrene for 24 hours. Lyn knockdown cells were

enriched by cell sorter (Cytomation Plus, Dako) (for pSicoR constructs) or selected in the

presence of 0.5 μg/ml puromycin for 2 weeks (for PLKO.1 constructs).

Immunofluorescence and live-cell imaging

For immunofluorescence studies, cells were suspended in mHBSS buffer (20 mM

HEPES, pH 7.4, 50 mM sodium chloride, 4 mM potassium chloride, 1 mM magnesium

37

chloride, 10 mM glucose) with 1% HSA, and then plated on fibrinogen-coated cover

glass (250 μg/ml fibrinogen) (Schymeinsky et al., 2005) for 20 minutes, washed briefly

and subsequently stimulated with fMLP for various times. Cells were fixed with 3.7%

paraformaldehyde in CSK buffer (10 mM HEPES, pH 7.4, 138 mM KCl, 3 mM MgCl2,

2mM EGTA, 150 mM sucrose) for 15 minutes, permeablized and blocked in CSK buffer

containing 5% donkey serum and 0.1% saponin (Fluka) for 1 hour, incubated with

primary antibodies O/N at 4°C followed by Alexa fluor 488 or 594 conjugated goat anti-

mouse or anti-rabbit secondary antibody. Primary antibodies were used at dilutions of

1:100 (Lyn), 1:100 (p-Src), 1:200 (p-MRLC), 1:100 (β2-integrin), 1:200 (CrkL) and

1:100 (C3G). Secondary antibodies were used at 2 μg/ml. For the detection of F-actin,

cells were incubated with Alexa fluor 488- conjugated phalloidin for 10 minutes at RT.

For live-cell imaging, cells were plated on human fibrinogen-coated surface for

20 minutes, washed briefly, and subsequently stimulated either with a uniform

concentration of 100 nM fMLP or a point source of 10 µM fMLP from a micropipette

(internal diameter of 1 µm, pulled from a glass capillary), as described (Servant et al.,

2000a). To analyze the effects of inhibiting de novo cell adhesion on chemotaxis, dHL-60

cells were plated on fibrinogen-coated surface for 20 minutes and washed briefly.

Adherent cells were then incubated with different concentrations of IB4 antibody (0.1

μg/mL or 0.5 μg/mL) or with mouse isotype IgG for 10 minutes prior to stimulation with

the fMLP-containing micropipette. DIC images, fluorescent images and combined

DIC/fluorescence images were collected with a Zeiss 40X NA 1.30 Fluar DIC objective

or 63X NA 1.4 Plan Apochromat DIC objective on a Zeiss Axiovert 200M microscope.

All images were collected with a cooled charge-coupled device camera (AxioCam MR3,

38

Zeiss) and processed by the Image J program. Kymograph analysis of leading edge

protrusion was conducted as described previously (Shin et al., 2010a).

Immunoprecipitation and western blotting

Cells (5 x 106) were suspended in 90 μl of mHBSS buffer and stimulated with 10

μl of 1 μM fMLP for various times. fMLP stimulation was terminated by the addition of

400 μl of 1.25 x RIPA buffer (50 mM Tris-HCl, pH 7.5, 1% NP-40, 150 mM NaCl, 1

mM EDTA) containing 1 mM Na3VO4, 1 mM NaF, 1 mM PMSF and a protease inhibitor

cocktail (Sigma). After centrifugation at 14,000 x g for 10 minutes, cell lysates were pre-

cleared with protein A beads and thereafter incubated with 2 µg of anti-Lyn polyclonal

antibody. After 2 h of incubation at 4ºC, protein A beads were added to the cell lysates.

The resulting immunoprecipitates were washed with ice-cold RIPA buffer and analyzed

further with SDS-PAGE.

For western blotting, cells were lysed in RIPA buffer with the aforementioned

protease inhibitors. Protein concentrations of the lysates were determined by using DC

Protein Assay Kit (Bio-Rad). Equivalent amounts of protein (20 μg) were resolved on a

10% disulfide-reduced SDS-PAGE gel, transferred to Trans-Blot Transfer Medium,

blocked with 5% nonfat milk in TBST for 1 hour at room temperature, and incubated

with the primary antibody O/N at 4°C followed by a secondary antibody conjugated to

horseradish peroxidase (HRP; GE Healthcare). The HRP signals were detected by using

enhanced chemiluminescence (SuperSignal West Pico). The western blot was stripped

and re-probed with appropriate antibody according to the manual instruction (Restore

Western Blot, Thermo Scientific).

39

Microfluidic gradient device

The microfluidic gradient device comprises of a polydimethylsiloxane (PDMS)

slab embedded with a Y-shaped fluidic channel and glass substrate as its base. The

microchannels were fabricated using rapid prototyping and soft lithography as described

previously (Herzmark et al., 2007; Jeon et al., 2000; Lin and Butcher, 2006). Inlets and

outlets for the fluids and cells were created in PDMS using a steel punch. The surface of

the PDMS replica and a clean glass coverslip (Fisher Scientific) were treated with air

plasma for 90 seconds in a plasma cleaner (Model PDC-001, Harrick Scientific) and

irreversibly bonded together to complete the device assembly. PTFE tubing (0.022"ID x

0.042"OD, Cole-Parmer) (Cole-Parmer) inserted into the ports were used to deliver fluid

to the microchannels. Bubbles were purged from the microchannels by first injecting

ethanol and then rinsing the channels with PBS. The channel region was then coated with

250 μg/mL fibrinogen for 1 hour at room temperature before use. Cells were washed,

suspended in mHBSS containing 1% HSA and injected in the microfluidic device. The

device was left in the incubator (37°C, 5% CO2) for 20 minutes to allow cells to attach to

the substrate. The floating cells in the channels were then gently washed away with

mHBSS.

To generate soluble gradients, fMLP (500 nM) and mHBSS were infused into the

device from separate inlets. Solutions were driven using a syringe infusion pump (Pump

22, Harvard Apparatus) at a flow rate of 0.03 ml/h to generate a concentration gradient in

the channel by diffusion. Phase contrast images were captured using Axiovision software

on a Zeiss Axiovert 200M microscope.

40

Rac-GTP /RhoA-GTP assay

To determine Rac-GTP or RhoA-GTP level in dHL-60 lysate, we used an

absorbance-based (490 nm) Rac G-LISA kit or RhoA G-LISA (Cytoskeleton, Inc). dHL-

60 cells with or without Lyn depletion were centrifuged for 5 minutes at 1,000 rpm at RT,

suspended in mHBSS (5 × 106 cells in 90 μl per condition), and stimulated or

unstimulated with 10 μl of 1 μM fMLP. The reaction was stopped by adding 100 μL of 2

× lysis buffer (provided with the kit) at 4°C. Subsequent steps were performed as

described in the protocol attached to the kit.

Cell adhesion assay

96-well plates were coated with 100 μg/ml fibrinogen at 4°C O/N and blocked

with 2 mg/ml heat-denatured BSA for 1 hour at 37°C. Plates were washed with PBS once

before seeding cells into wells. Cells were labeled with calcium AM (5 μM) for 30

minutes, washed and suspended in mHBSS. 4 x 105 cells (100 μl) were loaded into each

well. After 30 minutes incubation with buffer or fMLP (100 nM), each well was washed

4 times with pre-warmed mHBSS. The remaining cells were then lysed with 100 µl PBS

containing 0.1% Triton-X 100. The fluorescence intensity of each well was taken by

microplate reader (SpectraMax M2). The percentage of adherent cells was calculated as

the ratio of remaining fluorescence intensity and input fluorescence intensity.

Actin polymerization assay

The procedure for measuring polymerized actin has already been described

(Weiner et al., 2006). Briefly, cells were starved in serum-free medium for 1 hour at 37°C

and stimulated in mHBSS in suspension with 100 nM fMLP at RT. After fixation by

41

3.7% paraformaldehyde for 30 minutes at RT, cells were permeabilized with 0.2% Triton

X-100 and stained with Alexa 647-conjugated phalloidin (1:100) for 30 minutes at 37°C.

Stained cells were suspended in ice-cold PBS, and fluorescence was determined by flow

cytometry with a BD Biosciences LSR II System. The mean fluorescence intensity of the

cell population was determined.

Rap1-GTP assay

To determine the level of Rap1-GTP in dHL-60 cells, we used a Rap1-GTP Pull-

Down and Detection kit from Thermo Scientific. For suspension conditions, dHL-60 cells

were centrifuged (5 minutes,1,000 rpm) at RT, suspended in mHBSS (10 × 106 cells in

90 μl for one condition), and stimulated or unstimulated with 100 nM fMLP in a total

volume of 100 μl. The reaction was stopped by the addition of 0.4 ml ice-old 1.25 × lysis

buffer (25 mM Tris-HCl, pH 7.5, 150 mM NaCl, 5 mM MgCl2, 1% NP-40 and 5%

glycerol). For adhesion conditions, dHL-60 cells were suspended in mHBSS (5 × 106

cells in 1 ml for one condition) and plated on fibrinogen-coated 6-well plates for 20

minutes. Cells were stimulated with 100 nM fMLP for 5 minutes or left unstimulated, and

cells (both adherent and non-adherent) were lysed in 0.5 ml lysis buffer. Subsequent steps

were performed according to the manufacturer’s instructions.

Isolation of primary neutrophils

Primary neutrophils were isolated from venous blood from healthy human donors.

Blood was collected into heparin-containing Vacutainer tubes (BD Biosciences) and

neutrophil isolation procedure was performed within 30 minutes of blood collection using

PMN isolation medium (Matrix). Red blood cell contaminants were removed by Red

42

Blood Cell Lysis buffer (Roche), which produced more than 97% of neutrophil purity.

Neutrophils were suspended in RPMI 1640 medium supplemented with 10% fetal bovine

serum at 37°C until the time of experiments conducted within 8 hours after isolation.

Statistical analysis

Statistically significant differences between two groups were determined using

the heteroscedastic two-tailed Student’s t-Test. The difference with p value less the 0.05

is considered to be significant.

2.4 Results

Lyn expression, activation and localization

To understand how Lyn controls chemotaxis, we determined its expression,

activity and subcellular localization in neutrophils. Lyn was highly expressed in primary

human neutrophils and the neutrophil-like differentiated HL-60 cells (dHL-60), but not in

undifferentiated HL-60 cells or non-hematopoietic HEK293 cells (Figure 2.1A). The

upper and lower bands represent the Lyn A and Lyn B isoform, respectively, as

previously reported (Yi et al., 1991). Differentiated HL-60 cells have been used widely

for studying neutrophil polarity and chemotaxis (Bodin and Welch, 2005; Gomez-

Mouton et al., 2004; Hauert et al., 2002; Nuzzi et al., 2007; Schymeinsky et al., 2006):

when differentiated, they exhibit neutrophil morphologies, polarize in response to

attractants, and migrate in gradients of attractant at rates comparable to primary

neutrophils (Servant et al., 2000a; Wang et al., 2002a; Xu et al., 2003). Unlike primary

neutrophils, they can be continuously cultured and are genetically tractable (Neel et al.,

2009; Servant et al., 2000a; Srinivasan et al., 2003; Weiner et al., 2006; Xu et al., 2003).

43

In this study, we used fibrinogen as the ECM substrate to recapitulate the physiological

microenvironment for neutrophil migration. Fibrinogen is a known ligand for β2-integrin

in neutrophils (Altieri et al., 1990).

We analyzed the activity of Lyn by immunoprecipitating endogenous Lyn from

dHL-60 cell lysates, followed by assessment of the level of activated Lyn using an

antibody specific for phospho-Src proteins. Phosphorylation of tyrosine residues (at 397

for Lyn) in the activation loop of the kinase domain up-regulates enzyme activity of Src

kinase family (Hunter, 1987). Exposure of suspended cells (i.e., in the absence of ECM

attachment) to the bacteria-derived chemoattractant formyl-Met-Leu-Phe (fMLP) induced

rapid and robust activation of Lyn, reaching a maximum at 30 sec (6.1 ± 2.4 fold over

resting condition; mean ± s.e.m), as detected by the phospho-specific antibody (Figure

2.1B). Treatment of cells with pertussis toxin (PTX) completely abolished Lyn activation

(Figure 2.1C). Covalent modification of the α subunit of Gi by PTX inactivates Gi and

abrogates all frontness signals (Xu et al., 2003). In contrast, inhibition of PI3K with a

specific inhibitor LY294002 expectedly prevented fMLP-induced Akt phosphorylation

but failed to affect Lyn activation (Figure 2.1C and data not shown). Thus, fMLP-

stimulated Lyn activation depends upon Gi but not PI3K. Interestingly, when exposed to

fMLP, cells plated on the fibrinogen substrate exhibited a relatively small increase in

phosphorylated Lyn (10-20%), which sustained for over 10 minutes (data not shown).

The distinct patterns of Lyn activation between suspended and adherent cells suggested

differential regulation of Lyn activity under these conditions.

Immunofluorescence of Lyn in non-stimulated adherent dHL-60 cells was

distributed cortically around the plasma membranes, with some cytoplasmic localization,

44

but was preferentially distributed to the leading edge of 73% of cells (out of 415

examined) treated with fMLP, co-localizing with F-actin (Figure 2.2A). The

immunofluorescence was specific for Lyn, as verified by competition experiments with a

blocking peptide. The asymmetry of Lyn in fMLP-treated cells was not caused by

increased amounts of membranes at the front, as shown by dual imaging of Lyn and a

fluorescent membrane dye (DiD) in the same cells. While membranes were uniformly

located and in some cases slightly enriched at the leading edge, there was a significant

increase in Lyn, as shown by the merged image. fMLP-induced leading edge recruitment

of Lyn was also seen in primary neutrophils (Figure 2.2A, lower panel). In addition,

fMLP also stimulated asymmetric accumulation of phospho-Src (activated Src) proteins,

in patterns highly reminiscent of total Lyn (Figure 2.2B). Immunofluorescence of

phospho-Src was mostly attributed to phospho-Lyn protein, because RNAi-mediated Lyn

depletion (the next section) nearly completely abolished the phospho-Src

immunofluorescence. Thus, Lyn is activated and translocates to the leading edge in

response to fMLP.

Depletion of Lyn impairs neutrophil chemotaxis

We next assessed the effect of Lyn depletion on dHL-60 chemotaxis (Figure 2.3

and Figure 2.4). We transiently knocked down the expression of genes of interest in dHL-

60 cells by using the Amaxa Nucleofection System, allowing us to quickly test genes

involved in the regulation of neutrophil chemotaxis. Transfection of cells with a pool of

siRNAs targeting human Lyn reduced Lyn protein by ~80% after 48 hours (Figure

2.3A,B). In addition, we used a lentivirus-based system to stably express small hairpin

RNAs (shRNAs) that efficiently depleted Lyn in dHL-60 cells, enabling us to conduct

45

large-scale biochemical analyses. When exposed to an external fMLP gradient, delivered

by a micropipette, cells transfected with non-specific siRNAs formed well-defined

leading and trailing edges and migrated rapidly and persistently towards the tip of the

micropipette (Figure 2.3C). In contrast, although Lyn-depleted cells were capable of

responding to the fMLP gradient and establishing a leading edge in the direction of the

pipette, they protruded significantly slower than control cells. Kymograph analysis

showed that the protrusion speeds for control and Lyn-depleted cells were 6.32 ± 0.44

μm/min and 3.64 ± 0.31 μm/min (mean ± s.e.m.), respectively (Figure 2.3D,E).

Concomitant to decreased protrusion, the speed for trailing edge retraction was also

reduced in Lyn-depleted cells [5.03 ± 0.25 μm/min for control vs. 1.61 ± 0.23 μm/min for

Lyn depletion (mean ± s.e.m.)]. In addition, the bodies of Lyn-depleted cells were unable

to detach effectively from the substrate and remained largely stationary, resulting in a

stretched spindle-shaped morphology (Figure 2.3C) and long trailing edges [32.2 ± 1.3

μm vs. 19.8 ± 1.6 μm for controls (mean ± s.e.m.)]. In keeping with this result, the

elongation index (the ratio of the length of a polarized cell when fully extended to the

width of its pseudopod) of Lyn-depleted cells was greater than that of control cells [2.63

± 0.23 vs. 1.61 ± 0.11, respectively (mean ± s.e.m.)]. At late stage of stimulation (> 6

minutes), some of the Lyn-depleted cells (~26%) retracted their leading edges (Figure

2.3C; cells b and d).

Interestingly, treatment of dHL-60 cells with PP2, a highly specific inhibitor of

Src kinase family, failed to mimic the phenotypes of Lyn depletion. In the micropipette

assay, PP2 treatment caused cells to slightly increase the protrusion and migration speeds.

A similar result was reported: treatment of mouse and human neutrophils with PP2 did

46

not alter neutrophil migration in experiments with the transwell assay (Fumagalli et al.,

2007). The differential effects between Lyn depletion and PP2 treatment may be due to

potential complex interplay among Src kinase family. Expression of a number of Src

family kinases was documented in neutrophils (Korade-Mirnics and Corey, 2000; Lowell,

2004)

The chemotactic behaviors of Lyn-depleted cells were also examined with a

microfluidic gradient device, which assesses migration of cell populations in highly

stable gradients over a long distance (Figure 2.4). Time-lapse video-microscopy could

track individual cells as they migrated in the microfluidic chamber (Figure 2.4A). In this

assay, Lyn depletion markedly impaired neutrophil chemotaxis in an fMLP gradient (1.4

nM/μm). In contrast to untreated cells (Figure 2.4A,B, upper panel; Figure 2.4D), Lyn-

depleted cells protruded slower and exhibited an elongated morphology (Figure 2.4A,B,

lower panel; Figure 2.4D). These defects were similar to those in the micropipette assay.

Notably, the chemotactic index [the ratio of migration in the correct direction to the

actual length of the migration path (Xu et al., 2005)] was similar between control and

Lyn-depleted cells (0.93 and 0.91, respectively), suggesting that Lyn depletion did not

impair directionality of the cells (Figure 2.4C).

The chemotactic defects in Lyn-depleted cells were likely attributed to the lack of

Lyn kinase activity, Expression of a GFP-tagged dominant negative kinase-dead Lyn

mutant (LynK275D) in dHL-60 cells led to defects similar to Lyn depletion. Furthermore,

transfection of wild-type Lyn, but not the kinase-dead mutant, nearly rescued the defects

of Lyn depletion (protrusion speed: 6.08 ± 0.31 μm/min for control; 5.36 ± 0.21 μm/min

for Lyn-depleted cells with Lyn-EGFP rescue) (Figure 2.5A,B,C). Together, these results

47

suggest that Lyn kinase activity might be responsible for Lyn-mediated functions in

neutrophils during chemotaxis.

Lyn is not required for the frontness signals

We next assessed the mechanism by which Lyn controls neutrophil chemotaxis.

Lyn’s asymmetric localization and the effect of Lyn depletion on cell protrusion led us to

investigate a potential role in leading edge establishment. It was reported previously that

actin polymers, phospatidylinositol-3,4,5-phosphate (PIP3), and Rac-GTP are key

components of the “frontness” pathway that controls leading edge establishment during

neutrophil chemotaxis (Srinivasan et al., 2003; Xu et al., 2003). We therefore assessed

whether Lyn depletion reduced these signals. We took advantage of a unique feature of

neutrophils: their ability to polarize when stimulated with chemoattractants, even in

suspension. When exposed to fMLP, neutrophils in suspension quickly (within 1–2

minutes) establish a morphological leading edge (Zhelev et al., 2004). The leading edge

does not persist and retracts after stimulation (Fechheimer and Zigmond, 1983; Sklar et

al., 1985; Wang et al., 2002a; Ydrenius et al., 1997). Attractant stimulation of suspended

neutrophils also suffices to accumulation of polymerized actin, PIP3 and Rac-GTP, albeit

transiently, consistent with the morphological response (Srinivasan et al., 2003; Wang et

al., 2002a; Xu et al., 2003). This feature of neutrophils enables us to assess the frontness

response without input introduced by the outside-in signals from cell-ECM adhesion.

Under these conditions Lyn depletion failed to prevent the accumulation of polymerized

actin, Rac-GTP and phospho-Akt (activated Akt, readout for PIP3) in neutrophils

exposed to fMLP stimulation (Figure 2.6A,B,C), suggesting that signaling pathways

leading to PI3K, Rac and F-actin activation/formation are not altered by Lyn depletion.

48

Lyn is required for localized activation of β2-integrin and leading edge adhesion

During migration protrusion and adhesion of the leading edge are coupled (Ridley

et al., 2003). Therefore, the defects of Lyn depletion on neutrophil chemotaxis could be

due to inability of these cells to attach their protrusive pseudopods, resulting in slower

protrusion. This notion is supported by the observation that at late stage of fMLP

stimulation some Lyn-depleted neutrophils detached and retracted their leading edges,

implying that the cells failed to stably adhere to the ECM substrate at the front. Thus, Lyn

might regulate leading edge adhesion, leading us to assess whether Lyn controlled β2-

integrin in neutrophils. In control cells, β2-integrin immunofluorescence was mostly

distributed at the leading edge after fMLP stimulation, with some diffused localization in

the cell body. The asymmetric distribution of β2-integrin was confirmed quantitatively by

using the ratio of immunofluorescence intensity of β2-integrin between the leading edge

and the cell body (2.54 ± 0.31; mean ± s.e.m.) (Figure 2.7A). The use of ratio of mean

fluorescence intensity discounts variations in the levels of fluorescence among cells and

is thus more appropriate for assessing relative accumulation of the fluorescent signals.

Lyn depletion did not change the asymmetry of β2-integrin localization (2.64 ± 0.19)

(Figure 2.7A). In contrast, although activated β2-integrin, assessed by using antibody

m24, exhibited a similar leading edge enrichment in control cells (Ratio: 2.52 ± 0.27),

Lyn depletion markedly impaired this enrichment (1.35 ± 0.03) (Figure 2.7B,C).

To investigate whether attenuated β2-integrin activity might be responsible for the

defects of Lyn depletion, we assessed the effect of a β2-integrin function blocking

antibody (IB4) on neutrophil migration. We plated dHL-60 cells on fibrinogen and then

treated them with different doses of IB4 for a short term (10 minutes) prior to fMLP

49

stimulation. This procedure selectively prevented fMLP-induced de novo adhesion

without significantly altering global adhesion of cells to the fibrinogen substrates

(Materials and Methods). In the presence of non-specific IgG, dHL-60 cells migrated

towards the source of fMLP and maintained a highly polarized morphology with well-

developed pseudopods (Figure 2.7D, upper panel). In contrast, cells treated with the

intermediate dose of IB4 (0.1 μg/mL) migrated with slower-protruding leading edges

[3.12 ± 0.24 μm/min vs. 5.84±0.38 μm/min for IgG-treated cells (mean ± s.e.m.; n=33

cells for each condition)] and exhibited elongated morphologies (Figure 2.7D, middle

panel). Furthermore, at a higher dose (0.5 μg/ml), IB4 treatment caused some cells

(~28%) to only transiently polarize in response to fMLP, with much smaller leading

edges and a round-up morphology, most likely due to stronger inhibition of leading edge

adhesion (Figure 2.7D, lower panel). Thus, blocking cell adhesion with intermediate dose

of IB4 mirrored Lyn depletion (Figure 2.3C).

It was reported previously that deletion of Lyn causes mouse neutrophils to

become hyper-adhesive when they are adherent to surfaces coated with ECM proteins

(Pereira and Lowell, 2003). Similarly, Lyn-depleted dHL-60 cells exhibited increased

adhesion to fibrinogen in the absence of fMLP (Figure 2.8A). However, in contrast to

control cells, which markedly elevated adhesion after fMLP stimulation, Lyn-depleted

cells only showed a moderate increase (Figure 2.8A). There was no difference in β2-

integrin surface expression between control and Lyn-depleted cells (Figure 2.8B),

consistent with mouse neutrophils (Pereira and Lowell, 2003). To ask whether elevated

adhesion may be responsible for the chemotactic defects of Lyn-depleted cells, we treated

cells with Mn2+

, which triggers β2-integrin activation (Altieri, 1991), causing neutrophils

50

to tightly adhere to the fibrinogen substrate and fail to polarize morphologically in

response to a fMLP gradient (Figure 2.8C). Immunofluorescence studies showed that

while the level of activated β2-integrin was high after Mn2+

treatment, F-actin was largely

diffusely distributed (Figure 2.8D).

Notably, the defects of Lyn depletion are reminiscent of those induced by

inhibiting the backness signals, which caused neutrophils to form highly stretched long

tails (Xu et al., 2003). However, we showed that Lyn was not involved in the regulation

of the backness pathway or myosin II-mediated contractions (Figure 2.9A,B,C). Lyn

depletion failed to alter RhoA activity or the level of activated myosin II (phosphorylated

specifically at Ser19, p[19]-MLC; S3E,F,G). In contrast, treatment of cells with the p160-

ROCK inhibitor Y-26732 markedly reduced the level of phospho-myosin II.

Furthermore, in contrast to Lyn depletion, Y-27632 treatment, which prevented trailing

edge retraction, did not alter the speed for leading edge protrusion (5.91±0.37 μm/min)

(Figure 2.3D). This result is consistent with earlier findings (Shin et al., 2010b), again

demonstrating differential effects between Lyn depletion and inhibition of the backness

signals.

Rap1 is a major effector of Lyn in neutrophils during chemotaxis

The Ras-like small GTPase Rap1 has emerged as a major mediator of inside-out

signaling to integrins (Bos, 2005; Caron et al., 2000; Shimonaka et al., 2003). Rap1

controls chemokine-induced integrin activation and adhesion (Shimonaka et al., 2003),

leading us to assess whether Rap1 acted as a downstream target of Lyn in neutrophils

during chemotaxis.

51

We assayed Rap1-GTP (the activated form of Rap1) by using a previously

described pull-down method (Zwartkruis et al., 1998). Rap1-GTP levels increased

transiently in suspended dHL-60 cells after exposure to fMLP, reaching a maximum at 30

sec (>10 fold over unstimulated control; Figure 2.10A). fMLP stimulation also caused

adherent cells to markedly increase the levels of Rap1-GTP, which sustained after

stimulation. A similar activation pattern was reported in primary neutrophils (M'Rabet et

al., 1998). Rap1 activation was also observed in live cells responding to a gradient of

fMLP, which induced GFP-RalGDS translocation from the cytosol to plasma membranes

at the ruffling leading edge (Figure 2.10C,D, upper panel). GFP-RalGDS is a fluorescent

probe for Rap1-GTP (Wang et al., 2006). A quantitative analysis of the fluorescence

intensity across the cells revealed a steep gradient of GFP signals in untreated control

cells (78±21% decrease). In contrast, GFP-tagged Rap1 was largely uniformly

distributed in the cytoplasm in unstimulated cells and remained diffusely distributed after

fMLP stimulation (Figure 2.11D), suggesting that the asymmetric accumulation was

specific for activated Rap1.

Lyn depletion markedly inhibited fMLP-induced activation of Rap1. We found

that Lyn depletion reduced fMLP-induced Rap1 activation by 63±12% in suspended

cells (at 30 sec) (Figure 2.10A,B) and 41±9% in adherent cells. Interestingly, in

adherent cells without fMLP stimulation Lyn depletion increased the level of activated

Rap1, in keeping with increased cell adhesion and suggesting that Lyn might inhibit Rap1

activity under this condition . In addition, Lyn-depleted cells exhibited poor leading edge

recruitment of GFP-RalGDS, as reflected by much more diffused localization and smaller

GFP gradients across the cell (36±15 % decrease) (Figure 2.10C,D, lower panel).

52

Moreover, when compared with control cells, the ratio of mean fluorescence intensity of

GFP-RalGDS between the leading edge and the trailing edge was markedly reduced by

Lyn depletion (1.40 ± 0.25 for control vs. 1.17 ± 0.11 for Lyn depletion, respectively;

mean ± s.e.m.), again revealing that asymmetric localization of activated Rap1 was

impaired. Thus, Lyn is required for chemoattractant-induced activation and asymmetric

accumulation of Rap1-GTP at the neutrophil's leading edge.

In addition, Rap1 depletion induced chemotactic defects highly reminiscent of

those caused by Lyn depletion. Two shRNA constructs effectively reduced the level of

Rap1A (Figure 2.11A), the predominant isoform of Rap1 in neutrophils (Quilliam et al.,

1991). In accordance to previous reports (Duchniewicz et al., 2006; Li et al., 2007), Rap1

depletion decreased the number of adherent dHL-60 cells plated on fibrinogen-coated

substrate. When stimulated with a point source of fMLP, Rap1A-depleted cells protruded

much slower than cells containing non-targeting shRNA [3.43 ± 0.35 μm/min for Rap1A

shRNA1 and 2.96 ± 0.13 μm/min for Rap1A shRNA2 vs. 5.85 ± 0.20 μm/min for non-

targeting shRNA (mean ± s.e.m.; n=36 cells for each condition)]. Furthermore, like Lyn-

depleted cells, Rap1A-depleted cells retracted significantly slower (0.83 ± 0.12 μm/min

vs 4.66 ± 0.45 μm/min for control cells) and developed a spindle-shaped morphology

(Figure 2.11B,C). The elongation index for Rap1A depleted cells was 2.5±0.32 (mean ±

s.e.m.; n=23), close to the value of Lyn-depleted cells.

Together, these results suggested that Rap1 acts as a major downstream effector

of Lyn in neutrophils during chemotaxis. This inference was further confirmed by effects

of overexpression of GFP-Rap1A in Lyn-depleted cells. Ectopic expression of Rap1

nearly completely rescued the defects caused by Lyn depletion (Figure 2.11D). In GFP-

53

Rap1 overexpressing cells, the average speed for leading edge protrusion was increased

to 5.83±0.48 μm/min, and the elongation index was reduced to 1.74 ± 0.21 μm/min

(mean ± s.e.m.; n=15), even when Lyn was nearly entirely depleted. These values

approached those of the control cells without Lyn depletion.

Lyn recruits the CrkL/C3G complex to the leading edge

Previous studies demonstrated that the Crk family adaptors function downstream

of activated receptor tyrosine kinases, bind and regulate C3G, a guanine nucleotide

exchange factor (GEF) for Rap1, and consequently controls Rap1 activation (Arai et al.,

1999; Sattler and Salgia, 1998). We found that human primary neutrophils and dHL-60

cells express in high abundance Crk-like protein (CrkL), a member of the Crk family.

Interestingly, Crk-I, expressed in many cell types, was nearly undetectable. We thus

focused on CrkL and asked whether it was involved in the regulation of Rap1 by Lyn.

The tyrosine residue Y207, located in the small region between two SH3 domains of

CrkL, is critical for the regulation of CrkL activity: phosphorylation of Y207 promotes

intramolecular association of CrkL, leading to inhibition (de Jong et al., 1997; Rosen et

al., 1995; Senechal et al., 1998). Using a specific antibody, we found that level of

phosphorylation at Y207 (p-Y207) remained largely unaltered in control cells after fMLP

stimulation (Figure 2.12A). In contrast, the level of p-Y207 was significantly higher in

Lyn-depleted cells with and without fMLP stimulation (1.5-2.3 fold over control cells;

Figure 2.12A). Thus, Lyn modulates the removal of the phosphate group from Y207,

thereby relieving CrkL inhibition in neutrophils.

54

Lyn is also involved in the regulation of the subcellular localization of CrkL.

Immunofluorescence of CrkL was distributed diffusely in the cytosol and in the cortical

areas of the control neutrophils in the absence of fMLP and translocated and accumulated

at the leading edge, colocalizing with F-actin (Figure 2.12B). C3G exhibited a very

similar pattern (Figure 2.12C). In contrast, Lyn depletion impaired leading edge

recruitment of CrkL and C3G (Figure 2.12B,C). Both proteins were located more

diffusively throughout the cell bodies, even though the cells were morphologically

polarized. The ratios of mean fluorescence intensity between the leading edge and the

trailing edge were reduced from 3.41 ± 0.15 to 1.6 ± 0.34 for CrkL and from 2.64 ± 0.13

to 1.14 ± 0.09 for C3G (mean ± s.e.m.), respectively.

Consistent with earlier reports, we found that CrkL co-immunoprecipitated with

C3G in neutrophils. This interaction was detected in resting dHL-60 cells and was

unaltered after fMLP stimulation. Furthermore, the interaction between CrkL and C3G

was also detected in Lyn-depleted cells. Thus, Lyn appears unnecessary for the formation

or the stability of the CrkL/C3G protein complex but is instead essential for its spatial

organization.

CrkL is required for Rap1 activation and neutrophil chemotaxis

We next asked whether CrkL acted upstream of Rap1 in neutrophils during

chemotaxis. We found that CrkL depletion in dHL-60 cells caused Rap1 activity in

fMLP-stimulated cells to reduce by 41% (Figure 2.12D). CrkL depletion also caused

severe chemotactic defects (Figure 2.12E). First, CrkL-depleted cells protruded

significantly slower (3.01 ± 0.18 μm/min for CrkL shRNA1 and 2.57 ± 0.29 μm/min for

55

CrkL shRNA2 vs. 6.03 ± 0.36 μm/min for cells containing non-targeting shRNAs; mean

± s.e.m.). Furthermore, 68% (17 of 25 cells examined) developed a stretched elongated

morphology when responding a point source of fMLP, highly reminiscent of the defects

of Lyn- and Rap1-depleted cells. In addition, 20% of CrkL-depleted cells failed to

develop a stable leading edge, similar to cells challenged with high concentrations of IB4

(Figure 2.7D, Lower panel). Together, these results indicate that Rap1 is a major effector

of CrkL in neutrophils during chemotaxis.

2.5 Discussion

Cell migration is a tightly regulated process that requires adhesion of the extended

leading edge to the ECM substrate to ensure uninterrupted migration. Although the

understanding of cell adhesion in slow-migrating mesenchymal cells has been

significantly improved in the past two decades, this key step remains poorly defined in

highly polarized and rapidly moving amoeboid cells including neutrophils. Recent studies

with human neutrophils led to the discovery of signaling pathways essential for the

establishment of neutrophils’ protrusive leading edge. However, there are significant gaps

in our understanding of the mechanisms that govern integrin-mediated leading edge

adhesion during chemotaxis. In this study, we identified a novel signaling mechanism

that controls localized integrin activation and leading edge adhesion during neutrophil

chemotaxis. A model is proposed based on these findings, whereby a central regulatory

role is assigned to the non-receptor tyrosine kinase Lyn (Figure 2.13). In this scenario,

Gi-dependent Lyn activation and translocation to the leading edge is necessary for the

relief of inhibition and spatial organization of the CrkL/C3G complex at the leading edge,

which leads to localized activation of the small GTPase Rap1 and its downstream target

56

β2-integrin. Together, these results provide novel mechanistic insights into a critical but

poorly studied facet of neutrophil chemotaxis – adhesion of the protrusive leading edge.

We demonstrated a critical role for Lyn in regulating leading edge adhesion in

chemotaxing neutrophils. First, in response to chemoattractants, Lyn is activated and

recruited to leading edge of polarized neutrophils. Depletion of Lyn impairs leading edge

protrusion but fails to prevent chemoattractant-induced activation of Rac, polymerized

actin or PI3Ks, suggesting that Lyn regulates leading edge adhesion, not initial

establishment of the leading edge. Second, although some Lyn-depleted cells exhibit a

morphological defect similar to that of cells with the backness pathway impaired, Lyn

depletion does not impair the activity of this pathway. Indeed, a fraction of Lyn-depleted

cells retract their leading edges when exposed to fMLP, a defect rarely seen in cells with

the backness pathway inhibited. Moreover, treatment of cells with Y-27632, unlike Lyn

depletion, fails to impair leading edge protrusion. At the mechanistic level, Lyn depletion

impairs localized activation of β2-integrin, which appears responsible for the chemotactic

defects induced by Lyn depletion, because inhibition of de novo leading edge adhesion

with intermediate concentration of IB4 mimics the phenotype of Lyn-depletion. In

addition, we identified Rap1, a well-recognized regulator of β2-integrin activity, as a

major downstream effector of Lyn in neutrophils during chemotaxis.

How might partial inhibition of leading edge adhesion (e.g., with Lyn or Rap1

depletion, or the treatment with intermediate dose of IB4) lead to an elongated cellular

morphology in neutrophils? We speculate that this defect might result from inability of

the cells to establish firm adhesion sites at the front, a prerequisite for translocation of the

cell body and de-attachment of the trailing edge. Indeed, in each condition wherein

57

leading edge adhesion is impaired, there is a concomitant reduction in retraction speeds,

suggesting linkage between front adhesion and tail retraction. Furthermore, inhibition of

the backness pathway per se (e.g., with Y-27632) does not affect protrusion, again

implying that compromised retraction in Lyn- and Rap1A-depleted cells, as well as in

cells treated with intermediated dose of IB4, might originate from defects in leading edge

adhesion. More complete understanding of the detailed mechanisms underlying the

morphological defects awaits future experiments.

In comparison to earlier studies, our findings have provided a more detailed and

comprehensive delineation of Lyn’s functions in neutrophils during chemotaxis. In

keeping with earlier studies in Lyn-deficient mouse neutrophils (Pereira and Lowell,

2003), our data show that without fMLP stimulation, Lyn-depleted cells exhibit enhanced

adhesion to the fibrinogen substrate and exhibit higher Rap1 activity, suggesting that Lyn

might inhibit Rap1 activity under this condition. In contrast, in the presence of fMLP,

Lyn depletion markedly reduces Rap1 activity and consequently impairs local activation

of β2-integrin in dHL-60 cells. Furthermore, addition of Mn2+

, which induces β2-integrin

activation, causes neutrophils to tightly adhere to the fibrinogen substrate and fail to

polarize in response to a chemoattractant gradient - a phenotype distinct from Lyn-

depleted cells. Together, these results suggest differential regulation and wiring of the

chemotactic signaling network in cells with and without attractant stimulation. In

agreement with this notion, Gakidis et al (Gakidis et al., 2004) showed that Vav1, Vav2

and Vav3 (GEFs for Rac and Cdc42), are required for integrin-dependent functions of

mouse neutrophils, including sustained adhesion, spreading, and complement-mediated

phagocytosis but are dispensable for fMLP–induced signaling events and chemotaxis,

58

even when cells are assessed on the same ECM substrate (Gakidis et al., 2004). Together,

these results highlight the complexity of chemotactic signaling networks that control

neutrophil chemotaxis and underscore the importance of studying cell-ECM interactions

as an integral part of the global chemotactic response. Notably, we did not observe an

increase in total β2-integrin activity in Lyn-depleted dHL-60 cells after fMLP stimulation

in suspension (He, Y. and Wang, F., unpublished observations), in contrast to a previous

study in Lyn-inhibited hematopoietic cells with SDF-1α stimulation (Nakata et al., 2006).

The differential effects of Lyn depletion/inhibition might be attributed to differences in

cell types and chemoattractants. Alternatively, it might be that the experiments for the

detection of active β2-integrin were conducted in suspended cells with prolonged

incubation with the m24 β2-integrin antibody and fMLP, which did not recapitulate the

chemotaxis conditions and may also contribute to the variation.

Our results suggest that the CrkL/C3G protein complex acts as a key signaling

intermediate that controls Lyn-dependent spatial activation of Rap1 in neutrophils. In this

scenario, CrkL associates constitutively with the Rap1 GEF C3G, as in hematopoietic

cells and other cell types. Upon exposure to chemoattractant, the CrkL/C3G complex is

recruited to the leading edge of polarized neutrophils, providing spatially instructive cues

for localized Rap1 activation (Fig. 8). Lyn mediates the leading edge recruitment of

CrkL/C3G, unlikely via direct physical interactions: we did not detect Lyn protein in

CrkL or C3G immunoprecipitates (He, Y. and Wang, F., unpublished observations).

Instead, we speculate that other Lyn-interacting proteins such as hematopoietic cell-

specific Lyn substrate 1 (HS1) (Uruno et al., 2003) or the Shc adaptor protein (Ptasznik et

al., 1995) might play a special role in mediating the recruitment. In addition, we have

59

shown that Lyn modulate CrkL function via the removal of Y207 phosphorylation,

thereby relieving CrkL's intramolecular inhibition. It is of note that the kinetics of Y207

phosphorylation differs from Lyn activation: in response to fMLP the level of p-Y207

remains largely unaltered while Lyn activity is markedly elevated. Therefore, it is not

clear whether Lyn expression, or its activity, might modulate CrkL Y207

dephosphorylation. We postulate that the tyrosine phosphatase SH-2 domain phosphatase

(SHP), reportedly a downstream target of Lyn (Pereira and Lowell, 2003) (Figure 2.13),

might be responsible for the removal of Y207 phosphorylation. However, although SHP-

1 was recruited to the leading edge of neutrophils, the attempt to produce SHP-1-depleted

dHL-60 cells failed in our experiments (data not shown).

60

2.6 Figures

Western: p-Lyn

Western: Lyn

fMLP (100 nM)

0 30 60 120 300Time (s)

IP: Lyn

fMLP

PTX

LY

- + + +

- - + -

- - - +

p-Lyn

Lyn

HEK

293

dHL-60

uHL-60

HEK

293

dHL-60

uHL-60

GAPDH

Lyn

1neutro

phil

Figure 2.1 Lyn expression and activation. (A) ) Lyn expression in human

primary neutrophils (1° neutrophil), dHL-60 (dHL-60), non-differentiated HL-

60 (uHL-60) and HEK 293 cells. GAPDH was a loading control. (B)

Phosphorylation of Lyn in response to fMLP stimulation (100 nM) for various

time points. (C) Western blot analysis of tyrosine-phosphorylated Lyn (p-Lyn)

in dHL-60 cells pretreated with buffer, PTX (1 mg/ml, O/N) or LY-294002 (200

mM, 2 hours) and stimulated with fMLP (100 nM, 1 minute).

A

B

C

61

Lyn F-actin merge DIC

Un

tim

.+

fML

P+

fML

P

dH

L-6

0

Pri

ma

ry

ne

utr

op

hil

Lyn p-Src merge DIC

Figure 2.2 Lyn localization in dHL-60 cells and neutrophils. (A) Immunofluorescence of

Lyn (red) and F-actin (green) in cells plated on fibrinogen without or with fMLP stimulation

(100 nM, 2 minutes). Fluorescent images of Lyn (red), F-actin (green), Lyn/F-actin

merged images and DIC images of cells are shown. (B) Immunofluorescence studies of

Lyn (green) and phosphorylated Src (red) in cells plated on fibrinogen with a uniform

concentration of fMLP (100 nM, 2 minutes). Fluorescent images of Lyn (red), phospho-Src

(green), merged images and DIC images of cells are shown. All bars, 10 mm.

A

B

62

I II

Control

Lyn

depletion

d b

c

db

c

d

b

c

db

c

a aa

a

10 s

10 s

150 s

320 s

350 s

400 s

600 s

600 s

Control

Lyn

depletion

+Y27632

α-tubulin

Lyn

24 h 48 h

NSsiRNAs

LynsiRNAs

NSsiRNAs

LynsiRNAs

0

1

2

3

4

5

Control Lyn depletion

6

7

Pro

tru

sio

nsp

ee

d(µ

m/m

in)

A B

C

D E

Figure 2.3 Lyn depletion impairs neutrophil chemotaxis toward the fMLP delivered by

micropipette. (A) Lyn depletion in dHL-60 cells with Lyn-targeting siRNAs or non–specific (NS)

siRNAs. α-tubulin was a loading control. (B) Immunofluorescence of Lyn in cells transfected

with NS siRNAs (I) or Lyn siRNAs for 48 hours (II). (C) dHL-60 cells with or without Lyn

depletion were allowed to migrate toward chemoattractant-containing micropipette (fMLP, 10

µM) on a fibrinogen-coated substrate. (D) DIC kymographs of a dHL-60 cell transfected with

control siRNAs, Lyn siRNAs or treated with Y-27632 migrating toward chemoattractant-

containing micropipette (fMLP, 10 µM). The left panel shows a portion of the neutrophils’ leading

edge under various conditions. The rectangles indicate the regions of the cell used to generate

the kymographs (before fMLP stimulation). The right panel shows the DIC kymographs. In both

panels, white arrows indicate the direction of protrusion. Approximately 6 minutes of migration

was recorded. (E) Speed of leading edge protrusion for control (n = 30) and Lyn-depleted cells

(n=35) responding to a gradient of fMLP generated by the micropippette. (*, p < 0.001).

63

Co

ntr

ol

Lyn

dep

leti

onI’ II’

I II

-40

-20

0

20

40

20 40 60 80

-40

-20

0

20

40

20 40 60 80

-40

-20

0

20

40

20 40 60 80

-40

-20

0

20

40

20 40 60 80

Control Lyn depletion

(µm) (µm)

Dis

tan

ce (m

m)

0

1

2

3

4

5

Control Lyn depletion

6

Pro

tru

sio

nsp

ee

d(µ

m/m

in)

Figure 2.4 Lyn depletion impairs dHL-60 chemotaxis in a microfluidic gradient

device. (A) Phase-contrast images of control (I, II) and Lyn-depleted cells (I’, II”) in

the microfluidic chamber coated with the fibrinogen substrate before (I, I’) and after

(II, II’) exposure to an fMLP gradient (10 minutes) generated by the microfluidic

device. The gradient can be traced by fluorescent dyes such as fluorescein (left).

Bar, 50 mm. Movies of cells with or without Lyn depletion are available as Movies

S3 and S4. (B) Enlarged views of control cells (top) and Lyn-depleted cells

(bottom) 10 minutes after the stimulation of the fMLP gradient. Bar, 10 mm. (F) The

positions of control and Lyn-depleted cells at the end of the experiments were

plotted, in relation to their starting position at the origin. X axis shows the distance

along the fMLP gradient. Y axis shows the distance vertical to the fMLP gradient.

(C) Speed of leading edge protrusion for control and Lyn-depleted cells responding

to a gradient of fMLP generated by the microfluidic device. The values are means

± SEM (n = 47 for control, 46 for Lyn-depleted cells).

A B

C D

64

Lyn

Lyn-EGFP

Lyn shRNA

pcDNA3/LynKD-EGFP

pcDNA3/Lyn-EGFP - - + -

- - - +

GAPDH

- + + +

10 s 318 s 10 s 320 sD

ICL

yn

-EG

FP

DIC

Lyn

KD

-EG

FP

0

1

2

3

4

5

6

7

Lyn shRNA

pcDNA3/LynKD-EGFP

pcDNA3/Lyn-EGFP - - + -- - - +- + + +

Pro

tru

sio

nsp

ee

d(µ

m/m

in)

*

*

Figure 2.5 Kinase activity is required for the regulation of neutrophil chemotaxis by Lyn.

(A) Expression of Lyn-EGFP or LynKD-EGFP in Lyn-depleted cells. Lyn-depleted cells

were differentiated and transfected with GFP-tagged wild-type Lyn56 or kinase-dead

dominant negative mutant of Lyn56 (K275D) containing silent mutations in the shRNA-

targeting region. Cell lysates were analyzed for Lyn. GAPDH is the loading control. (B)

Wild-type Lyn, but not kinase-dead (K275D) Lyn mutant, rescues the defects of Lyn-

depleted cells. Time-lapse images of representative cells for both conditions are shown.

Bar, 10 μm. (C) Speed of leading edge protrusion for control, Lyn-depleted, Lyn rescued

cells in micropipette assay. The values are means ± SEM. The asterisk indicates that the

cells differ statistically from the control cells (*, p < 0.001)

A

B

C

65

p-Akt

Akt

fMLP stimulation

0 0.5 1 2 5 (min)

Control Lyn depletion

0 0.5 1 2 5Time

p

0

0.1

0.2

0.3

0.4

0.5

0.6

0 30 60 120 300

Time (s)

Ab

so

rban

ce

(490

nm

)

Control

Lyn depletion

Rac-GTP

0

0.5

1

1.5

2

2.5

3

0 30 45 90 300

Time (s)

Re

lati

ve

F-a

cti

n l

eve

l Control

Lyn depletion

0

0.5

1

1.5

2

2.5

3

-

C

B

A

Figure 2.6 Lyn depletion does not affect frontness pathways. (A) dHL-60 cells with or

without Lyn depletion were stimulated for indicated times and lysed. Akt phosphorylation

(Thr308) was assessed by western blotting. Representative blots for phospho- and total

Akt are shown. (B) Quantification of Rac-GTP in dHL-60 cells with or with Lyn depletion.

Cells were stimulated with fMLP (100 nM) for times indicated and lysed. The level of

Rac-GTP was determined by the use of an absorbance-based Rac G-LISA kit. The y

axis represents the absorbance at 490 nm. Each bar represents the mean ± s.e.m.

(error bars) (n=4). (C) Quantification of F-actin polymerization. Cells with or without Lyn

depletion were stimulated with fMLP (100 nM) for various times in suspension and fixed

for staining with rhodamine-phalloidin. Fluorescence in stained cells was determined by

flow cytometry. Values are mean ± s.e.m. (n = 4).

66

Lyn

de

ple

tio

n

b2-integrin F-actin Merge DIC

Co

ntr

ol

Activated

b2-integrin F-actin Merge DIC

Co

ntr

ol

Lyn

de

ple

tio

n

A

B

C

Figure 2.7

67

5 s 305 s 605 sC

on

tro

lIg

G

5 s 305 s 405 s

5 s 305 s 505 s

IB4

(0.5μ

g/m

L)

IB4

(0.1μ

g/m

L)

D

Figure 2.7(cont.) Lyn depletion attenuates β2-integrin activation at the leading

edge. (A) Immunofluorescence of β2-integrin (red) in cells with or without Lyn

depletion, stimulated with a uniform concentration of fMLP (100 nM, 2 minutes).

Fluorescent images of F-actin (green), F-actin/β2-integrin merged images and DIC

images of cells are also shown. Bar, 10 mm. (B) Immunofluorescence of active β2-

integrin (red), stained by antibody m24, in cells with or without Lyn depletion,

stimulated with a uniform concentration of fMLP (100 nM, 2 minutes). Fluorescent

images of F-actin (green), F-actin/active β2-integrin merged images and DIC

images of cells are also shown. Bar, 10 mm. (C) Front vs back ratio of fluorescence

intensity in (A) and (B). The region containing actin staining is defined as front,

while the rest of the cell is defined as back. Fluorescence intensity in the regions of

front and back was quantified by Image J. The ratios for control cells (37 cells for

total β2-integrin and 45 cells for active β2-integrin) and Lyn-depleted cells (53 cells

for total β2-integrin and 46 for active β2-integrin) were calculated. (D) Blocking front

de novo adhesion mimics the defects of Lyn depletion in chemotaxis. Time lapse

images from four representative experiments are shown.

68

0

1

2

3

4

Re

lati

ve

ad

he

sio

n

ControlLyn depletion

No Ab b2-integrin Ab

- fMLP + fMLP - fMLP + fMLPFluorescence intensity

100

101

102

103

104

0

63

125

188

250

b2-intgrinControl

Lyn depletion

Nu

mb

er

of

ce

lls

(10

3)

5 s 105 s 205 s 305 s

Activated

b2-integrin F-actin Merge DIC

A

D

C

B

Figure 2.8 impaired chemotaxis of Lyn depleted cells is not due to altered global adhesion.

(A) dHL-60 cells with or without Lyn depletion were plated on fibrinogen in the absence or

presence of β2-integrin function-blocking antibody (IB4, 1 µg/ml), not stimulated or

stimulated by fMLP (100 nM, 30 min). After four washes the degree of cell adhesion was

assessed. Values were normalized to adhesion in control cells without fMLP or IB4 (=1) and

are means±SEM (n=4). (B) Flow cytometry analysis of surface expression of β2 integrin in

control and Lyn-depleted cells after fMLP stimulation (100 nM, 10 min). The black line

indicates background fluorescence from the isotype control IgG. (C) Time-lapse images of

cells treated with Mn2+ (0.2 mM, 20 min) and exposed to the point source of fMLP from a

micropipette for indicated times. Bar, 10 mm. A typical experiment from six independent

experiments is shown. (D) Immunofluorescence studies of active β2-integrin (red) in cells

with Mn2+ (0.2 mM, 20 min) and thereafter stimulated with a uniform concentration of fMLP

(100 nM, 2 minutes). Fluorescent images of F-actin (green), F-actin/active β2-integrin

merged images and DIC images of cells are also shown. Bar, 10 mm

69

0

0.1

0.2

0.3

0.4

-fMLP +fMLP

Ab

so

rba

nc

e(4

90

nm

) Control

Lyn depletion

Rho-GTP

p-MLC

MHC

fMLP

Y 27632

Control Lyn depletion

+ + + +

+ +

-

- -

-

- -

Control

Lyn depletion

p-MLC / F-actin DIC

Y-27632

A

B

C

Figure 2.9 The backness pathway is not altered in Lyn depleted cells. (A)

Quantification of RhoA-GTP in dHL-60 cells with or with Lyn depletion, not

stimulated or stimulated with fMLP (100 nM, 2 min). Each bar represents the

mean±SEM (error bars) (n=4). (B) Western blot analysis of activated myosin II

(p-MLC) in control cells, Lyn-depleted cells and cells treated with Y-27632 (10

μM, 30 min). All cells were stimulated or not stimulated in fMLP (100 nM) for 2

min in suspension, and the cell lysates were subjected to SDS-PAGE analysis. A

typical experiment from four separate experiments is shown. Myosin heavy chain

(MHC) was used as a loading control. (C) Merged fluorescent images of p-MLC

(red) and F-actin (green) in control, Lyn-depleted cells and cells pretreated with

Y-27632 (30 μM, 30 min). The corresponding DIC images are shown. The white

arrow indicates the loss of p-MLC fluorescence at the back of the Y-27632-

treated cell. Bars, 10 μm.

70

fMLP (100 nM)

- 30 60 120 - 30 60 120

Control Lyn depletion

fMLP (100 nM)

Time (s)

Rap1-GTP

Total Rap1

Lyn

GAPDH

EG

FP

-RalG

DS

DIC

DIC

Co

ntr

ol

Ly

n d

ep

leti

on

EG

FP

-RalG

DS

5 s 105 s 205 s 305 s

0

0.2

0.4

0.6

0.8

1

1.2

Control Lyn depletion

Re

lati

ve

Ra

p1

ac

tiva

tio

n

*

Co

ntr

ol

Lyn

de

ple

tio

n

Figure 2.10 Lyn is required for Rap1 activation and translocation to the leading edge.

(A) The levels of Rap1-GTP in dHL-60 cells under various conditions. dHL-60 cells with

or without Lyn depletion were stimulated with 100 nM of fMLP for various time points

and lysed for the pull-down assay. Levels of total Rap1 and GAPDH were used to show

cell lysate input. The blot for Lyn is also shown. (B) Quantification of relative levels of

Rap1-GTP level in dHL-60 cells with and without Lyn depletion 30 seconds after fMP

stimulation. Each bar represents the mean±SEM (error bars). Values are normalized to

the level of Rap1-GTP (=1) in cells without Lyn depletion. (*, p<0.001). (C) dHL-60 cells

with or without Lyn depletion were transfected with EGFP-RalGDS and stimulated by a

fMLP-containing micropipette for indicated times. EGFP-RalGDS fluorescence and the

corresponding DIC images are shown. (D) Enlarged views of EGFP-RalGDS-

expressing cells with or without Lyn depletion after stimulation with fMLP. The arrow

indicates leading edge recruitment of EGFP-RalGDS.

BA

C D

71

Rap 1

GAPDH

Control

shRNA1

shRNA2

5 s 100 s 305 s155 s

5 s 215 s 435 s

Co

ntr

ol

Rap

1A

sh

RN

A1

5 s 255 s 505 s

Rap

1A

sh

RN

A2

0

1

2

3

4

5

6

7

Control Rap1AshRNA1

Rap1AshRNA2

Pro

tru

sio

nsp

ee

d(µ

m/m

in)

A

B

C

Figure 2.11

72

EG

FP

-Rap

1A

EG

FP

-Rap

1A

DIC

DIC

Co

ntr

ol

Lyn

dep

leti

on

5 s 105 s 205 s 255 s

5 s 105 s 205 s 255 s

D

Figure 2.11 (cont.) Rap1A depletion mimics the chemotactic defects of Lyn

depletion. (A) Western blotting of Rap1A in dHL-60 cells with or without shRNAs

targeting Rap1A. GAPDH is a loading control. (B) dHL-60 cells with Rap1A

depletion(Rap1 A shRNAs) or without (Control, with non-targeting shRNAs)

stimulated on a fibrinogen-coated substrate by a point source of fMLP. The four

images in each row show the positions of individual cells after exposure to fMLP. (C)

Speed of leading edge protrusion for control and Rap1A-depleted cells in the

micropipette assay. The values are means ± SEM. The asterisk indicates that the

Rap1A-depleted cells differ statistically from the control cells (*, p < 0.001) (D) dHL-

60 cells with or without Lyn depletion were transfected with EGFP-Rap1A and

stimulated with a fMLP-containing micropipette. All bars, 10 μm.

73

Control Lyn depletion

Unst

im.

30 s

1m

in3

min

5m

inUnst

im.

30 s

1m

in3

min

5m

in

fMLP fMLP

p-CrkL

Lyn

CrkL

a-tubulinC

on

tro

lL

yn

dep

leti

on

CrkL F-actin DICMerge

Pri

mary

neu

tro

ph

ils

Co

ntr

ol

Lyn

dep

leti

on

C3G F-actin DICMerge

A

B

C

Figure 2.12

74

C3G

CrkL

Rap1

Rap1-GTP

fMLP - + - +

Control CrkL depletion

5 s

105 s

345 s

Control

5 s

105 s

345 s

505 s

a

b

a

b

a

b

a

b

5 s

105 s

345 s

405 s

CrkL shRNA1 CrkL shRNA2D E

Figure 2.12 (cont.) Lyn is required for the functions of CrkL/C3G complex in

neutrophils. (A) Western blotting of phosphorylated CrkL (p-CrkL, at Y207) in dHL-60

cells with or without Lyn depletion. Cells were unstimulated or stimulated in

suspension with 100 nM fMLP for various times. CrkL and α-tubulin were loading

controls. Lyn expression was also analyzed. (B and C) Immunofluorescence of CrkL

and C3G in dHL-60 cells with or without Lyn depletion. Cells were stimulated with a

uniform concentration of fMLP (100 nM, 2 minutes). Immunofluorescence of CrkL in

human primary neutrophil is also shown. (D) Levels of Rap1-GTP in dHL-60 cells with

or without CrkL depletion 30 seconds after fMLP stimulation. 30 seconds of fMLP

stimulation gave rise to maximal Rap1 activation . Blot for total Rap1 shows input

from cell lysates for each condition. Blots for CrkL and C3G are also shown. (E) dHL-

60 cells with or without CrkL depletion were stimulated with a micropipette containing

fMLP. The images in each row show the positions of individual cells (identified with a

superimposed letter) after exposure to fMLP. All bars, 10 μm.

75

Gi Lyn CrkL C3G

GPCR

Rap-GTP

SHP ?

Rap-GDPRecruitment

Activation

Relief of inhibitionRecruitment

a b

b2-integrin

pY398 Y207

Figure 2.13 A biochemical pathway is required for leading edge adhesion,

localized b2-integrin activation and persistent migration in neutrophils during

chemotaxis. Lyn is a activated downstream effector of ligated GPCR and

translocated to leading edge. Active Lyn then recruits and activates CrKL/C3G

complex, which further spatially activate Rap1 in the leading edge. Activated

Rap1 up-regulates β2 integrin and enhances the front adhesion of chemotactic

neutrophils. GPCR: G-protein coupled receptor.

76

CHAPTER 3 MAMMALIAN TARGET OF RAPAMYCIN COMPLEX 2

(mTORC2) REGULATES ACTIN POLYMERIZATION AND NEUTROPHIL

CHEMOTAXIS

3.1 Summary

Chemotaxis allows neutrophils to seek out sites of infection and inflammation.

The signaling mechanism underlying neutrophil polarity and chemotaxis has been

intensively studied and is still not fully understood. The asymmetric accumulation of

filamentous actin in the leading edge is essential for the development and maintenance of

neutrophil polarity and has been postulated as the driving force for neutrophil motility.

By using RNA interference, pharmacological and biochemical approaches, we

demonstrated that mTORC2 is an essential regulator of actin cytoskeleton in neutrophils

and mTORC2 is required for neutrophil chemotaxis. Depletion of mTORC2 impaired

chemotaxis of dHL-60 cells, a neutrophil-like cell line. Although depletion of mTORC2

abolished AKT phosphorylation, a downstream event of mTORC2 activation, inhibition

of AKT phosphorylation failed to recapitulate the migratory defect of mTORC2 depletion

in both dHL-60 cells and neutrophils. Furthermore, inhibition of kinase activity of

mTORC2 also failed to alter neutrophil chemotaxis. At the molecular level, mTORC2

depletion severely impaired actin polymerization induced by chemoattractant stimulation.

Consistent with this finding, mTORC2 depletion reduced chemoattractant-induced Rac

activation under both adherent and non-adherent conditions. We propose here that

mTORC2 regulates neutrophil chemotaxis through Rac-mediated signal pathways.

77

3.2 Introduction

Neutrophils are critical for the host innate immune response against invading

microbes. Chemotaxis, the directed movement of cells, allows circulating neutrophils in

the bloodstream to crawl toward sites of infection and inflammation. Upon stimulation by

chemoattractants, neutrophils respond by establishing a leading edge (pseudopod), which

protrudes toward the source of the chemoattractant, and a trailing edge (uropoda), which

faces away from the chemoattractant gradient. Recently work has begun to unfold the

mechanism underlying the polarity and chemotaxis of neutrophils. In the leading edge,

signaling pathways, including PI3Kγ and Rac, control actin polymerization for membrane

protrusion (Sasaki et al., 2000; Weiner et al., 2002; Weiner et al., 1999). RhoA-regulated

actin-myosin contractility is responsible for the retraction of the trailing edge (Xu et al.,

2003). The asymmetric accumulation of filamentous actin (F-actin) in the leading edge is

the hallmark of neutrophil polarity and also has been postulated to provide the driving

force for neutrophil chemotaxis. Thus, the delineation of signaling pathways regulating

actin polymerization induced by chemoattractants is the key for better understanding the

molecular mechanism underpinning neutrophil polarity and chemotaxis.

mTORC2 has emerged as the key regulator for actin cytoskeleton in mammalian

cell lines (Jacinto et al., 2004; Sarbassov et al., 2004). In contrast to mTORC1 in

mammals, which has been intensively studied for many years and shown to be the key

regulator for cell growth in response to nutrient signals, the functions of mTORC2 remain

elusive (Jacinto and Hall, 2003; Loewith et al., 2002). In budding yeast, Torc2 has been

shown to be resistant to rapamycin treatment and regulates bud formation (Loewith et al.,

2002). Recently the counterpart of yeast Torc2 in mammals, mTORC2, has been

78

identified in mammals and is also insensitive to rapamycin treatment (Jacinto et al., 2004;

Sarbassov et al., 2004). Depletion of mTORC2 in HeLa cells and 3T3 fibroblasts

disrupted stress fibers and dramatically altered the structure of actin cytoskeleton.

Attempts to generate rictor knockout mice failed due to embryonic lethality (Guertin et

al., 2006; Shiota et al., 2006). In Dictyostelium discoideum, homologues of mTORC2

components have been identified: LST8, RIP3 (AVO1) and Pia (AVO3, mAVO3, Rictor)

(Cai et al., 2010; Lee et al., 2005). The deletion of TORC2 did not affect survival, but

impaired D. discoideum chemotaxis induced by cAMP. TORC2 acted downstream of Ras

and regulates PKBR1 and PKBA phosphorylation which was required for actin

polymerization in D. discoideum chemotaxis (Cai et al., 2010). The embryonic lethality

of mTORC2 has hampered studies of mTORC2 functions in mammalian tissues and

cells, including neutrophils. During preparation of our paper, a study with Rictor

depletion in PBL cell line showed that mTORC2 is required for neutrophil chemotaxis by

regulating cAMP production and RhoA activation (Liu et al., 2010). Interestingly, the

actin polymerization in cells with mTORC2 depletion is shown to be comparable to the

control in their report.

Here by using a more commonly used neutrophil-like cell line HL-60 cell, we

reported that mTORC2 depletion impaired neutrophil chemotaxis through disrupting the

reorganization of actin cytoskeleton. mTORC2 was required for fMLP-induced actin

polymerization and motility of neutrophils. Importantly, this requirement was not

dependent on kinase activity of mTOR and the downstream event of AKT

phosphorylation. Furthermore, we showed that mTORC2 depletion inhibited Rac

activation, which is required for actin polymerization during neutrophil chemotaxis.

79

3.3 Materials and methods

Reagents and plasmids

Human fibrinogen, human serum albumin (HSA) and fMLP were from Sigma-

Aldrich. Rapamycin, Y27632 and Akti1/2 were from Calbiochem. Torin1 was kindly

provided by Drs. Nathanael S. Gray (Dana-Farber Cancer Institute, Harvard) and David

M. Sabatini (Whitehead Institute, MIT). Rabbit anti-Rictor antibody and Rictor blocking

peptide were from Bethyl laboratories (Montgometry, TX). Rabbit anti-mTOR antibody,

rabbit anti-Raptor antibody, rabbit anti-Akt antibody, mouse anti-phospho [Ser473] and

rabbit anti-phospho [Thr308]-Akt antibodies, mouse anti-phospho [Thr389]-S6K1, rabbit

anti-phospho [Ser19] myosin light chain antibody, were from Cell Signaling Technology.

Alexa fluor 488 phalloidin, alexa fluor 647 phalloidin and alexa fluor 594 conjugated

secondary antibodies were from Invitrogen. Goat anti-GAPDH polyclonal antibody was

from GenScript. All secondary antibodies for western blotting were from Jackson

ImmunoResearch Laboratories. Other reagents if not mentioned specifically were from

Sigma.

Three shRNA constructs targeting human mTOR were from Sigma [SHCLNG-

NM_004958:TRCN0000039783,TRCN0000039784 and TRCN0000039785).Two

shRNA constructs targeting human Raptor (id # 1857 and 1858 ) or Rictor (id # 1853 and

1854) were requested from Addgene.

80

Cell culture, lentiviral production and infection

Cultivation and differentiation of HL-60 cells were described previously (Wang et

al., 2002a). HL-60 cells stably expressing Actin-YFP were from Dr. Henry Bourne lab

(University of California at San Francisco).

For lentiviral production, shRNA-containing plasmids were co-transfected with

three packaging plasmids pPL1, pPL2 and pPL/VSVG (Invitrogen) into actively growing

HEK293T cells. After 3 days of transfection, supernatant containing the lentiviral

particles was concentrated by Lenti-X concentrator (Clontech) and added to growing HL-

60 cells in the presence of 6 μg/ml polybrene for 18-24 hours. Infected cells were

transferred to fresh culture medium containing 1.3% DMSO and 0.25 µg/ml puromycin

and differentiated for 4-5 days.

Immunofluorescence and live-cell imaging

For immunofluorescence studies, cells were suspended in mHBSS buffer (20 mM

HEPES, pH 7.4, 50 mM sodium chloride, 4 mM potassium chloride, 1 mM magnesium

chloride, 10 mM glucose) with 1% HSA, and then plated on fibrinogen-coated cover

glass (250 μg/ml fibrinogen) (Schymeinsky et al., 2005) for 20 minutes, washed briefly

and subsequently stimulated with a uniform concentration of 100 nM fMLP for indicated

times. After stimulation, cells were immediately fixed with 3.7% paraformaldehyde in

CSK buffer (10 mM HEPES, pH 7.4, 138 mM KCl, 3 mM MgCl2, 2mM EGTA, 150 mM

sucrose) for 15 minutes, washed and permeablized for 10 minutes by using CSK buffer

containing 0.2% Trinton X-100. Cells were blocked in CSK buffer containing 1% BSA

for 1 hour, incubated with primary antibodies overnight at 4°C followed by Alexa fluor

81

594 conjugated goat anti-rabbit or anti-mouse secondary antibody. Primary antibodies

were used at dilutions of 1:100 (Rictor) and 1:200 (p-MRLC). Secondary antibodies were

used at 2 μg/ml. For the detection of F-actin, cells were incubated with Alexa fluor 488-

conjugated phalloidin for 10 minutes at RT.

For live-cell imaging, cells were plated on human fibrinogen-coated surface for

20 minutes, washed briefly, and subsequently stimulated with a point source of 10 µM

fMLP from a micropipette (internal diameter of 1 µm, pulled from a glass capillary), as

described (Servant et al., 2000a). DIC images, fluorescent images and combined

DIC/fluorescence images were collected with a Zeiss 40X NA 1.30 Fluar DIC objective

or 63X NA 1.4 Plan Apochromat DIC objective on a Zeiss Axiovert 200M microscope.

All images were collected with a cooled charge-coupled device camera (AxioCam MR3,

Zeiss) and processed by the Image J program.

Western blotting

Cells (5 x 106) were suspended in 90 μl of mHBSS buffer and stimulated with 10

μl of 1 μM fMLP for various times. fMLP stimulation was terminated by the addition of

equal volume of 2 x RIPA buffer (50 mM Tris-HCl, pH 7.5, 1% NP-40, 150 mM NaCl, 1

mM EDTA) containing 1 mM Na3VO4, 1 mM NaF, 1 mM PMSF and a protease inhibitor

cocktail (Sigma). Cell lysates were clarified by centrifugation at 14,000 x g for 10

minutes. Protein concentrations of the lysates were determined by using DC Protein

Assay Kit (Bio-Rad). Equivalent amounts of protein (20 μg) were resolved on a 8%

disulfide-reduced SDS-PAGE gel, transferred to Trans-Blot Transfer Medium, blocked

with 5% nonfat milk in TBST for 1 hour at room temperature, and incubated with the

82

primary antibody O/N at 4°C followed by a secondary antibody conjugated to

horseradish peroxidase (HRP; GE Healthcare). The HRP signals were detected by using

enhanced chemiluminescence (SuperSignal West Pico). The western blot was stripped

and re-probed with appropriate antibody according to the manual (Restore Western Blot,

Thermo Scientific).

Microfluidic gradient device

The microfluidic gradient device comprises of a polydimethylsiloxane (PDMS)

slab embedded with a Y-shaped fluidic channel and glass substrate as its base. The

microchannels were fabricated using rapid prototyping and soft lithography as described

previously (Herzmark et al., 2007; Jeon et al., 2000; Lin and Butcher, 2006). Cells were

washed, suspended in mHBSS containing 1% HSA and injected in the microfluidic

device. The device was left in the incubator (37°C, 5% CO2) for 20 minutes to allow

cells to attach to the substrate. The channels were then gently rinsed with mHBSS to

wash away floating cells

To generate soluble gradients, fMLP (500 nM) and mHBSS were infused into the

device from separate inlets. Solutions were driven using a syringe infusion pump (Pump

22, Harvard Apparatus) at a flow rate of 0.03 ml/h to generate a concentration gradient in

the channel by diffusion. Phase contrast images were captured using Axiovision software

on a Zeiss Axiovert 200M microscope.

Actin polymerization assay

The procedure for measuring polymerized actin was according to previous report

with some modifications (Weiner et al., 2006). Briefly, 1×106 cells were starved in

83

serum-free medium for 1 hour at 37°C and suspended in mHBSS buffer. Cells were

stimulated with 100 nM fMLP for indicated times. After stimulation, cells were fixed by

adding equal volume of 7.4% paraformaldehyde for 30 minutes at room temperature.

Fixed cells were permeabilized and stained in PBS buffer containing 0.1% Triton X-100

and Alexa 647-conjugated phalloidin (1:100) for 30 minutes at 37°C in dark. Stained

cells were washed once and suspended in ice-cold PBS, and fluorescence was

immediately determined by flow cytometry with a BD Biosciences LSR II System. The

mean fluorescence intensity of the cell population was determined.

Rac pull-down assay

GST (glutathione S-transferase) – PAK1 – PBD (p21-binding domain) fusion

protein was prepared according to previous report (Benard et al., 1999). dHL-60 cells (5

× 106 cells) containing NT shRNA or Rictor shRNA were starved in serum-free medium

for 1 hour at 37°C. Cells were then suspended in mHBSS buffer and stimulated by 100

nM fMLP for indicated times. Cell stimulation was stopped by adding 0.4 ml 1.25 × lysis

buffer(1× lysis buffer: 25 mM Tris-HCl, pH 7.5, 100 mM NaCl, 5 mM MgCl2, 1% NP-

40, and 5% glycerol, 1 mM phenylmethylsulfonyl fluoride, 1 mM Diisopropyl

phosphorofluoridate, and protease inhibitor). Cell lysates were immediately put on ice

and pipetted for 4-5 times. The Subsequent steps were performed as previous description

(Benard et al., 1999). For adhesion condition, cells were plated on fibrinogen (100 µg/ml)

coated 6-well plates for 30 minutes and then stimulated with 100 nM fMLP for indicated

times. Cells including adherent and non-adherent were lysed, and the same procedures as

above mentioned were performed to pull down the activated Rac.

84

RhoA activation assay

An absorbance-based (490 nm) RhoA G-LISA kit (Cytoskeleton, Inc) was used to

determine RhoA-GTP levels in dHL-60. Cells with or without Rictor depletion were

suspended in mHBSS (5 × 106 cells in 90 μl per condition), and stimulated or

unstimulated with 10 μl of 1 μM fMLP. The reaction was stopped by adding 100 μL of 2

× lysis buffer (provided with the kit) at 4°C. Subsequent steps were performed as

described in the protocol attached to the kit. For adhesion condition, cells were plated on

fibrinogen (100 µg/ml) coated 6-well plates for 30 minutes and then stimulated with 100

nM fMLP for indicated times. Cells including adherent and non-adherent were lysed, and

the same procedures as described in the kit were performed to determine RhoA activity.

Isolation of primary neutrophils

Primary neutrophils were isolated from venous blood from healthy human donors.

Blood was collected into heparin-containing Vacutainer tubes (BD Biosciences) and

neutrophil isolation procedure was performed within 30 minutes of blood collection using

PMN isolation medium (Matrix). Red blood cell contaminants were removed by Red

Blood Cell Lysis buffer (Roche), which produced more than 97% of neutrophil purity.

Neutrophils were suspended in RPMI 1640 medium supplemented with 10% fetal bovine

serum at 37°C until the time of experiments conducted within 8 hours after isolation.

Statistical analysis

Statistically significant differences between two groups were determined using

the heteroscedastic two-tailed Student’s t-Test. The difference with p value less than 0.05

is considered to be significant.

85

3.4 Results

mTORC2 is required for neutrophil chemotaxis

To explore the role of mTOR signaling in neutrophil chemotaxis, we assessed the

effects of mTORC1 and mTORC2 depletion on neutrophil chemotaxis (Figure 3.1 and

Figure 3.2). A human promyelocytic leukemia cell (HL-60), which can be differentiated

into neutrophil-like cells in the presence of DMSO and is genetically tractable, has been

widely used as a model for studying neutrophil chemotaxis (Bodin and Welch, 2005;

Hauert et al., 2002; Nuzzi et al., 2007; Schymeinsky et al., 2005; Servant et al., 2000b;

Weiner et al., 2006). We identified that both HL-60 cells (uHL-60) and differentiated

HL-60 cells (dHL-60) expressed components of mTOR complexes, mTOR, Rictor and

Raptor (Figure 3.1A). The expression of mTOR, Rictor and Raptor in human primary

neutrophils was confirmed in this study.

By using a lentiviral transfection system, we were able to transiently knock down

the expression of genes of interest in dHL-60 cells, which allows us to quickly test gene

function in dHL-60 cell chemotaxis. HL-60 cells were incubated with lentivirus overnight

and subsequently differentiated for 4-5 days. In contrast to non-target shRNA, shRNAs

for mTOR, Raptor and Rictor efficiently knocked down their respective proteins (Figure

3.1B-C). Protein levels were reduced after the introduction of appropriate shRNAs:

mTOR shRNA-1, 36.3% ± 4.1 %; mTOR shRNA-2, 20.8% ± 2.8 %, mTOR shRNA-3,

5.9% ± 5.0 %; Raptor shRNA-1, 21.4% ± 5.2%; Raptor shRNA-2, 13.4% ± 3.8%; Rictor

shRNA-1, 17.1% ± 9.1 %; Rictor shRNA-2, 15.2% ± 7.8 %. The chemotactic behaviors

of mTOR, Raptor or Rictor-depleted cells were examined by using a microfluidic

gradient device, which enabled us to watch populations of cells moving in a highly stable

86

gradient over a long distance (150 µm). Time-lapse video-microscopy was used to track

individual cells as they migrated in the microfluidic chamber. In this assay, a majority of

cells with non-target shRNA (70.6% ± 3.8%) and Raptor shRNAs (Raptor shRNA-1,

69.4% ± 4.8%; Raptor shRNA-2, 69.1% ± 7.9%) polarized and moved toward the

concentration gradient (1.4 nM/μm) of the bacteria-derived chemoattractant formyl-Met-

Leu-Phe (fMLP), resulting in few cells in the low concentration region of the chamber

(Figure 3.2A-D). As shown in the cell trajectory, most control cells or cells with Raptor

knockdown developed typical polarized morphology during chemotaxis, exhibiting a flat

broad front and a narrow tail (Figure 3.2C). In contrast, cells with mTOR or Rictor

depletion showed severe defects in chemotaxis (Figure 3.2A-D). Few cells migrated

toward the gradient of fMLP (mTOR shRNA-2, 22.1% ± 7.0%; mTOR shRNA-3, 11.9%

± 3.2%; Rictor shRNA-1, 9.3% ± 1.1%; Rictor shRNA-2, 12.9% ± 5.0%) and remained

in the lower concentration region of the chamber. At the end of stimulation, most cells

with mTOR or Rictor depletion failed to develop polarized morphology (Figure 3.2B).

These cells randomly extended small protrusions along the membrane (Figure 3.2C). The

chemotactic behaviors of cells treated with shRNAs were also analyzed in a point source

of fMLP gradient delivered by a micropipette (Figure 3.2E). Similarly, cells with mTOR

or Rictor depletion failed to migrate toward the tip. Taken together, mTORC2, not

mTOC1, regulated chemotaxis in dHL-60 cells.

TOR signaling pathways play an essential role in cell growth. To assess whether

the migratory defect of cells with mTOR or Rictor depletion is due to the non-specific

toxicity of protein depletion, we examined the percentage of apoptotic cells in

differentiated cells transfected by shRNAs (Figure 3.3). Rictor depletion (Rictor shRNA-

87

1) in dHL-60 did not increase apoptotic cells within 4-5 days when compared to control,

although mTOR depletion caused an increase of apoptotic cells (Figure 3.3A). In

addition, the differentiation process to neutrophils, indicated as the expression level of

beta 2 integrin, appeared to be unaffected in Rictor depleted cells. However, the depletion

of mTOR kinase dramatically reduced the expression of beta 2 expression (Figure 3.3B).

Furthermore, AKT phosphorylation (pT308), a downstream event of fMLP-induced PI3K

activation, was not affected in cells with Rictor depletion (data not shown).

mTORC2-dependent AKT phosphorylation is not required for neutrophils chemotaxis

mTORC2 is the key kinase that phosphorylates a serine site (S473) at the

hydrophobic motif of AKT in response to nutrient signals (Sarbassov et al., 2005). In

budding yeast, TorC2 regulates actin polymerization through the ACG kinase YPK2, the

yeast AKT homologue (Kamada et al., 2005; Schmidt et al., 1996). Similarly, TORC2 in

D. discoideum activates PKBR1 and PKBA, the Dictyostelium AKT homologue, in the

leading edge of cells, and plays an important role in the regulation of actin cytoskeleton

and chemotaxis (Kamimura et al., 2008; Lee et al., 2005). To examine whether the

mTORC2-AKT signaling pathway plays a similar role in neutrophil chemotaxis, we

examined the phosphorylation of AKT in neutrophils stimulated by fMLP, and how the

inhibition of AKT phosphorylation affected neutrophil chemotaxis (Figure 3.4).

Differentiated HL-60 cells and human primary neutrophils were stimulated with

fMLP in suspension, which excluded the influence of extracellular substrate interaction.

The endogenous level of phosphorylated AKT was assessed by using a specific antibody

for p-AKT (Ser473). Exposure of suspended dHL-60 cells (i.e., in the absence of ECM

88

attachment) to the fMLP induced rapid and robust phosphorylation of AKT, reaching a

maximum at 60 seconds, as detected by the phospho-specific antibody (Figure 3.4A-B).

A similar pattern of AKT phosphorylation was also observed in human primary

neutrophils after fMLP stimulation, although the peak of phosphorylation happened at a

later time (data not shown). Rictor depletion, which specifically disrupts mTORC2,

abolished AKT phosphorylation at serine 473 (Figure 3.4C). The level of AKT

phosphorylation at serine 473 in Rictor-depleted cells was reduced to 12.4 % ± 11.2% of

control cells at the stimulation time point of 1 minute (Figure 3.4D). In contrast, the

phosphorylation of AKT at theorine 308, which is catalyzed by PDK1, was not altered in

Rictor-depleted cells (data not shown). This result suggested that in neutrophil-like cells,

dHL-60 cells, mTORC2 is also the major regulator for AKT phosphorylation at serine

473. To ask whether the inhibition of AKT phosphorylation at serine 473 is responsible

for the migratory defect observed in Rictor depleted cells, like the case in Dictyostelium,

we used a specific inhibitor Akti-1/2, shown to abolish the phosphorylation at both sites

in vitro and in vivo (Barnett et al., 2005), to test whether the inhibition recapitulates the

chemotactic defect of cells with mTORC2 depletion. We observed the potent inhibition

of AKT phosphorylation at both sites even at low concentration of Akti-1/2 (0.1 µM)

(Figure 3.4E). However, pretreatment of Akti-1/2 had no effect on chemotaxis of dHL-60

cells and human primary neutrophils in the gradient of fMLP generated by microfluid

device (Figure 3.4F and data not shown). In the micropipette migration assay, Akti-1/2

treated dHL-60 cells moved toward the micropipette tip with a similar polarized

morphology and speed comparable to control cells (data not shown). These data indicate

89

that AKT phosphorylation is not the essential downstream event of mTORC2-mediated

pathway in the regulation of neutrophil chemotaxis.

mTORC2 regulates neutrophil chemotaxis through kinase activity- independent pathways

Since AKT phosphorylation at serine 473 was not involved in mTORC2-mediated

signaling pathway regulating neutrophil chemotaxis, what would be the downstream

targets for this mTORC2-mediated pathway? Rapamycin, a potent and specific inhibitor

for mTORC1, has been shown to inhibit many signaling pathways of mTORC1, although

some rapamycin insensitive activities have been reported in mTORC1 (Wullschleger et

al., 2006). Recently Torin1, an ATP analogue for mTOR kinase, has been developed.

Exposure of cells to Torin 1 inhibits a variety of mTORC1 and mTORC2 activities

(Thoreen et al., 2009). To further understand whether other kinase activities of mTORC2,

instead of AKT phosphorylation, are responsible for the regulation of neutrophil

chemotaxis by mTORC2, we examined the effects of Torin 1 treatment on neutrophil

chemotaxis (Figure 3.5). In this study, we also included the rapamycin treatment as a

comparison.

Rapamycin treatment reduced S6K1 phosphorylation (1 minute time point:

control, 88.8% ± 11.2%; rapamycin, 20.5% ± 14.3%; Means ± s.e.m), while there was no

effect on AKT phosphorylation (Figure 3.5A-C). In contrast, Torin 1 treatment reduced

the phosphorylation level of both S6K1 and AKT(p-T389 S6K1: control, 88.8% ± 11.2%;

torin 1, 12.6% ± 6.8%. p-S473 AKT: control, 93.5% ± 6.5%; torin 1, 12.6% ± 6.8%. 1

minute time point) (Figure 3.5A-C). Furthermore, Torin1 also inhibited AKT

phosphorylation induced by fMLP in isolated human primary neutrophils (data not

90

shown). Next we assessed the effects of these inhibitor treatments on neutrophil and

dHL-60 chemotaxis by using the microfluid device. Both rapamycin and Torin 1 had no

detectable effect on chemotaxis of dHL-60 and neutrophils (Figure 3.5D,E and data not

shown). Both the speed and chemotaxis index of rapamycin or torin 1-treated cells were

comparable to those of control cells (Table 1). These data indicated that mTOR kinase

activity was not required for mTORC2-mediated signaling pathway regulating neutrophil

chemotaxis.

mTORC2 accumulation in the leading edge of chemotactic neutrophils

After chemoattractant stimulation, neutrophils respond by establishing the

polarity with asymmetric accumulation of signaling molecules in the leading edge or

trailing edge: PI3Ps, active Rac and filamentous actin accumulate in the leading edge

(Wang, 2009); RhoA and actin-myosin accumulate in the trailing edge (Xu et al., 2003).

The spatial distribution of these molecules indicates their distinct functions in neutrophil

polarity and chemotaxis. In D. discoideum, TORC2 has been postulated to activate

PKBR1 and PKBA in the leading edge (Kamimura et al., 2008). To explore the

mechanism of mTORC2 regulation of neutrophil chemotaxis, we examined the cellular

localization of mTORC2 before and after fMLP stimulation by Rictor immuno-staining.

At the resting condition, Rictor concentrated in the nuclear region of dHL-60 cells with a

little accumulation at the surrounding region beneath the membrane. In the same cells,

filamentous actin accumulated at the membrane cortical region (Figure 3.6A, top panel).

After 3 minutes of uniform fMLP exposure, we observed an accumulation of Rictor in

the leading edge, co-localizing with F-actin (Figure 3.6A, Middle panel). The

fluorescence intensity of stained Rictor and F-actin across the body was revealed in the

91

line scanning profile (Figure 3.6B). We defined the first 4 microns of leading edge as the

“front” and the remaining cell body as the “back”. The fluorescence intensity ratio of

front vs back was 1.89 ± 0.09 (Mean ± s.e.m) (Figure 3.6C). The specificity of Rictor

staining was further confirmed by the control experiment with blocking peptide (Figure

3.6A, the low panel).

The co-localization of Rictor and F-actin in the leading edge of cells raised the

question of whether they directly associated with each other. We used a biochemical

approach to ask whether Rictor or mTORC2 directly associate with filamentous actin.

Resting and stimulated dHL-60 cells were lysed in Triton X-100 lysis buffer with

different concentrations of Triton x-100 (1% vs 0.1%). In control experiments, a large

proportion of Rictor was released into the soluble fraction of cell lysate, independent of

the concentrations of detergent used. After stimulation, we observed an increased

proportion of actin retained in the insoluble fraction. Treatment with Latrunculin B

completely eliminated the actin from the insoluble fraction of the cell lysate, suggesting

that actin polymerization was inhibited in cells. However, the protein level of Rictor in

Triton-X 100 insoluble pellet was not altered by the treatment of Latrunculin B. This

evidence indicates that Rictor does not directly associate with the actin cytoskeleton (data

not shown).

mTORC2 is required for actin polymerization in neutrophils stimulated by fMLP

As both AKT phosphorylation and mTOR kinase activity were found not to be in

the regulatory pathway of neutrophil chemotaxis by mTORC2, what might be the

mechanism used by mTORC2 to regulate neutrophil chemotaxis? Previous reports have

92

suggested a physiological role for TORC2 in regulation of the actin cytoskeleton

(Wullschleger et al., 2006). TORC2 in budding yeast regulated the cell cycle-dependent

polarization of actin cytoskeleton (Schmidt et al., 1996). In D. discoideum, TORC2

controlled actin polarization and cAMP-induced chemotaxis (Kamimura et al., 2008; Lee

et al., 2005). In HeLa cells and 3T3 fibroblasts, depletion of mTORC2 caused structural

changes in the actin cytoskeleton, leading to disassembly of stress fibers (Jacinto et al.,

2004; Sarbassov et al., 2004). Both dHL-60 cells and primary neutrophils undergo

reorganization of the actin cytoskeleton in response to chemoattractants during

chemotaxis. Inhibition of actin cytoskeletal dynamics by inhibitors impaired neutrophil

chemotaxis.

To examine whether the defects of chemotaxis in Rictor- depleted cells is indeed

due to the inhibition of actin polymerization, we measured the content of filamentous

actin (F-actin) in cells with or without Rictor depletion (Figure 3.7). Exposure of

suspended dHL-60 cells induced transient actin polymerization, which could be measured

by flow cytometry after fixation. In control cells (Non-target shRNA), fMLP treatment

induced a quick and transient F-actin formation, with a peak at around 1 minute (2.44 ±

0.14 fold of resting cells, n=3) (Figure 3.7A). Rictor depletion caused the reduction of F-

actin levels even at resting condition (Rictor shRNA-1, 0.74 ± 0.07; Rictor shRNA-2,

0.75 ±0.12; non-target shRNA) (Figure 3.7A). In addition, treatment with torin 1 or Akti-

1/2 failed to alter F-actin formation (data not shown). When cells were plated on the

substrate fibrinogen and exposed to uniform concentration of fMLP, control cells

developed polarized morphology with F-actin concentrated in the leading edge as

revealed by fluorescence-conjugated phallodin staining (Figure 3.7B, the top panel).

93

Under the same conditions, cells with Rictor depletion failed to develop fully polarized

morphology (Figure 3.8B, the middle and bottom panel). Along the membrane, we often

observed small protrusions containing patched F-actin. After exposure to uniform fMLP

stimulation, about 12% of Rictor-depleted cells polarized with a typical morphology,

compared to about 70% of control cells (Figure 3.7C).

In live cell experiments, we expressed non-target shRNA or rictor shRNA in

actin-YFP stable expressing cells and assessed the localization of actin-YFP in live cells.

Control cells polarized and migrated toward the micropipette tip. During the process of

migration, initially diffuse actin-YFP translocated to the membrane and finally

concentrated at the leading front of chemotaxing cells (Figure 3.7D). In contrast, actin-

YFP remained diffuse in cells with Rictor depletion through the course of experiment

(Figure 3.7D). Taken together, mTORC2 disruption by Rictor depletion significantly

inhibits fMLP-induced actin polymerization in dHL-60 cells.

Disruption of mTORC2 inhibits Rac activation

How might mTORC2 regulate actin polymerization during neutrophil

chemotaxis? Among the Rho GTPases, Rac proteins have been documented as the key

regulators for actin cytoskeleton in leukocytes and many other cells (Etienne-Manneville

and Hall, 2002; Sanchez-Madrid and del Pozo, 1999). Rac2 has been identified only in

hematopoietic cells and is more abundant than Rac1 in neutrophils. Rac2 knockout in

mice or expression of dominant negative Rac2 mutant in human patients caused the

dramatic attenuation of actin polymerization in neutrophils (Roberts et al., 1999;

Williams et al., 2000). In NIH 3T3 cells, mTORC2 depletion inhibited Rac activation

94

induced by serum (Jacinto et al., 2004). To examine whether mTORC2 depletion also

affects Rac activation induced by chemoattractants, we assessed the level of GTP-bound

Rac by using GST-PBD pull down assay (Figure 3.8). After fMLP stimulation, we

observed that the level of Rac-GTP in control cells quickly increased within 30 seconds

and returned back to the resting level after 5 minutes. In Rictor-depleted cells, the level of

active Rac was reduced to 37.6% ± 3.7% of control (Figure 3.8A,B). The reduced level of

Rac-GTP corresponded well to the level of F-actin in Rictor depleted cells. When cells

were plated on fibrinogen-coated plates, Rac activation was already observed in control

cells even without chemoattractant stimulation, which is consistent with a previous report

that integrin ligation can induce Rac activation (Price et al., 1998). After fMLP

stimulation, the level of Rac-GTP increased about 10~20 % in 1 minute and returned to

the level of unstimulated condition after 5 minutes. Rictor depletion also reduced Rac

activation under adherent conditions (Figure 3.8C, D).

Recently, Liu and colleagues reported that mTORC2 negatively regulated RhoA

through activation of adenylyl cyclase 9 (AC9) (Liu et al., 2010). Rictor depletion caused

the inhibition of cAMP production and resulted in elevated RhoA activation. In

suspended cells, we also observed elevated content of RhoA-GTP in 1 minute after fMLP

stimulation (Figure 3.9A). However, under adherent conditions, no significant difference

of RhoA activation was observed between control and Rictor-depleted cells (Figure

3.9B). Myosin phosphorylation (MLRC p-S19) has been shown as the downstream event

of activated RhoA. In Rictor-depleted cells, we observed increased staining of myosin

phosphorylation in the back region of the cell body, compared to that of control.

Incubation with Y27632, an inhibitor for ROCK, completely abolished phosphorylated

95

Myosin light chain staining in cells (Figure 3.9C). In conclusion, although Rictor

depletion affected the backness pathways, the failure of polarization in Rictor-depleted

cells strongly indicates that the chemotactic defect was mainly due to inhibition of actin

polymerization.

3.5 Discussion

Previous studies have established the key roles of TOR signaling in cell growth

(Wullschleger et al., 2006). At least two TOR signaling branches have been identified.

The TORC1 signaling branch, which is sensitive to rapamycin, regulates the temporal

aspect of cell growth and has been intensively studied over the past decades. In contrast,

the TORC2 signaling branch, insensitive to rapamycin, is not well understood. A line of

evidence indicates that TORC2 regulates actin cytoskeleton in budding yeast and

mammalian cell lines. Recently, TORC2 has been documented as an important regulator

of D. discoideum chemotaxis in response to cAMP (Cai et al., 2010; Lee et al., 2005; Liu

et al., 2010). In this study, we found that mTORC2 was required for fMLP-induced

neutrophil chemotaxis. Based on our findings, a model has been proposed to explain the

function of mTORC2 in neutrophil chemotaxis. As described in Figure 3.10, mTORC2

acts downstream of activated Gi protein and regulates actin polymerization and

neutrophil chemotaxis through controlling Rac activation. This mTORC2-mediated

regulation is independent of AKT activation and mTOR kinase activity. First, inhibiton

of AKT and mTOR kinase activity failed to recapitulate the defect of chemotaxis

observed in mTORC2-depleted cells. Second, mTORC2 depletion profoundly impairs

chemoattractant-induced actin polymerization and neutrophil polarity. Third, we found

mTORC2 depletion inhibited the activation of Rac proteins, essential regulators of actin

96

cytoskeleton in neutrophils. Our findings further confirm and expand the role of

mTORC2 in organization of actin cytoskeleton in neutrophils.

How might mTORC2 regulate actin cytoskeleton during chemotaxis? In D.

discoideum, TORC2 depletion reduced cell polarity and migration speed. This defect

seemed dependent on the inhibition of PKB/PKBR phosphorylation due to TORC2

knockout (Kamimura et al., 2008; Lee et al., 2005). Furthermore, Ras has been shown to

function upstream to control TORC2 activation and downstream events (Cai et al., 2010).

Interestingly, TORC2 deletion in D. discoideum only modestly affected filamentous actin

formation and the structure of the actin cytoskeleton. TORC2 depletion strongly impairs

the activation of adenylyl cyclase which is required for cAMP production. Our findings

differ from the above reports. Depletion of mTORC2 dramatically impaired chemotaxis

of dHL-60 cells. After stimulation by fMLP, most cells could not persistently protrude

and failed to form stable polarity in the absence of mTORC2. The downstream effector

AKT is probably not the mediator of mTORC2 in neutrophil chemotaxis, as we showed

that the inhibition of AKT failed to affect neutrophil chemotaxis (Figure 3.4).

Furthermore, inhibition of mTOR kinase activity did not affect neutrophil chemotaxis

either, further suggesting that kinase activity-indepenent function of mTORC2 regulates

actin cytoskeleton and neutrophil chemotaxis (Figure 3.5). Evidence suggests that kinase-

independent functions exist in TORC to control myogenesis (Erbay and Chen, 2001; Ge

et al., 2009). Taken together, mammalian cells like neutrophils might have acquired new

functions of mTORCs to modulate actin cytoskeleton and cell motility, which are

independent on mTOR kinase activities and the mTORC downstream effector AKT.

97

Very recently, Liu et al reported that mTORC2 controlled neutrophil chemotaxis

through cAMP and in a RhoA-dependent fashion (Liu et al., 2010). In their study, cells

with Rictor depletion failed to polarize, but had comparable filamentous actin formation

to control cells in reponse to chemoattractant. This is in contrast to our finding that

mTORC2 depletion dramatically impaired actin cytoskeleton even without

chemoattractants. It is worth noting that in HeLa cells and 3T3 fibroblasts, mTORC2

depletion affected actin polymerization and dramatically altered the structure of actin

cytoskeleton characterized by the disappearance of stress fibers. Our findings support a

conserved role of mTORC2 in the regulation of actin cytoskeleton among mammalian

cells. Furthermore, the phenotypic defect of Rictor depletion was very different from the

adenylyl cyclase 9-depleted cells which developed long tails in response to

chemoattractants. Based on our findings, a major role of mTORC2 is in the regulation of

actin polymerization and thus is involved in leading edge formation. It is possible that the

defect in the leading edge of mTORC2-depleted cells might affect the contractility

machinery in the back, as indicated by the increase of RhoA and myosin II activation

(Figure 3.10). These discrepancies may be due to the differences in cell lines used in

these experiments. Further studies are needed to clarify these discrepancies of mTORC2

function in neutrophil chemotaxis.

Similar to NIH 3T3 cells, Rac proteins have been identified as the downstream

regulator of mTORC2 in neutrophil chemotaxis. Studies with knockout mouse and

human primary neutrophils with dominant negative Rac mutant indicate that Rac proteins

are the major regulators of actin cytoskeleton. In mTORC2 depleted cells, reduced Rac

activation was observed after fMLP stimulation. How mTORC2 regulates Rac activation

98

is an interesting question for future studies. In neutrophils, DOCK2 and p-Rex have been

identified as two nucleotide exchange factors for Rac activation. It is intriguing to

determine whether one or both GEFs are regulated by mTORC2. However, the severity

of chemotaxis in mTORC2-depleted cells suggests that there may be other additional

mechanisms used by mTORC2 to regulate actin cytoskeleton.

99

3.6 Figures and table

Neutr

ophil

dH

L-6

0

uH

L-6

0

mTOR

Rictor

Raptor

GAPDH

NS

shR

NA

mT

OR

shR

NA

-1

mT

OR

shR

NA

-2

Rap

tor

shR

NA

-1

Rapto

rshR

NA

-2

Ric

tor

shR

NA

-1

Ric

tor

shR

NA

-2

shRNAs:

mTOR

Rictor

Raptor

GAPDH

mT

OR

shR

NA

-3

Pro

tein

le

ve

l (%

)

0

20

40

60

80

100

120

Figure 3.1 Expression and depletion of mTOR, Rictor and Raptor. (A) mTOR, Rictor and

Raptor are expressed in neutrophil-like cell line HL-60 cells and human primary

neutrophils. Human neutrophils, undifferentiated HL-60 cells (uHL-60) and differentiated

HL-60 cells (dHL-60) were lysed in RIPA buffer and analyzed by western blotting.

GAPDH is the loading control. (B) mTORC depletion in dHL-60 cells. HL-60 cells were

infected with lenti-viral particles containing respective shRNAs overnight and then

induced to differentiate for 5 days. Cell lysates were analyzed by western blotting.

GAPDH is the loading control. (B) The knockdown efficiency of mTOR, Raptor or Rictor

by respective shRNAs. The Y axis represents the densitometry of remaining protein after

knockdown. Values are means SEM (n = 3)

A B

C

100

fM LP

fM LP

10s 10s 10s 10s

910s 910s 910s 910s

NT shRN A m TOR shRN A -3 Rictor shRNA-1 Raptor shRNA-1

NT shRNA mTOR shRNA-3 Rictor shRNA-1 Raptor shRNA-1

A

B

C

Figure 3.2

101

Ce

ll m

igra

tio

n (

%)

0

10

20

30

40

50

60

70

80

90

NT shRNA mTOR shRNA-3

Rictor shRNA-1 Raptor shRNA-1

5 s 307 s 5 s 287 s

5 s 406 s 5 s 297 s

D

E

Figure 3.2 (cont.) mTORC2 depletion impairs dHL-60 chemotaxis. (A) Chemotaxis of control

dHL-60 cells (NT shRNA) or dHL-60 cells with depletion of mTOR, Rictor or Raptor in microfluidic

chamber. Cells (2 106) were loaded to fibrinogen – coated microfluid device chamber for 20

minutes and exposed to an fMLP gradient (20 minutes) generated by the microfluidic device. As

shown are the images for cells at two time points in each condition. Bar, 50 µm. (B) The enlarged

view of cells after the stimulation of the fMLP gradient. Bar, 10 µm. (C) The outlines of cells

responding to the fMLP stimulation in microfluidic chamber. Each set of outlines represents a

single cell observed at indicated intervals (denoted by different colors) after exposure to fMLP.

Two representative cells were shown for each condition. (D) Quantification of chemotactic dHL-60

cells with or without mTORC depletion in microfluid device. During the experiments, cells within

the 200 µm region of the lowest fMLP concentration (bottom) were examined and the percentage

of cell migration is calculated as the ratio of chemotactic cells vs total cells. Values are means

SEM (n = 3). (E) Cell migration in response to the point source of fMLP. Cells treated with non-

target shRNA, mTOR shRNA-3, Rictor shRNA-1 or Raptor shRNA-1 were plated on fibrinogen

coated slides for 20 minutes and then stimulated by fMLP delivered from a micropipette. The two

time lapse images for each condition were shown. The bar represents 10 µm.

102

FITC-H

PI-

H

100

101

102

103

104

100

101

102

103

104

0.18% 1.77%

88.74%

9.31%

FITC-H

PI-

H

100

101

102

103

104

100

101

102

103

104

0.54% 3.05%

87.47%

8.95%

FITC-H

PI-

H

100

101

102

103

104

100

101

102

103

104

3.38% 6.00%

77.91%

12.71%

NT shRNA Rictor shRNA-1

mTOR shRNA-3

APC-A

Count

100

101

102

103

104

0

15

30

45

60

NT shRNAIsotype IgG

Rictor shRNA-1 mTOR shRNA-3

A

B

Figure 3.3 Rictor depletion did not affect HL-60 survival and differentiation. (A)

Apoptotic assay of control cells or mTORC2 depleted cells. Cells were

infected with lenti-viral particles containg non-target shRNA, mTOR shRNA-3

or Rictor shRNA-1 and differentiated for 5 days. Cells were prepared

according to the manual (FITC Annexin V Apoptosis Detection Kit I, BD

Pharmingen) and analyzed by flow cytometry. (C) Flow cytometry analysis of

surface expression of β2 integrin in control and mTORC2 depleted cells after

fMLP stimulation (100 nM, 10 min). The black line indicates background

fluorescence from the isotype control IgG.

103

0 0.5 1 2 3 5 10Time (min)

fMLP (100 nM)

p-S473 AKT

AKT

0

20

40

60

80

100

120

0 30 60 120 180 300 600

Re

lative

le

ve

l o

f p

-AK

T

fMLP stimulation (s)

p-S473 AKT

AKT

Rictor

NT shRNA Rictor shRNA-1

fMLP stim. (min) 0 1 5 0 1 5

A

B

C

Figure 3.4

104

Re

lative

le

ve

l o

f p

-AK

T

0

20

40

60

80

100

120

NT shRNA Rictor shRNA-1

AKT

p-S473 AKT

p-T308 AKT

un

stim

.

0 0.1 0.5 1 2 5 10AKTi -1/2 (μM)

fMLP (100 nM)

5s 905s

fMLP

AKTi -1/2 (1 µM)D

E

F

Figure 3.4 (cont.) The downstream effector AKT is not involved in mTORC2

mediated regulation of neutrophil chemotaxis. (A) Phosphorylation of AKT (p-S473)

in response to fMLP stimulation (100 nM) for various time points. (B) The average

level of AKT phosphorylation (p-S473) at various time points after fMLP stimulation.

Values are means SEM (n = 3). (C) Rictor depletion inhibits AKT phosphorylation

(p-S473) induced by fMLP. Total AKT is the loading control, and Rictor blot is also

shown. (D) Quantification of AKT phosphorylation (p-S473) in dHL-60 cells with or

without Rictor depletion at the time point of 1 minute. Values are means SEM (n =

3). (E) AKTi 1/2 inhibits fMLP induced AKT phosphorylation. Cells were treated with

various concentrations of AKTi 1/2 for 30 minutes and stimulated with 100 nM fMLP

for 1 minute. The level of AKT phosphorylation, p-S473 or p-T308, were assessed by

western blotting. (F) The chemotaxis of dHL-60 cells treated with AKTi 1/2 (1µM) in

micorfluid device. Bar, 50 µm.

105

0 1 5

Vehicle Rapamycin

0 1 5 0 1 5

Torin 1

fMLP (min)

p-S473 AKT

p-T389 S6K1

AKT

fmLP (min) 0 1 5 0 1 5 0 1 5

Vehicle + + + - - - - - -

Rapamycin - - - + + + - - -

Torin 1 - - - - - - + + +

0

20

40

60

80

100

120

Re

lative

le

ve

l o

f p

-AK

T

A

B

Figure 3.5

106

fmLP (min) 0 1 5 0 1 5 0 1 5

Vehicle + + + - - - - - -

Rapamycin - - - + + + - - -

Torin 1 - - - - - - + + +

0

20

40

60

80

100

120

Re

lative

le

ve

l o

f p

-S6

K1

Vehicle Rapamycin T o r in 1

10s

489s

10s

483s

10s

480s

fMLP

fMLP

C

D

Figure 3.5 (cont.)

107

flowflow flow

gradientgradient gradient

Vehicle Rapamycin Torin 1

Figure 3.5 (cont.) mTORC inhibition by rapamycin or torin 1 failed to affect

neutrophil chemotaxis. (A) mTORC inhibition by rapamycin or torin 1. dHL-60 cells

were pretreated with vehicle (DMSO), rapamycin (100 nM) or torin1 (250 nM) for 30

minutes and stimulated with fMLP (100 nM) for indicated time points.

Phosphorylation of AKT (p-S473) and S6K1 (p-T389) was analyzed by western

blotting. AKT is the loading control. (B) Quantification of AKT phosphorylation (p-

S473) in dHL-60 cells with or without mTORC inhibition. The values are normalized

to the peak level of AKT phosphorylation (1 minute). Values are means SEM (n =

3). (C) Quantification of S6K1 phosphorylation (p-T389) in in dHL-60 cells with or

without mTORC inhibition. The values are normalized to the peak level of S6K1

phosphorylation (5 minutes). Values are means SEM (n = 3). (D) chemotaxis of

dHL-60 treated with vehicle, rapamycin (100 nM) or torin 1(250 nM). Cells were pre-

treated with above reagents for 30 minutes and then loaded to the fibrinogen-coated

microfluidic device. The chemotactic cells in each condition were monitored by time-

lapse microscopy. As shown are the images for cells at two time points in each

condition. Bar, 50 µm. (E) trajectory of dHL-60 cells. dHL-60 cells were pretreated

with vehicle (DMSO), rapamycin (100 nM) or torin 1(250 nM) for 30 minutes and

then plated on fibrinogen coated microfluid device chamber. Cell migration was

recorded by time lapse microscope. The migration path of individual cells was

analyzed by Image J and plotted.

E

108

Cells Parameters DMSO (veh.) Rapamycin Torin 1

dHL-60 Speed

(μm/min)

4.36± 0.16 4.39± 0.12 4.67± 0.13

Chemotaxis

index

0.97± 0.01 0.95± 0.01 0.96± 0.01

PMN Speed

(μm/min)

7.05± 0.12 7.40± 0.55 7.47± 0.21

Chemotaxis

index

0.96± 0.01 0.97± 0.01 0.96± 0.01

Table 1 Effects of drug inhibition on dHL-60 and PMN chemotaxis

109

0

20

40

60

80

100

120

0 1 3 4 6 7 9 101113141617192022

Rictor

F-actin

micron

flu

ore

scen

ce in

ten

sit

y

-fMLP

0

20

40

60

80

100

120

140

160

180

200

0 1 3 4 6 7 9 10111314161719202223

Rictor

F-actin

micron

flu

ore

scen

ce in

ten

sit

y

+fMLP

A

B

Rictor F-actin Merge DIC

-fM

LP

+fM

LP

+fM

LP

+B

loc

kin

g p

ep

Figure 3.6

110

0

1

2

3

4

5

6

7

Front Back

F-actin

Rictor

Rela

tive M

FI

C

Figure 3.6 (cont.) mTORC2 accumulates in the leading edge of chemotactic

dHL-60 cells. (A) Immunofluorescence of Rictor (red) and F-actin (green) in

cells plated on fibrinogen without or with fMLP stimulation (100 nM, 2 minutes).

Fluorescent images of Rictor (red), F-actin (green), Rictor/F-actin merged

images and DIC images of cells are shown. Also shown is the staining of Rictor

and F-actin in the presence of anti-Rictor blocking peptide for fMLP treated

cells. Bar, 10 µm. (B) The fluorescence line profile of Rictor (Red) and F-actin

(Green) in stained dHL-60 cells. The fluorescence intensity cross the center of

cell body (unstimulated cell) or from the front to back of cell body (stimulated

cell) was measured with Image J and plotted. (C) The relative fluorescence

intensity of Rictor and F-actin in the leading edge of fMLP-stimulated dHL-60

cells. The first 4 µm from the edge of front is designated as “front” and the left

of cell body is defined as “back”. The ratio of fluorescence intensity for both F-

actin and Rictor (front vs back) was measured by Image J. As shown is the

result from 41 stained cells from at least 3 independent experiments.

111

fMLP stimulation (s)

Re

lative

F-a

ctin

0

0.5

1

1.5

2

2.5

3

0 30 60 180

NT shRNA

Rictor shRNA-1

Rictor shRNA-2

DICF-actin

NT

sh

RN

AR

icto

r sh

RN

A-1

Ric

tor

sh

RN

A-2

NT

shRNARictor

shRNA-1

Rictor

shRNA-2

po

lari

za

tio

n (

%)

0

10

20

30

40

50

60

70

80

90A

B

C

Figure 3.7

112

5s 30s 60 s 240 s

5s 30s 60 s 240 s

Actin

-YF

PD

ICA

ctin

-YF

PD

IC

NT

sh

RN

AR

icto

r sh

RN

A-1

D

Figure 3.7 (cont.) mTORC2 depletion impairs actin polymerization in chemotactic dHL-60

cells. (A) The content of F-actin in dHL-60 cells. dHL-60 cells with or without Rictor depletion

were stimulated with fMLP (100 nM) for indicated time points. Cells were immediately fixed,

permeablized and stained by fluorescence conjugated phalloidin. The binding of phalloidin to

the cells was analyzed by flow cytometry. As shown are the values from four independent

experiments. Means s.e.m. (B) F-actin staining in fMLP stimulated dHL-60 cells with or

without Rictor depletion. Cells were plated on fibrinogen-coated slide for 20 minutes and

stimulated with uniform fMLP (100 nM) for 2 minutes. Cells were immediately fixed,

permeablized and stained by fluorescence conjugated phalloidin. The images of F-actin and

DIC are shown. Bar,10 µm. (C) Quantification of palorized cells in response to uniform fMLP

(100 nM). Cells were treated as described in (B). Cells were examined from at least three

independent experiments: non-targeted shRNA (252 cells), Rictor shRNA-1 (255 cells) and

Rictor shRNA-2 (304 cells). (D) Actin-YFP dynamics in live dHL-60 cells. Actin-YFP stable

expressing cells were treated with non-target shRNA or Rictor shRNA-1 and differentiated for

5 days. Cells were plated on fibrinogen-coated slides and stimulated by a point source of

fMLP delivered by micropipette. The four images in each row show the positions of individual

cells after exposure to fMLP. all bars represent 10 µm.

113

Rac-GTP

Total Rac

fMLP stm. (s) 0 30 60 300 0 30 60 300

NT shRNA Rictor shRNA-1

suspension

0

20

40

60

80

100

120

NT shRNA Rictor shRNA-1

Re

lative

R

ac-G

TP

*

Rac-GTP

Total Rac

fMLP stim. (min) 0 1 5 0 1 5

NT shRNA Rictor shRNA-1

adhesion

A B

CD

Figure 3.8 mTORC2 depletion inhibits fMLP induced Rac activation. (A) Rac-GTP

pulldown in dHL-60 cells with or without Rictor depletion under suspension condition.

cells were stimulated with 100 nM of fMLP for indicated time points and lysed for the pull-

down assay. Levels of total Rap1 was used to show equal cell lysate input. (B)

Quantification of relative levels of Rac-GTP level in dHL-60 cells with and without Rictor

depletion 30 seconds after fMP stimulation. Each bar represents the mean SEM (error

bars). Values are normalized to the level of Rac-GTP (=1) in cells without Rictor

depletion. Asterisks indicate that the value for cells with Rictor depletion differs

statistically from the control (*, p<0.001, n=3). (C) The levels of Rac-GTP in adherent

dHL-60 cells under various conditions. Control cells or Rictor-depleted cells plated on

fibrinogen-coated plates were unstimulated or stimulated for 1 or 5 minutes with a

uniform concentration of fMLP (100 nM) and lysed for pull-down assay. Levels of total

Rac was used to show cell lysate input. (D) Quantification of relative levels of Rac-GTP

level in experiments described in (C). Values are normalized to the level of Rac-GTP (=1)

in cells without Rictor depletion stimulated with fMLP. Each bar represents the

mean SEM (error bars) (n=3).

114

Re

lative

Rh

oA

-GT

P

fMLP stimulation (s)

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

1.8

0 30 60 300

NT shRNA

Rictor shRNA-1

suspension*

0

0.2

0.4

0.6

0.8

1

1.2

1.4

0 60 300

NT shRNA

Rictor shRNA-1

Re

lative

Rh

oA

-GT

P

fMLP stimulation (s)

adhesion

NT

sh

RN

AR

icto

r sh

RN

A-1

NT

sh

RN

A

+Y

27

63

2

F-actin

p-MRLCDICA

B

C

Figure 3.9 mTORC2 depletion enhance fMLP induced RhoA activation and myosin

phosphorylation. (A) Quantification of RhoA-GTP in dHL-60 cells with or with Rictor

depletion under suspension condition. Cells were stimulated with fMLP (100 nM) for

times indicated and lysed. The level of RhoA-GTP was determined by the use of an

absorbance-based RhoA G-LISA kit. The y axis represents the absorbance at 490

nm. Each bar represents the mean s.e.m. (error bars) (n=4) (B) Quantification of

RhoA-GTP dHL-60 cells with or with Rictor depletion under adherent condition. Cells

were suspended in mHBSS buffer and plated on fibrinogen-coated plates for 30

minutes. Cells were then stimulated by 100 nM fMLP for indicated times and lysed.

The level of RhoA-GTP was determined by the use of an absorbance-based RhoA G-

LISA kit. The y axis represents the absorbance at 490 nm. Each bar represents the

mean s.e.m. (error bars) (n=3). (C) Merged fluorescent images of p-MLC (red) and

F-actin (green) in control, Rictor-depleted cells and cells pretreated with Y-27632 (30

μM, 30 min). The corresponding DIC images are shown. The white arrow indicates

the loss of p-MLC fluorescence at the back of the Y-27632-treated cell. Bars, 10 μm.

115

mTORC2

AKTRac

F-actin

RhoA

Myosin II

GPCR

Figure 3.10 The current model of mTORC2 in neutrophil chemotaxis. mTORC2 is

activated by Gi protein and controls Rac activation and actin polymerization through

the fashion of AKT phosphorylation –independent and kinase activity-independent

pathways. Concomitantly, mTORC2 down-regulates RhoA activation, as shown

increased myosin phosphorylation.

116

CHAPTER 4 CONCLUSION

This dissertation is the summary of my research work conducted in Dr. Fei

Wang’s laboratory during my graduate studies at the University of Illinois at Urbana-

Champaign. My major research goal was to dissect the molecular mechanism of signal

transduction during neutrophil chemotaxis. As described in previous chapters, neutrophil

chemotaxis, as well as other types of cell migration, is a complex process regulated by

the integration of multiple signaling pathways. I have worked on two research projects:

the signaling mechanisms regulating interaction between chemotactic neutrophils and

substrate, and the mechanisms of actin cytoskeleton dynamics. By using the genetically

tractable cell line (dHL-60), we have documented a novel signaling pathway by which

non-receptor tyrosine kinase Lyn controls leading edge adhesion. We have also found

that mTORC2 is an essential regulator of actin polymerization during neutrophil

chemotaxis.

In Chapter 2, I have described the function of Lyn kinase in neutrophil

chemotaxis. Although Lyn kinase has been shown to be activated by many cell surface

receptors, the molecular details of Lyn kinase function in neutrophil chemotaxis has not

been fully explored. The data presented in this chapter show that Lyn kinase was

activated and translocated to the leading edge upon fMLP stimulation. Using the

approach of RNA interference, we demonstrated that Lyn kinase was required for the

persistent migration of dHL-60 cells in the migration settings of micropipette and

microfluid device. Lyn depletion did not affect the signaling pathways regulating PI3K,

Rac activation and actin polymerization. Although Lyn-depleted cells had a similar

phenotype to cells with defects in the backness pathways, Lyn depletion did not affect

117

RhoA activation and myosin phosphorylation. Furthermore, we observed that Lyn

depletion reduced the accumulation of active β2 integrin in the leading edge without

altering the total β2 distribution. Interestingly, blocking de novo adhesion by β2 antibody

recapitulated the defects observed in Lyn-depleted cells. At the molecular level, Lyn

depletion inhibited fMLP-induced Rap1 activation, a known upstream event of β2

integrin activation pathways. In Lyn-depleted cells, the level of active Rap1 was reduced

in the leading edge of live chemotactic cells when compared to that of control cells. We

further found that the reduced activation of Rap1 in Lyn-depleted cells corresponded to

the inhibition of CrKL/C3G recruitment to the leading edge. In summary, our data in

Chapter 2 have revealed a novel signaling pathway by which Lyn kinase regulates

leading edge adhesion during neutrophil chemotaxis. However, there are some

unanswered questions about how Lyn kinase regulates CrKL/C3G recruitment and, thus,

integrin activation.

In Chapter 3, I have presented the results on the role of mTORC2 on neutrophil

chemotaxis. Dynamics of the actin cytoskeleton are thought to be essential for neutrophil

chemotaxis as well as other types of cell migration. We report in this chapter that

mTORC2 disruption severely inhibited dHL-60 cell chemotaxis when cells were exposed

to a point source of fMLP by micropipette or a linear fMLP gradient by microfluid

device. This migratory defect was not due to inhibition of mTORC2 kinase activity, as

shown that inhibition of AKT and mTOR kinase did not affect chemotaxis of dHL-60

cells and neutrophils. mTORC2 depletion strongly inhibited actin polymerization induced

by fMLP in dHL-60 cells. When mTORC2-depleted cells were plated on fibrinogen

substrate, they failed to form the typical polarized morphology as seen in control cells. At

118

the molecular level, mTORC2 depletion inhibited Rac activation by fMLP stimulation.

How mTORC2 regulates Rac activation and actin polymerization are under investigation.

The answer to these questions would also shed light on the regulatory mechanism of

mTORC2 on actin cytoskeleton in other types of cells.

As mentioned previously, we have begun to unravel the complex signaling

mechanism underpinning neutrophil chemotaxis by using the transgenic mouse and

neutrophil-like cell lines. However, there are many remaining questions regarding how

neutrophils respond to attractants at the molecular and cellular levels, especially the

signaling cascades involved in the regulation of cell adhesion. In the future, more insights

on the integrated signaling pathways which temporally and spatially regulate neutrophil

chemotaxis will be obtained through the development of new methodologies and

technologies, such as the application of intravital microsopy to observe chemotaxing

neutrophils in the animal body.

119

REFERENCES

Altieri, D.C. 1991. Occupancy of CD11b/CD18 (Mac-1) divalent ion binding site(s)

induces leukocyte adhesion. J Immunol. 147:1891-1898.

Altieri, D.C., F.R. Agbanyo, J. Plescia, M.H. Ginsberg, T.S. Edgington, and E.F. Plow.

1990. A unique recognition site mediates the interaction of fibrinogen with the

leukocyte integrin Mac-1 (CD11b/CD18). J Biol Chem. 265:12119-12122.

Ambruso, D.R., C. Knall, A.N. Abell, J. Panepinto, A. Kurkchubasche, G. Thurman, C.

Gonzalez-Aller, A. Hiester, M. deBoer, R.J. Harbeck, R. Oyer, G.L. Johnson, and

D. Roos. 2000. Human neutrophil immunodeficiency syndrome is associated with

an inhibitory Rac2 mutation. Proc Natl Acad Sci U S A. 97:4654-4659.

Arai, A., Y. Nosaka, H. Kohsaka, N. Miyasaka, and O. Miura. 1999. CrkL activates

integrin-mediated hematopoietic cell adhesion through the guanine nucleotide

exchange factor C3G. Blood. 93:3713-3722.

Arcaro, A., and M.P. Wymann. 1993. Wortmannin is a potent phosphatidylinositol 3-

kinase inhibitor: the role of phosphatidylinositol 3,4,5-trisphosphate in neutrophil

responses. Biochem J. 296 ( Pt 2):297-301.

Avdi, N.J., B.W. Winston, M. Russel, S.K. Young, G.L. Johnson, and G.S. Worthen.

1996. Activation of MEKK by formyl-methionyl-leucyl-phenylalanine in human

neutrophils. Mapping pathways for mitogen-activated protein kinase activation. J

Biol Chem. 271:33598-33606.

Baggiolini, M., B. Dewald, and B. Moser. 1994. Interleukin-8 and related chemotactic

cytokines--CXC and CC chemokines. Adv Immunol. 55:97-179.

Ballestrem, C., B. Hinz, B.A. Imhof, and B. Wehrle-Haller. 2001. Marching at the front

and dragging behind: differential alphaVbeta3-integrin turnover regulates focal

adhesion behavior. J Cell Biol. 155:1319-1332.

Barnett, S.F., D. Defeo-Jones, S. Fu, P.J. Hancock, K.M. Haskell, R.E. Jones, J.A.

Kahana, A.M. Kral, K. Leander, L.L. Lee, J. Malinowski, E.M. McAvoy, D.D.

Nahas, R.G. Robinson, and H.E. Huber. 2005. Identification and characterization

of pleckstrin-homology-domain-dependent and isoenzyme-specific Akt inhibitors.

Biochem J. 385:399-408.

Baruzzi, A., E. Caveggion, and G. Berton. 2008. Regulation of phagocyte migration and

recruitment by Src-family kinases. Cell Mol Life Sci. 65:2175-2190.

Benard, V., B.P. Bohl, and G.M. Bokoch. 1999. Characterization of rac and cdc42

activation in chemoattractant-stimulated human neutrophils using a novel assay

for active GTPases. J Biol Chem. 274:13198-13204.

Beningo, K.A., M. Dembo, I. Kaverina, J.V. Small, and Y.L. Wang. 2001. Nascent focal

adhesions are responsible for the generation of strong propulsive forces in

migrating fibroblasts. J Cell Biol. 153:881-888.

Berton, G., and C.A. Lowell. 1999. Integrin signalling in neutrophils and macrophages.

Cell Signal. 11:621-635.

Bivona, T.G., H.H. Wiener, I.M. Ahearn, J. Silletti, V.K. Chiu, and M.R. Philips. 2004.

Rap1 up-regulation and activation on plasma membrane regulates T cell adhesion.

J Cell Biol. 164:461-470.

120

Bodin, S., and M.D. Welch. 2005. Plasma membrane organization is essential for

balancing competing pseudopod- and uropod-promoting signals during neutrophil

polarization and migration. Mol Biol Cell. 16:5773-5783.

Bokoch, G.M., B.P. Bohl, and T.H. Chuang. 1994. Guanine nucleotide exchange

regulates membrane translocation of Rac/Rho GTP-binding proteins. J Biol

Chem. 269:31674-31679.

Bos, J.L. 2005. Linking Rap to cell adhesion. Curr Opin Cell Biol. 17:123-128.

Bourne, H.R., and O. Weiner. 2002. A chemical compass. Nature. 419:21.

Bunting, M., E.S. Harris, T.M. McIntyre, S.M. Prescott, and G.A. Zimmerman. 2002.

Leukocyte adhesion deficiency syndromes: adhesion and tethering defects

involving beta 2 integrins and selectin ligands. Curr Opin Hematol. 9:30-35.

Burridge, K., and M. Chrzanowska-Wodnicka. 1996. Focal adhesions, contractility, and

signaling. Annu Rev Cell Dev Biol. 12:463-518.

Cai, H., S. Das, Y. Kamimura, Y. Long, C.A. Parent, and P.N. Devreotes. 2010. Ras-

mediated activation of the TORC2-PKB pathway is critical for chemotaxis. J Cell

Biol. 190:233-245.

Caron, E., A.J. Self, and A. Hall. 2000. The GTPase Rap1 controls functional activation

of macrophage integrin alphaMbeta2 by LPS and other inflammatory mediators.

Curr Biol. 10:974-978.

Carp, H. 1982. Mitochondrial N-formylmethionyl proteins as chemoattractants for

neutrophils. J Exp Med. 155:264-275.

Cockcroft, S. 1992. G-protein-regulated phospholipases C, D and A2-mediated signalling

in neutrophils. Biochim Biophys Acta. 1113:135-160.

Coffer, P.J., N. Geijsen, L. M'Rabet, R.C. Schweizer, T. Maikoe, J.A. Raaijmakers, J.W.

Lammers, and L. Koenderman. 1998. Comparison of the roles of mitogen-

activated protein kinase kinase and phosphatidylinositol 3-kinase signal

transduction in neutrophil effector function. Biochem J. 329 ( Pt 1):121-130.

Corey, S.J., P.M. Dombrosky-Ferlan, S. Zuo, E. Krohn, A.D. Donnenberg, P. Zorich, G.

Romero, M. Takata, and T. Kurosaki. 1998. Requirement of Src kinase Lyn for

induction of DNA synthesis by granulocyte colony-stimulating factor. J Biol

Chem. 273:3230-3235.

Davies, S.P., H. Reddy, M. Caivano, and P. Cohen. 2000. Specificity and mechanism of

action of some commonly used protein kinase inhibitors. Biochem J. 351:95-105.

de Jong, R., J. ten Hoeve, N. Heisterkamp, and J. Groffen. 1997. Tyrosine 207 in CRKL

is the BCR/ABL phosphorylation site. Oncogene. 14:507-513.

Devreotes, P.N., and S.H. Zigmond. 1988. Chemotaxis in eukaryotic cells: a focus on

leukocytes and Dictyostelium. Annu Rev Cell Biol. 4:649-686.

Duchniewicz, M., T. Zemojtel, M. Kolanczyk, S. Grossmann, J.S. Scheele, and F.J.

Zwartkruis. 2006. Rap1A-deficient T and B cells show impaired integrin-

mediated cell adhesion. Mol Cell Biol. 26:643-653.

Dusi, S., M. Donini, F. Wientjes, and F. Rossi. 1996. Tyrosine phosphorylation and

subcellular redistribution of p125 ras guanosine triphosphatase-activating protein

in human neutrophils stimulated with FMLP. FEBS Lett. 383:181-184.

Ehrengruber, M.U., P. Boquet, T.D. Coates, and D.A. Deranleau. 1995a. ADP-

ribosylation of Rho enhances actin polymerization-coupled shape oscillations in

human neutrophils. FEBS Lett. 372:161-164.

121

Ehrengruber, M.U., T.D. Coates, and D.A. Deranleau. 1995b. Shape oscillations: a

fundamental response of human neutrophils stimulated by chemotactic peptides?

FEBS Lett. 359:229-232.

Erbay, E., and J. Chen. 2001. The mammalian target of rapamycin regulates C2C12

myogenesis via a kinase-independent mechanism. J Biol Chem. 276:36079-

36082.

Etienne-Manneville, S., and A. Hall. 2002. Rho GTPases in cell biology. Nature.

420:629-635.

Fechheimer, M., and S.H. Zigmond. 1983. Changes in cytoskeletal proteins of

polymorphonuclear leukocytes induced by chemotactic peptides. Cell Motil.

3:349-361.

Firtel, R.A., and R. Meili. 2000. Dictyostelium: a model for regulated cell movement

during morphogenesis. Curr Opin Genet Dev. 10:421-427.

Francis, K., B. Palsson, J. Donahue, S. Fong, and E. Carrier. 2002. Murine Sca-1(+)/Lin(-

) cells and human KG1a cells exhibit multiple pseudopod morphologies during

migration. Exp Hematol. 30:460-463.

Friedl, P. 2004. Prespecification and plasticity: shifting mechanisms of cell migration.

Curr Opin Cell Biol. 16:14-23.

Friedl, P., S. Borgmann, and E.B. Brocker. 2001. Amoeboid leukocyte crawling through

extracellular matrix: lessons from the Dictyostelium paradigm of cell movement.

J Leukoc Biol. 70:491-509.

Friedl, P., K.S. Zanker, and E.B. Brocker. 1998. Cell migration strategies in 3-D

extracellular matrix: differences in morphology, cell matrix interactions, and

integrin function. Microsc Res Tech. 43:369-378.

Fumagalli, L., H. Zhang, A. Baruzzi, C.A. Lowell, and G. Berton. 2007. The Src family

kinases Hck and Fgr regulate neutrophil responses to N-formyl-methionyl-leucyl-

phenylalanine. J Immunol. 178:3874-3885.

Funk, C.D. 2001. Prostaglandins and leukotrienes: advances in eicosanoid biology.

Science. 294:1871-1875.

Gakidis, M.A., X. Cullere, T. Olson, J.L. Wilsbacher, B. Zhang, S.L. Moores, K. Ley, W.

Swat, T. Mayadas, and J.S. Brugge. 2004. Vav GEFs are required for beta2

integrin-dependent functions of neutrophils. J Cell Biol. 166:273-282.

Gaudry, M., A.C. Caon, C. Gilbert, S. Lille, and P.H. Naccache. 1992. Evidence for the

involvement of tyrosine kinases in the locomotory responses of human

neutrophils. J Leukoc Biol. 51:103-108.

Gaudry, M., C. Gilbert, F. Barabe, P.E. Poubelle, and P.H. Naccache. 1995. Activation of

Lyn is a common element of the stimulation of human neutrophils by soluble and

particulate agonists. Blood. 86:3567-3574.

Ge, Y., A.L. Wu, C. Warnes, J. Liu, C. Zhang, H. Kawasome, N. Terada, M.D. Boppart,

C.J. Schoenherr, and J. Chen. 2009. mTOR regulates skeletal muscle regeneration

in vivo through kinase-dependent and kinase-independent mechanisms. Am J

Physiol Cell Physiol. 297:C1434-1444.

Gerard, C., and N.P. Gerard. 1994. The pro-inflammatory seven-transmembrane segment

receptors of the leukocyte. Curr Opin Immunol. 6:140-145.

Gilbert, S.F. 2010. Developmental Biology. Sinauer, Sunderland, MA.

122

Glogauer, M., J. Hartwig, and T. Stossel. 2000. Two pathways through Cdc42 couple the

N-formyl receptor to actin nucleation in permeabilized human neutrophils. J Cell

Biol. 150:785-796.

Gomez-Mouton, C., R.A. Lacalle, E. Mira, S. Jimenez-Baranda, D.F. Barber, A.C.

Carrera, A.C. Martinez, and S. Manes. 2004. Dynamic redistribution of raft

domains as an organizing platform for signaling during cell chemotaxis. J Cell

Biol. 164:759-768.

Grinnell, F. 1994. Fibroblasts, myofibroblasts, and wound contraction. J Cell Biol.

124:401-404.

Guertin, D.A., D.M. Stevens, C.C. Thoreen, A.A. Burds, N.Y. Kalaany, J. Moffat, M.

Brown, K.J. Fitzgerald, and D.M. Sabatini. 2006. Ablation in mice of the mTORC

components raptor, rictor, or mLST8 reveals that mTORC2 is required for

signaling to Akt-FOXO and PKCalpha, but not S6K1. Dev Cell. 11:859-871.

Haston, W.S., and J.M. Shields. 1985. Neutrophil leucocyte chemotaxis: a simplified

assay for measuring polarizing responses to chemotactic factors. J Immunol

Methods. 81:229-237.

Hauert, A.B., S. Martinelli, C. Marone, and V. Niggli. 2002. Differentiated HL-60 cells

are a valid model system for the analysis of human neutrophil migration and

chemotaxis. Int J Biochem Cell Biol. 34:838-854.

Hegerfeldt, Y., M. Tusch, E.B. Brocker, and P. Friedl. 2002. Collective cell movement in

primary melanoma explants: plasticity of cell-cell interaction, beta1-integrin

function, and migration strategies. Cancer Res. 62:2125-2130.

Herzmark, P., K. Campbell, F. Wang, K. Wong, H. El-Samad, A. Groisman, and H.R.

Bourne. 2007. Bound attractant at the leading vs. the trailing edge determines

chemotactic prowess. Proc Natl Acad Sci U S A. 104:13349-13354.

Hirsch, E., V.L. Katanaev, C. Garlanda, O. Azzolino, L. Pirola, L. Silengo, S. Sozzani, A.

Mantovani, F. Altruda, and M.P. Wymann. 2000. Central role for G protein-

coupled phosphoinositide 3-kinase gamma in inflammation. Science. 287:1049-

1053.

Hunter, T. 1987. A tail of two src's: mutatis mutandis. Cell. 49:1-4.

Huttenlocher, A., S.P. Palecek, Q. Lu, W. Zhang, R.L. Mellgren, D.A. Lauffenburger,

M.H. Ginsberg, and A.F. Horwitz. 1997. Regulation of cell migration by the

calcium-dependent protease calpain. J Biol Chem. 272:32719-32722.

Huttenlocher, A., R.R. Sandborg, and A.F. Horwitz. 1995. Adhesion in cell migration.

Curr Opin Cell Biol. 7:697-706.

Jacinto, E., and M.N. Hall. 2003. Tor signalling in bugs, brain and brawn. Nat Rev Mol

Cell Biol. 4:117-126.

Jacinto, E., R. Loewith, A. Schmidt, S. Lin, M.A. Ruegg, A. Hall, and M.N. Hall. 2004.

Mammalian TOR complex 2 controls the actin cytoskeleton and is rapamycin

insensitive. Nat Cell Biol. 6:1122-1128.

Jeon, N.J., S.K.W. Dertinger, D.T. Chiu, I.S. Choi, A.D. Stroock, and G.M. Whitesides.

2000. Generation of solution and surface gradients using microfluidic systems.

Langmuir. 16:8311-8316.

Johansson, M.W., M. Patarroyo, F. Oberg, A. Siegbahn, and K. Nilsson. 1997.

Myeloperoxidase mediates cell adhesion via the alpha M beta 2 integrin (Mac-1,

CD11b/CD18). J Cell Sci. 110 ( Pt 9):1133-1139.

123

Kamada, Y., Y. Fujioka, N.N. Suzuki, F. Inagaki, S. Wullschleger, R. Loewith, M.N.

Hall, and Y. Ohsumi. 2005. Tor2 directly phosphorylates the AGC kinase Ypk2 to

regulate actin polarization. Mol Cell Biol. 25:7239-7248.

Kamimura, Y., Y. Xiong, P.A. Iglesias, O. Hoeller, P. Bolourani, and P.N. Devreotes.

2008. PIP3-independent activation of TorC2 and PKB at the cell's leading edge

mediates chemotaxis. Curr Biol. 18:1034-1043.

Kawaguchi, A., M. Ohmori, K. Harada, S. Tsuruoka, K. Sugimoto, and A. Fujimura.

2000. The effect of a Rho kinase inhibitor Y-27632 on superoxide production,

aggregation and adhesion in human polymorphonuclear leukocytes. Eur J

Pharmacol. 403:203-208.

Kent, J.D., S. Sergeant, D.J. Burns, and L.C. McPhail. 1996. Identification and regulation

of protein kinase C-delta in human neutrophils. J Immunol. 157:4641-4647.

King, C.A., T.M. Preston, R.H. Miller, and P. Donovan. 1980. Cell-substrate interactions

during amoeboid locomotion of neutrophil leukocytes. Exp Cell Res. 126:453-

458.

Knall, C., G.S. Worthen, and G.L. Johnson. 1997. Interleukin 8-stimulated

phosphatidylinositol-3-kinase activity regulates the migration of human

neutrophils independent of extracellular signal-regulated kinase and p38 mitogen-

activated protein kinases. Proc Natl Acad Sci U S A. 94:3052-3057.

Knall, C., S. Young, J.A. Nick, A.M. Buhl, G.S. Worthen, and G.L. Johnson. 1996.

Interleukin-8 regulation of the Ras/Raf/mitogen-activated protein kinase pathway

in human neutrophils. J Biol Chem. 271:2832-2838.

Knaus, U.G., and G.M. Bokoch. 1998. The p21Rac/Cdc42-activated kinases (PAKs). Int

J Biochem Cell Biol. 30:857-862.

Korade-Mirnics, Z., and S.J. Corey. 2000. Src kinase-mediated signaling in leukocytes. J

Leukoc Biol. 68:603-613.

Kreienbuhl, P., H. Keller, and V. Niggli. 1992. Protein phosphatase inhibitors okadaic

acid and calyculin A alter cell shape and F-actin distribution and inhibit stimulus-

dependent increases in cytoskeletal actin of human neutrophils. Blood. 80:2911-

2919.

Krugmann, S., K.E. Anderson, S.H. Ridley, N. Risso, A. McGregor, J. Coadwell, K.

Davidson, A. Eguinoa, C.D. Ellson, P. Lipp, M. Manifava, N. Ktistakis, G.

Painter, J.W. Thuring, M.A. Cooper, Z.Y. Lim, A.B. Holmes, S.K. Dove, R.H.

Michell, A. Grewal, A. Nazarian, H. Erdjument-Bromage, P. Tempst, L.R.

Stephens, and P.T. Hawkins. 2002. Identification of ARAP3, a novel PI3K

effector regulating both Arf and Rho GTPases, by selective capture on

phosphoinositide affinity matrices. Mol Cell. 9:95-108.

Landry, J., and J. Huot. 1995. Modulation of actin dynamics during stress and

physiological stimulation by a signaling pathway involving p38 MAP kinase and

heat-shock protein 27. Biochem Cell Biol. 73:703-707.

Landry, J., and J. Huot. 1999. Regulation of actin dynamics by stress-activated protein

kinase 2 (SAPK2)-dependent phosphorylation of heat-shock protein of 27 kDa

(Hsp27). Biochem Soc Symp. 64:79-89.

Laudanna, C., D. Mochly-Rosen, T. Liron, G. Constantin, and E.C. Butcher. 1998.

Evidence of zeta protein kinase C involvement in polymorphonuclear neutrophil

integrin-dependent adhesion and chemotaxis. J Biol Chem. 273:30306-30315.

124

Lauffenburger, D.A., and A.F. Horwitz. 1996. Cell migration: a physically integrated

molecular process. Cell. 84:359-369.

Lawson, M.A., and F.R. Maxfield. 1995. Ca(2+)- and calcineurin-dependent recycling of

an integrin to the front of migrating neutrophils. Nature. 377:75-79.

Lee, S., F.I. Comer, A. Sasaki, I.X. McLeod, Y. Duong, K. Okumura, J.R. Yates, 3rd,

C.A. Parent, and R.A. Firtel. 2005. TOR complex 2 integrates cell movement

during chemotaxis and signal relay in Dictyostelium. Mol Biol Cell. 16:4572-

4583.

Ley, K., C. Laudanna, M.I. Cybulsky, and S. Nourshargh. 2007. Getting to the site of

inflammation: the leukocyte adhesion cascade updated. Nat Rev Immunol. 7:678-

689.

Li, Y., J. Yan, P. De, H.C. Chang, A. Yamauchi, K.W. Christopherson, 2nd, N.C.

Paranavitana, X. Peng, C. Kim, V. Munugalavadla, R. Kapur, H. Chen, W. Shou,

J.C. Stone, M.H. Kaplan, M.C. Dinauer, D.L. Durden, and L.A. Quilliam. 2007.

Rap1a null mice have altered myeloid cell functions suggesting distinct roles for

the closely related Rap1a and 1b proteins. J Immunol. 179:8322-8331.

Li, Z., H. Jiang, W. Xie, Z. Zhang, A.V. Smrcka, and D. Wu. 2000. Roles of PLC-beta2

and -beta3 and PI3Kgamma in chemoattractant-mediated signal transduction.

Science. 287:1046-1049.

Lin, F., and E.C. Butcher. 2006. T cell chemotaxis in a simple microfluidic device. Lab

Chip. 6:1462-1469.

Liu, L., S. Das, W. Losert, and C.A. Parent. 2010. mTORC2 regulates neutrophil

chemotaxis in a cAMP- and RhoA-dependent fashion. Dev Cell. 19:845-857.

Loewith, R., E. Jacinto, S. Wullschleger, A. Lorberg, J.L. Crespo, D. Bonenfant, W.

Oppliger, P. Jenoe, and M.N. Hall. 2002. Two TOR complexes, only one of

which is rapamycin sensitive, have distinct roles in cell growth control. Mol Cell.

10:457-468.

Lowell, C.A. 2004. Src-family kinases: rheostats of immune cell signaling. Mol Immunol.

41:631-643.

M'Rabet, L., P. Coffer, F. Zwartkruis, B. Franke, A.W. Segal, L. Koenderman, and J.L.

Bos. 1998. Activation of the small GTPase rap1 in human neutrophils. Blood.

92:2133-2140.

Machesky, L.M., and R.H. Insall. 1998. Scar1 and the related Wiskott-Aldrich syndrome

protein, WASP, regulate the actin cytoskeleton through the Arp2/3 complex. Curr

Biol. 8:1347-1356.

Machesky, L.M., R.D. Mullins, H.N. Higgs, D.A. Kaiser, L. Blanchoin, R.C. May, M.E.

Hall, and T.D. Pollard. 1999. Scar, a WASp-related protein, activates nucleation

of actin filaments by the Arp2/3 complex. Proc Natl Acad Sci U S A. 96:3739-

3744.

Marasco, W.A., S.H. Phan, H. Krutzsch, H.J. Showell, D.E. Feltner, R. Nairn, E.L.

Becker, and P.A. Ward. 1984. Purification and identification of formyl-

methionyl-leucyl-phenylalanine as the major peptide neutrophil chemotactic

factor produced by Escherichia coli. J Biol Chem. 259:5430-5439.

Martin, P., and S.J. Leibovich. 2005. Inflammatory cells during wound repair: the good,

the bad and the ugly. Trends Cell Biol. 15:599-607.

125

Maxwell, M.J., Y. Yuan, K.E. Anderson, M.L. Hibbs, H.H. Salem, and S.P. Jackson.

2004. SHIP1 and Lyn Kinase Negatively Regulate Integrin alpha IIb beta 3

signaling in platelets. J Biol Chem. 279:32196-32204.

McKay, D.A., J.R. Kusel, and P.C. Wilkinson. 1991. Studies of chemotactic factor-

induced polarity in human neutrophils. Lipid mobility, receptor distribution and

the time-sequence of polarization. J Cell Sci. 100 ( Pt 3):473-479.

Meili, R., C. Ellsworth, S. Lee, T.B. Reddy, H. Ma, and R.A. Firtel. 1999.

Chemoattractant-mediated transient activation and membrane localization of

Akt/PKB is required for efficient chemotaxis to cAMP in Dictyostelium. Embo J.

18:2092-2105.

Mocsai, A., M. Zhou, F. Meng, V.L. Tybulewicz, and C.A. Lowell. 2002. Syk is required

for integrin signaling in neutrophils. Immunity. 16:547-558.

Model, M.A., and G.M. Omann. 1996. Cell polarization as a possible mechanism of

response termination. Biochem Biophys Res Commun. 224:516-521.

Molski, T.F., and R.I. Sha'afi. 1987. Intracellular acidification, guanine-nucleotide

binding proteins, and cytoskeletal actin. Cell Motil Cytoskeleton. 8:1-6.

Murphy, P.M. 1994. The molecular biology of leukocyte chemoattractant receptors. Annu

Rev Immunol. 12:593-633.

Murphy, P.M. 1996. Chemokine receptors: structure, function and role in microbial

pathogenesis. Cytokine Growth Factor Rev. 7:47-64.

Naccache, P.H., S. Levasseur, G. Lachance, S. Chakravarti, S.G. Bourgoin, and S.R.

McColl. 2000. Stimulation of human neutrophils by chemotactic factors is

associated with the activation of phosphatidylinositol 3-kinase gamma. J Biol

Chem. 275:23636-23641.

Nakata, Y., B. Tomkowicz, A.M. Gewirtz, and A. Ptasznik. 2006. Integrin inhibition

through Lyn-dependent cross talk from CXCR4 chemokine receptors in normal

human CD34+ marrow cells. Blood. 107:4234-4239.

Neel, N.F., M. Barzik, D. Raman, T. Sobolik-Delmaire, J. Sai, A.J. Ham, R.L. Mernaugh,

F.B. Gertler, and A. Richmond. 2009. VASP is a CXCR2-interacting protein that

regulates CXCR2-mediated polarization and chemotaxis. J Cell Sci. 122:1882-

1894.

Neptune, E.R., T. Iiri, and H.R. Bourne. 1999. Galphai is not required for chemotaxis

mediated by Gi-coupled receptors. J Biol Chem. 274:2824-2828.

Nick, J.A., N.J. Avdi, S.K. Young, C. Knall, P. Gerwins, G.L. Johnson, and G.S.

Worthen. 1997. Common and distinct intracellular signaling pathways in human

neutrophils utilized by platelet activating factor and FMLP. J Clin Invest. 99:975-

986.

Niggli, V. 1999. Rho-kinase in human neutrophils: a role in signalling for myosin light

chain phosphorylation and cell migration. FEBS Lett. 445:69-72.

Niggli, V. 2000. A membrane-permeant ester of phosphatidylinositol 3,4, 5-trisphosphate

(PIP(3)) is an activator of human neutrophil migration. FEBS Lett. 473:217-221.

Niggli, V., and H. Keller. 1997. The phosphatidylinositol 3-kinase inhibitor wortmannin

markedly reduces chemotactic peptide-induced locomotion and increases in

cytoskeletal actin in human neutrophils. Eur J Pharmacol. 335:43-52.

126

Nijhuis, E., J.W. Lammers, L. Koenderman, and P.J. Coffer. 2002. Src kinases regulate

PKB activation and modulate cytokine and chemoattractant-controlled neutrophil

functioning. J Leukoc Biol. 71:115-124.

Nuzzi, P.A., M.A. Senetar, and A. Huttenlocher. 2007. Asymmetric localization of

calpain 2 during neutrophil chemotaxis. Mol Biol Cell. 18:795-805.

O'Laughlin-Bunner, B., N. Radosevic, M.L. Taylor, Shivakrupa, C. DeBerry, D.D.

Metcalfe, M. Zhou, C. Lowell, and D. Linnekin. 2001. Lyn is required for normal

stem cell factor-induced proliferation and chemotaxis of primary hematopoietic

cells. Blood. 98:343-350.

Parent, C.A., B.J. Blacklock, W.M. Froehlich, D.B. Murphy, and P.N. Devreotes. 1998.

G protein signaling events are activated at the leading edge of chemotactic cells.

Cell. 95:81-91.

Pereira, S., and C. Lowell. 2003. The Lyn tyrosine kinase negatively regulates neutrophil

integrin signaling. J Immunol. 171:1319-1327.

Philips, M.R., A. Feoktistov, M.H. Pillinger, and S.B. Abramson. 1995. Translocation of

p21rac2 from cytosol to plasma membrane is neither necessary nor sufficient for

neutrophil NADPH oxidase activity. J Biol Chem. 270:11514-11521.

Price, L.S., J. Leng, M.A. Schwartz, and G.M. Bokoch. 1998. Activation of Rac and

Cdc42 by integrins mediates cell spreading. Mol Biol Cell. 9:1863-1871.

Ptasznik, A., E.R. Prossnitz, D. Yoshikawa, A. Smrcka, A.E. Traynor-Kaplan, and G.M.

Bokoch. 1996. A tyrosine kinase signaling pathway accounts for the majority of

phosphatidylinositol 3,4,5-trisphosphate formation in chemoattractant-stimulated

human neutrophils. J Biol Chem. 271:25204-25207.

Ptasznik, A., A. Traynor-Kaplan, and G.M. Bokoch. 1995. G protein-coupled

chemoattractant receptors regulate Lyn tyrosine kinase.Shc adapter protein

signaling complexes. J Biol Chem. 270:19969-19973.

Quilliam, L.A., H. Mueller, B.P. Bohl, V. Prossnitz, L.A. Sklar, C.J. Der, and G.M.

Bokoch. 1991. Rap1A is a substrate for cyclic AMP-dependent protein kinase in

human neutrophils. J Immunol. 147:1628-1635.

Quinn, M.T., T. Evans, L.R. Loetterle, A.J. Jesaitis, and G.M. Bokoch. 1993.

Translocation of Rac correlates with NADPH oxidase activation. Evidence for

equimolar translocation of oxidase components. J Biol Chem. 268:20983-20987.

Rickert, P., O.D. Weiner, F. Wang, H.R. Bourne, and G. Servant. 2000. Leukocytes

navigate by compass: roles of PI3Kgamma and its lipid products. Trends Cell

Biol. 10:466-473.

Ridley, A.J. 2001. Rho proteins, PI 3-kinases, and monocyte/macrophage motility. FEBS

Lett. 498:168-171.

Ridley, A.J., M.A. Schwartz, K. Burridge, R.A. Firtel, M.H. Ginsberg, G. Borisy, J.T.

Parsons, and A.R. Horwitz. 2003. Cell migration: integrating signals from front to

back. Science. 302:1704-1709.

Roberts, A.W., C. Kim, L. Zhen, J.B. Lowe, R. Kapur, B. Petryniak, A. Spaetti, J.D.

Pollock, J.B. Borneo, G.B. Bradford, S.J. Atkinson, M.C. Dinauer, and D.A.

Williams. 1999. Deficiency of the hematopoietic cell-specific Rho family GTPase

Rac2 is characterized by abnormalities in neutrophil function and host defense.

Immunity. 10:183-196.

127

Rosen, M.K., T. Yamazaki, G.D. Gish, C.M. Kay, T. Pawson, and L.E. Kay. 1995. Direct

demonstration of an intramolecular SH2-phosphotyrosine interaction in the Crk

protein. Nature. 374:477-479.

Sanchez-Madrid, F., and M.A. del Pozo. 1999. Leukocyte polarization in cell migration

and immune interactions. Embo J. 18:501-511.

Sarbassov, D.D., S.M. Ali, D.H. Kim, D.A. Guertin, R.R. Latek, H. Erdjument-Bromage,

P. Tempst, and D.M. Sabatini. 2004. Rictor, a novel binding partner of mTOR,

defines a rapamycin-insensitive and raptor-independent pathway that regulates the

cytoskeleton. Curr Biol. 14:1296-1302.

Sarbassov, D.D., D.A. Guertin, S.M. Ali, and D.M. Sabatini. 2005. Phosphorylation and

regulation of Akt/PKB by the rictor-mTOR complex. Science. 307:1098-1101.

Sasaki, T., J. Irie-Sasaki, R.G. Jones, A.J. Oliveira-dos-Santos, W.L. Stanford, B. Bolon,

A. Wakeham, A. Itie, D. Bouchard, I. Kozieradzki, N. Joza, T.W. Mak, P.S.

Ohashi, A. Suzuki, and J.M. Penninger. 2000. Function of PI3Kgamma in

thymocyte development, T cell activation, and neutrophil migration. Science.

287:1040-1046.

Sattler, M., and R. Salgia. 1998. Role of the adapter protein CRKL in signal transduction

of normal hematopoietic and BCR/ABL-transformed cells. Leukemia. 12:637-

644.

Schiffmann, E. 1982. Leukocyte chemotaxis. Annu Rev Physiol. 44:553-568.

Schmidt, A., J. Kunz, and M.N. Hall. 1996. TOR2 is required for organization of the

actin cytoskeleton in yeast. Proc Natl Acad Sci U S A. 93:13780-13785.

Schmitt, M., and B. Bultmann. 1990. Fluorescent chemotactic peptides as tools to

identify the f-Met-Leu-Phe receptor on human granulocytes. Biochem Soc Trans.

18:219-222.

Schymeinsky, J., A. Sindrilaru, D. Frommhold, M. Sperandio, R. Gerstl, C. Then, A.

Mocsai, K. Scharffetter-Kochanek, and B. Walzog. 2006. The Vav binding site of

the non-receptor tyrosine kinase Syk at Tyr 348 is critical for {beta}2 integrin

(CD11/CD18)-mediated neutrophil migration. Blood.

Schymeinsky, J., C. Then, and B. Walzog. 2005. The non-receptor tyrosine kinase Syk

regulates lamellipodium formation and site-directed migration of human

leukocytes. J Cell Physiol. 204:614-622.

Senechal, K., C. Heaney, B. Druker, and C.L. Sawyers. 1998. Structural requirements for

function of the Crkl adapter protein in fibroblasts and hematopoietic cells. Mol

Cell Biol. 18:5082-5090.

Servant, G., O.D. Weiner, P. Herzmark, T. Balla, aacute, J.W. Sedat, and H.R. Bourne.

2000a. Polarization of Chemoattractant Receptor Signaling During Neutrophil

Chemotaxis. Science. 287:1037-1040.

Servant, G., O.D. Weiner, P. Herzmark, T. Balla, J.W. Sedat, and H.R. Bourne. 2000b.

Polarization of chemoattractant receptor signaling during neutrophil chemotaxis.

Science. 287:1037-1040.

Servant, G., O.D. Weiner, E.R. Neptune, J.W. Sedat, and H.R. Bourne. 1999. Dynamics

of a chemoattractant receptor in living neutrophils during chemotaxis. Mol Biol

Cell. 10:1163-1178.

128

Shimonaka, M., K. Katagiri, T. Nakayama, N. Fujita, T. Tsuruo, O. Yoshie, and T.

Kinashi. 2003. Rap1 translates chemokine signals to integrin activation, cell

polarization, and motility across vascular endothelium under flow. J Cell Biol.

161:417-427.

Shin, M.E., Y. He, D. Li, S. Na, F. Chowdhury, Y.C. Poh, O. Collin, P. Su, P. de

Lanerolle, M.A. Schwartz, N. Wang, and F. Wang. 2010a. Spatiotemporal

organization, regulation and functions of tractions during neutrophil chemotaxis.

Blood.

Shin, M.E., Y. He, D. Li, S. Na, F. Chowdhury, Y.C. Poh, O. Collin, P. Su, P. de

Lanerolle, M.A. Schwartz, N. Wang, and F. Wang. 2010b. Spatiotemporal

organization, regulation, and functions of tractions during neutrophil chemotaxis.

Blood. 116:3297-3310.

Shiota, C., J.T. Woo, J. Lindner, K.D. Shelton, and M.A. Magnuson. 2006. Multiallelic

disruption of the rictor gene in mice reveals that mTOR complex 2 is essential for

fetal growth and viability. Dev Cell. 11:583-589.

Sklar, L.A., G.M. Omann, and R.G. Painter. 1985. Relationship of actin polymerization

and depolymerization to light scattering in human neutrophils: dependence on

receptor occupancy and intracellular Ca++. J Cell Biol. 101:1161-1166.

Small, J.V., K. Anderson, and K. Rottner. 1996. Actin and the coordination of protrusion,

attachment and retraction in cell crawling. Biosci Rep. 16:351-368.

Smith, A., Y.R. Carrasco, P. Stanley, N. Kieffer, F.D. Batista, and N. Hogg. 2005. A

talin-dependent LFA-1 focal zone is formed by rapidly migrating T lymphocytes.

J Cell Biol. 170:141-151.

Spangrude, G.J., F. Sacchi, H.R. Hill, D.E. Van Epps, and R.A. Daynes. 1985. Inhibition

of lymphocyte and neutrophil chemotaxis by pertussis toxin. J Immunol.

135:4135-4143.

Srinivasan, S., F. Wang, S. Glavas, A. Ott, F. Hofmann, K. Aktories, D. Kalman, and

H.R. Bourne. 2003. Rac and Cdc42 play distinct roles in regulating PI(3,4,5)P3

and polarity during neutrophil chemotaxis. J Cell Biol. 160:375-385.

Stasia, M.J., A. Jouan, N. Bourmeyster, P. Boquet, and P.V. Vignais. 1991. ADP-

ribosylation of a small size GTP-binding protein in bovine neutrophils by the C3

exoenzyme of Clostridium botulinum and effect on the cell motility. Biochem

Biophys Res Commun. 180:615-622.

Stephens, L., L. Milne, and P. Hawkins. 2008. Moving towards a better understanding of

chemotaxis. Curr Biol. 18:R485-494.

Stephens, L., A. Smrcka, F.T. Cooke, T.R. Jackson, P.C. Sternweis, and P.T. Hawkins.

1994. A novel phosphoinositide 3 kinase activity in myeloid-derived cells is

activated by G protein beta gamma subunits. Cell. 77:83-93.

Stephens, L.R., A. Eguinoa, H. Erdjument-Bromage, M. Lui, F. Cooke, J. Coadwell, A.S.

Smrcka, M. Thelen, K. Cadwallader, P. Tempst, and P.T. Hawkins. 1997. The G

beta gamma sensitivity of a PI3K is dependent upon a tightly associated adaptor,

p101. Cell. 89:105-114.

Suire, S., J. Coadwell, G.J. Ferguson, K. Davidson, P. Hawkins, and L. Stephens. 2005.

p84, a new Gbetagamma-activated regulatory subunit of the type IB

phosphoinositide 3-kinase p110gamma. Curr Biol. 15:566-570.

129

Suire, S., A.M. Condliffe, G.J. Ferguson, C.D. Ellson, H. Guillou, K. Davidson, H.

Welch, J. Coadwell, M. Turner, E.R. Chilvers, P.T. Hawkins, and L. Stephens.

2006. Gbetagammas and the Ras binding domain of p110gamma are both

important regulators of PI(3)Kgamma signalling in neutrophils. Nat Cell Biol.

8:1303-1309.

Sullivan, S.J., G. Daukas, and S.H. Zigmond. 1984. Asymmetric distribution of the

chemotactic peptide receptor on polymorphonuclear leukocytes. J Cell Biol.

99:1461-1467.

Tamariz, E., and F. Grinnell. 2002. Modulation of fibroblast morphology and adhesion

during collagen matrix remodeling. Mol Biol Cell. 13:3915-3929.

Tapon, N., and A. Hall. 1997. Rho, Rac and Cdc42 GTPases regulate the organization of

the actin cytoskeleton. Curr Opin Cell Biol. 9:86-92.

Thelen, M., M. Uguccioni, and J. Bosiger. 1995. PI 3-kinase-dependent and independent

chemotaxis of human neutrophil leukocytes. Biochem Biophys Res Commun.

217:1255-1262.

Thompson, H.L., M. Shiroo, and J. Saklatvala. 1993. The chemotactic factor N-

formylmethionyl-leucyl-phenylalanine activates microtubule-associated protein 2

(MAP) kinase and a MAP kinase kinase in polymorphonuclear leucocytes.

Biochem J. 290 ( Pt 2):483-488.

Thoreen, C.C., S.A. Kang, J.W. Chang, Q. Liu, J. Zhang, Y. Gao, L.J. Reichling, T. Sim,

D.M. Sabatini, and N.S. Gray. 2009. An ATP-competitive mammalian target of

rapamycin inhibitor reveals rapamycin-resistant functions of mTORC1. J Biol

Chem. 284:8023-8032.

Tilton, B., M. Andjelkovic, S.A. Didichenko, B.A. Hemmings, and M. Thelen. 1997. G-

Protein-coupled receptors and Fcgamma-receptors mediate activation of

Akt/protein kinase B in human phagocytes. J Biol Chem. 272:28096-28101.

Toker, A., and L.C. Cantley. 1997. Signalling through the lipid products of

phosphoinositide-3-OH kinase. Nature. 387:673-676.

Uruno, T., P. Zhang, J. Liu, J.J. Hao, and X. Zhan. 2003. Haematopoietic lineage cell-

specific protein 1 (HS1) promotes actin-related protein (Arp) 2/3 complex-

mediated actin polymerization. Biochem J. 371:485-493.

Vanhaesebroeck, B., and M.D. Waterfield. 1999. Signaling by distinct classes of

phosphoinositide 3-kinases. Exp Cell Res. 253:239-254.

Verkhovsky, A.B., T.M. Svitkina, and G.G. Borisy. 1999. Self-polarization and

directional motility of cytoplasm. Curr Biol. 9:11-20.

Vlahos, C.J., and W.F. Matter. 1992. Signal transduction in neutrophil activation.

Phosphatidylinositol 3-kinase is stimulated without tyrosine phosphorylation.

FEBS Lett. 309:242-248.

Vlahos, C.J., W.F. Matter, R.F. Brown, A.E. Traynor-Kaplan, P.G. Heyworth, E.R.

Prossnitz, R.D. Ye, P. Marder, J.A. Schelm, K.J. Rothfuss, and et al. 1995.

Investigation of neutrophil signal transduction using a specific inhibitor of

phosphatidylinositol 3-kinase. J Immunol. 154:2413-2422.

Walter, R.J., and W.A. Marasco. 1987. Direct visualization of formylpeptide receptor

binding on rounded and polarized human neutrophils: cellular and receptor

heterogeneity. J Leukoc Biol. 41:377-391.

130

Wang, F. 2009. The signaling mechanisms underlying cell polarity and chemotaxis. Cold

Spring Harb Perspect Biol. 1:a002980.

Wang, F., P. Herzmark, O.D. Weiner, S. Srinivasan, G. Servant, and H.R. Bourne. 2002a.

Lipid products of PI(3)Ks maintain persistent cell polarity and directed motility in

neutrophils. Nat Cell Biol. 4:513-518.

Wang, W., J.B. Wyckoff, V.C. Frohlich, Y. Oleynikov, S. Huttelmaier, J. Zavadil, L.

Cermak, E.P. Bottinger, R.H. Singer, J.G. White, J.E. Segall, and J.S. Condeelis.

2002b. Single cell behavior in metastatic primary mammary tumors correlated

with gene expression patterns revealed by molecular profiling. Cancer Res.

62:6278-6288.

Wang, Z., T.J. Dillon, V. Pokala, S. Mishra, K. Labudda, B. Hunter, and P.J. Stork. 2006.

Rap1-mediated activation of extracellular signal-regulated kinases by cyclic AMP

is dependent on the mode of Rap1 activation. Mol Cell Biol. 26:2130-2145.

Weiner, O.D. 2002. Regulation of cell polarity during eukaryotic chemotaxis: the

chemotactic compass. Curr Opin Cell Biol. 14:196-202.

Weiner, O.D., P.O. Neilsen, G.D. Prestwich, M.W. Kirschner, L.C. Cantley, and H.R.

Bourne. 2002. A PtdInsP(3)- and Rho GTPase-mediated positive feedback loop

regulates neutrophil polarity. Nat Cell Biol. 4:509-513.

Weiner, O.D., M.C. Rentel, A. Ott, G.E. Brown, M. Jedrychowski, M.B. Yaffe, S.P.

Gygi, L.C. Cantley, H.R. Bourne, and M.W. Kirschner. 2006. Hem-1 complexes

are essential for Rac activation, actin polymerization, and myosin regulation

during neutrophil chemotaxis. PLoS Biol. 4:e38.

Weiner, O.D., G. Servant, M.D. Welch, T.J. Mitchison, J.W. Sedat, and H.R. Bourne.

1999. Spatial control of actin polymerization during neutrophil chemotaxis. Nat

Cell Biol. 1:75-81.

Williams, D.A., W. Tao, F. Yang, C. Kim, Y. Gu, P. Mansfield, J.E. Levine, B.

Petryniak, C.W. Derrow, C. Harris, B. Jia, Y. Zheng, D.R. Ambruso, J.B. Lowe,

S.J. Atkinson, M.C. Dinauer, and L. Boxer. 2000. Dominant negative mutation of

the hematopoietic-specific Rho GTPase, Rac2, is associated with a human

phagocyte immunodeficiency. Blood. 96:1646-1654.

Worthen, G.S., N. Avdi, A.M. Buhl, N. Suzuki, and G.L. Johnson. 1994. FMLP activates

Ras and Raf in human neutrophils. Potential role in activation of MAP kinase. J

Clin Invest. 94:815-823.

Worthylake, R.A., S. Lemoine, J.M. Watson, and K. Burridge. 2001. RhoA is required

for monocyte tail retraction during transendothelial migration. J Cell Biol.

154:147-160.

Wullschleger, S., R. Loewith, and M.N. Hall. 2006. TOR signaling in growth and

metabolism. Cell. 124:471-484.

Wymann, M.P., and L. Pirola. 1998. Structure and function of phosphoinositide 3-

kinases. Biochim Biophys Acta. 1436:127-150.

Xiao, Z., N. Zhang, D.B. Murphy, and P.N. Devreotes. 1997. Dynamic distribution of

chemoattractant receptors in living cells during chemotaxis and persistent

stimulation. J Cell Biol. 139:365-374.

131

Xu, J., F. Wang, A. Van Keymeulen, P. Herzmark, A. Straight, K. Kelly, Y. Takuwa, N.

Sugimoto, T. Mitchison, and H.R. Bourne. 2003. Divergent signals and

cytoskeletal assemblies regulate self-organizing polarity in neutrophils. Cell.

114:201-214.

Xu, J., F. Wang, A. Van Keymeulen, M. Rentel, and H.R. Bourne. 2005. Neutrophil

microtubules suppress polarity and enhance directional migration. Proc Natl Acad

Sci U S A. 102:6884-6889.

Yan, S.R., L. Fumagalli, and G. Berton. 1996. Activation of SRC family kinases in

human neutrophils. Evidence that p58C-FGR and p53/56LYN redistributed to a

Triton X-100-insoluble cytoskeletal fraction, also enriched in the caveolar protein

caveolin, display an enhanced kinase activity. FEBS Lett. 380:198-203.

Yasui, K., M. Yamazaki, M. Miyabayashi, T. Tsuno, and A. Komiyama. 1994. Signal

transduction pathway in human polymorphonuclear leukocytes for chemotaxis

induced by a chemotactic factor. Distinct from the pathway for superoxide anion

production. J Immunol. 152:5922-5929.

Ydrenius, L., L. Molony, J. Ng-Sikorski, and T. Andersson. 1997. Dual action of cAMP-

dependent protein kinase on granulocyte movement. Biochem Biophys Res

Commun. 235:445-450.

Yi, T.L., J.B. Bolen, and J.N. Ihle. 1991. Hematopoietic cells express two forms of lyn

kinase differing by 21 amino acids in the amino terminus. Mol Cell Biol. 11:2391-

2398.

Yokota, M., N. Tatsumi, and I. Yano. 1993. Effect of okadaic acid on leukocyte

movement. Biochem Mol Biol Int. 29:799-805.

Yoshinaga-Ohara, N., A. Takahashi, T. Uchiyama, and M. Sasada. 2002. Spatiotemporal

regulation of moesin phosphorylation and rear release by Rho and

serine/threonine phosphatase during neutrophil migration. Exp Cell Res. 278:112-

122.

Zhelev, D.V., A.M. Alteraifi, and D. Chodniewicz. 2004. Controlled pseudopod

extension of human neutrophils stimulated with different chemoattractants.

Biophys J. 87:688-695.

Zheng, L., J. Eckerdal, I. Dimitrijevic, and T. Andersson. 1997. Chemotactic peptide-

induced activation of Ras in human neutrophils is associated with inhibition of

p120-GAP activity. J Biol Chem. 272:23448-23454.

Zigmond, S.H. 1977. Ability of polymorphonuclear leukocytes to orient in gradients of

chemotactic factors. J Cell Biol. 75:606-616.

Zigmond, S.H., M. Joyce, J. Borleis, G.M. Bokoch, and P.N. Devreotes. 1997.

Regulation of actin polymerization in cell-free systems by GTPgammaS and

Cdc42. J Cell Biol. 138:363-374.

Zigmond, S.H., H.I. Levitsky, and B.J. Kreel. 1981. Cell polarity: an examination of its

behavioral expression and its consequences for polymorphonuclear leukocyte

chemotaxis. J Cell Biol. 89:585-592.

Zu, Y.L., J. Qi, A. Gilchrist, G.A. Fernandez, D. Vazquez-Abad, D.L. Kreutzer, C.K.

Huang, and R.I. Sha'afi. 1998. p38 mitogen-activated protein kinase activation is

required for human neutrophil function triggered by TNF-alpha or FMLP

stimulation. J Immunol. 160:1982-1989.

132

Zwartkruis, F.J., R.M. Wolthuis, N.M. Nabben, B. Franke, and J.L. Bos. 1998.

Extracellular signal-regulated activation of Rap1 fails to interfere in Ras effector

signalling. Embo J. 17:5905-5912.