Molecular Mechanism of Podosome Formation and Proteolytic ... · critical for podosome formation,...

Molecular Mechanism of Podosome Formation

and Proteolytic Function in Human Bronchial

Epithelial Cells

By

Helan Xiao

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Graduate Department of Physiology

University of Toronto

Copyright by Helan Xiao 2009

ii

Molecular Mechanism of Podosome Formation and Proteolytic Function in

Human Bronchial Epithelial Cells

Helan Xiao

Doctor of Philosophy

Graduate Department of Physiology

University of Toronto

2009

Abstract

In the lung, epithelial cell migration plays a key role in both physiological and

pathophysiological conditions. When the respiratory epithelium is injured, the epithelial lining in

the respiratory system can be seriously damaged. Spreading and migrating of the surviving cells

neighboring a wound are essential for airway epithelial repair. When the repair process is

affected, aberrant remodelling may occur, which is important in the pathogenesis of lung

diseases. However, in comparison with other cellular and molecular functions in the respiratory

system, our understanding on lung epithelial cell migration and invasion is limited.

To gain insight into the molecular mechanisms that govern these cellular processes, I

asked whether normal (non-cancerous) human airway epithelial cells can form podosomes, a

cellular structure discovered from cancer and mesenchymal cells that controls cell migration and

invasion. I found that phorbol-12, 13-dibutyrate (PDBu), a protein kinase C (PKC) activator,

induced podosome formation in primary normal human bronchial epithelial cells, and in normal

human airway epithelial BEAS2B cells. PDBu-induced podosomes were capable of degrading

iii

fibronectin-gelatin-sucrose matrix. PDBu also increased the invasiveness of these epithelial cells.

I further demonstrated that PDBu-induced podosome formation was mainly mediated through

redistribution of conventional PKCs, especially PKC , from the cytosol to the podosomes,

whereas atypical PKC played a dominant role in the proteolytic activity of podosomes through

recruitment of MMP-9 to podosomes, and MMP-9 secretion and activiation. I also found that

that PDBu can activate PI3K/Akt/Src and ERK1/2 and JNK but not p38. PI3K, Akt and Src were

critical for podosome formation, whereas ERK1/2 and JNK mediated the proteolytic activity of

podosomes via MMP-9 recruitment, gene expression, release and activation without affecting

podosome assembly.

Podosomes are important for epithelial cell migration and invasion, thus contributing to

respiratory epithelial repair and regeneration. My thesis work unveils the molecular mechanisms

that regulate podosomal formation and proteolytic function in normal human bronchial epithelial

cells. These novel findings may enhance our understanding of cell migration and invasion in

lung development and repair. Similar mechanisms may be also applicable to other cell types in

distinct organs.

iv

Acknowledgements

The thesis is to my beloved parents Mr. Qingcai Xiao and Ms. Dong Li. I sincerely

appreciate their love and support throughout my life.

First and foremost, I would like to give my greatest gratitude to my supervisor Dr.

Mingyao Liu, who has provided me opportunity, knowledge, mentorship and assistance in all

these six years. As an international student and non-native English speaker, I met numerous

obstacles when I first came to Toronto. Dr. Liu always encourages me and tries his best to cheer

me up whenever I had troubles. Thanks to all of my current and previous academic committee

members, Dr. Andras Kapus, Dr. Alan S. Mak, Dr. Wei-Yang Lu, Dr. John F. MacDonald and

Dr. Daniela Rotin, for their continuous guidance, help, support, thoughtful discussion, friendship

and inspiration. I will always remember their serious scientific attitudes, passion for research,

dedication to science, and industrious work.

I would like to give my special gratitution to Dr. Shaf Keshavjee, Dr. Thomas Waddell,

Dr. Marc de Perrot and Dr. David Hwang for their support, leadership and help. I would like to

especially express my thanks to Dr. Jing Xu, Dr. Monika Lodyga, Dr. Feng Xu, Dr. Bing Han,

Ms. Xiao-Hui Bai, and Dr. Jeya Nadesalingam who have provided me their technical expertise,

suggestion, kindly assistance, thoughtful discussion, encouragement and friendship. Special

thanks to Ms. Xiao-Hui Bai to teach me Real-time RT-PCR and PKC assay, to Mr. Rob Eves to

teach me in situ zymography and gelatin gel zymography assay. I am also fortunate the dear

friendship, help and support from all of my current and previous lab members: Mr. Peter S.

Tang, Dr. Atsushi Shiozaki, Ms. Yu Zhang, Dr. Xiaolin He, Dr. Marco Mura, Dr. Takeshi

Oyaizu, Ms. Roli Bawa, Mr. Matthew Rubacha, Dr. Yongfang Yuan, Dr. Shane Fung and Ms.

v

Ivone Ornelas. Many thanks to the help from all of my summer students: Mr. David Laugren,

Mr. Wayne Kan, Ms. Christina Yep, Ms. Debbie Li, and Mr. Wasim Kagzi. I would give my

special acknowledgements to our colaboraters in Queen’s University for their contributions to

my publications, technical support, contributive and thoughtful discussions: Dr. Alan S. Mak,

Mr. Rob Eves, Ms. Lily Jia, and Dr. Bradley Webb. Great thanks to our entire research team

members and local collaborators: Dr. Lowell Langille, Ms. Hanna Zhihong Yun, Dr. Amy

Wong, Dr. Masaaki Sato, Dr. Kota Ishizawa, Dr. Masashi Gotoh, Dr. Shin Hirayama, Dr. Licun

Wu, Dr. Macelo Cypel, Dr. Masaki Anruku, Mr. Pascal Dunchesneau, Mr. Paul Chartrand, Dr.

Rongyu Jin, Dr. Yingzhe Zhou, Dr. Peter Sabatini, Mr. Marc L. Chretien, Dr. Rosalind

Silverman and Dr. Lorelei Silverman.

I would like to give my sincere gratitude to the current and previous graduate

coordinators, graduate administrators and graduate assistants in Department of Physiology for

their help, kind suggestion, consultant and encouragement, Dr. Milton P. Charlton, Dr. Martin

Wojtowicz, Dr. Denise Belsham, Ms. Julie Weedmark, Ms. Rosalie Pang and Ms. Colleen Shea.

Great thanks to all professors and teaching assistants in the courses I have taken. And many

thanks to all the people who ever kindly helped me during my PhD study.

I am also grateful for the financial support from Peterborough K. M. Hunter Graduate

Studentship for cancer research.

vi

Table of Contents

ABSTRACT.................................................................................................................................. II

ACKNOWLEDGEMENTS .......................................................................................................IV

TABLE OF CONTENTS ...........................................................................................................VI

LIST OF FIGURES .................................................................................................................. XII

LIST OF TABLES ...................................................................................................................XVI

LIST OF ABBREVIATIONS ............................................................................................... XVII

LIST OF PUBLICATIONS .................................................................................................XXIII

1. CHAPTER ONE. GENERAL BACKGROUND - EPITHELIAL CELL MIGRATION

IN HUMAN LUNG PHYSIOLOGY, PATHOPHYSIOLOGY AND RESPIRATORY

DISEASES ..................................................................................................................................... 1

1.1 Overview - Lung epithelial cell migration ........................................................................ 3

1.2 Respiratory epithelial cell migration is an early event in the repair process ................ 4

1.2.1 Epithelial cells in the airway and lung parenchyma................................................. 4

1.2.2 Airway epithelial cell migration occurs after tissue injury as an early event ........ 6

1.3 Regulation of airway epithelial cell migration ............................................................... 10

1.4 Alveolar epithelial cell migration occurs after tissue injury......................................... 13

vii

1.5 Perspective: insight from epithelial cell migration to understanding of injury and

repair of lung diseases ............................................................................................................ 15

2. CHAPTER TWO. GENERAL BACKGROUND - CELLULAR STRUCTURES FOR

CELL ADHESION, MIGRATION AND INVASION ............................................................ 17

2.1 Molecular mechanisms of cell migration ........................................................................ 18

2.2 Focal adhesion ................................................................................................................... 21

2.3 Podosome ........................................................................................................................... 24

2.4 Invadopodia ....................................................................................................................... 28

2.5 Molecular components of podosomes.............................................................................. 28

2.5.1 Src................................................................................................................................ 29

2.5.2 PI3K/Akt..................................................................................................................... 31

2.5.3 MAPKs........................................................................................................................ 34

2.5.4 Matrix metalloproteinases......................................................................................... 36

2.6 Focal adhesion vs. podosome: what is the difference? .................................................. 42

2.7 Podosome vs. invadopodia: what is the difference?....................................................... 42

3. CHAPTER THREE. GENERAL BACKGROUND - PROTEIN KINASE C FAMILY IN

CYTOSKELETON REARRANGEMENT AND CELL MOTILITY................................... 47

3.1 Introduction of PKC family ............................................................................................. 48

3.1.1 Molecular structure of PKC family.......................................................................... 49

3.1.2 Mechanism of PKC activation .................................................................................. 51

3.2 The role of PKCs in cell motility ..................................................................................... 54

3.2.1 Classical PKCs in cell motility .................................................................................. 54

viii

3.2.2 Novel PKCs in cell motility ....................................................................................... 56

3.2.3 Atypical PKCs in cell motility................................................................................... 59

3.2.4 PKCµ in cell motility ................................................................................................. 62

4. CHAPTER FOUR. RATIONALE, HYPOTHESIS AND SPECIFIC AIMS.................... 64

4.1 Rationale ............................................................................................................................ 65

4.2 Hypothesis.......................................................................................................................... 66

4.3 Specific aims ...................................................................................................................... 66

5. CHAPTER FIVE. MATERIALS AND METHODS ........................................................... 68

5.1 Reagents and antibodies ................................................................................................... 69

5.2 Cell culture ........................................................................................................................ 70

5.3 siRNA transfection............................................................................................................ 71

5.4 Immunofluorescence staining .......................................................................................... 71

5.5 Microscopy and image analysis ....................................................................................... 72

5.6 Live cell imaging ............................................................................................................... 73

5.7 Fibronectin based in situ zymography............................................................................ 74

5.8 Gelatin based in situ zymography ................................................................................... 75

5.9 Cell invasion assay ............................................................................................................ 75

5.10 Western blotting.............................................................................................................. 76

5.11 Gelatinase zymography gel ............................................................................................ 76

5.12 Real-Time quantitative RT-PCR................................................................................... 77

5.13 Statistics ........................................................................................................................... 78

ix

6. CHAPTER SIX. PHORBOL ESTER-INDUCED PODOSOMES IN NORMAL HUMAN

BRONCHIAL EPITHELIAL CELLS...................................................................................... 79

6.1 Summary............................................................................................................................ 80

6.2 Introduction....................................................................................................................... 80

6.3 Results ................................................................................................................................ 82

6.3.1 PDBu induces reorganization of cytoskeleton in normal human bronchial

epithelial cells ...................................................................................................................... 82

6.3.2 Identification of ventral podosomes and circular dorsal ruffles ........................... 86

6.3.3 Formation of podosomes in primary normal human bronchial epithelial cells... 89

6.3.4 Accumulation of actin-associate proteins and regulators in podosome-like

structures ............................................................................................................................. 93

6.3.5 Recruitment of integrin and associated proteins in podosome-like structures .... 96

6.3.6 Lack of vinculin in PDBu-induced small dots in normal human lung epithelial

cells ....................................................................................................................................... 99

6.3.7 Increased protein tyrosine kinases in podosome-like structures......................... 102

6.3.8 Distribution of MMPs in podosome-like structures ............................................. 102

6.3.9 PDBu-induced podosome-like structures can degrade extracellular matrix ..... 107

6.4 Discussion......................................................................................................................... 110

7. CHAPTER SEVEN. PKC REGULATES RECRUITMENT OF MMP-9 TO

PODOSOMES, AND ITS RELEASE AND ACTIVATION ................................................ 115

7.1 Summary.......................................................................................................................... 116

7.2 Introduction..................................................................................................................... 117

7.3 Results .............................................................................................................................. 118

x

7.3.1 PDBu-induced PKC phosphorylation and translocation to podosomes and

circular ruffles................................................................................................................... 118

7.3.2 PDBu-induced podosome assembly depends on cPKCs....................................... 124

7.3.3 PDBu-induced matrix degradation can be blocked by siRNAs of multiple PKC

isoforms.............................................................................................................................. 127

7.3.4 PKC mediates PDBu-induced proteolytic activity of podosomes, MMP-9

recruitment to podosome, and MMP-9 release and activation..................................... 130

7.3.5 PDBu-induced MMP-9 is involved in degradation of gelatin matrix ................. 136

7.3.6 Recruitment of PKC to PDBu-induced posodosmes depended on nPKC......... 139

7.4 Discussion......................................................................................................................... 144

8. CHAPTER EIGHT. COORDINATED REGULATION OF PKC ACTIVATION-

INDUCED PODOSOME FORMATION AND PROTEOLYTIC ACTIVITIES OF PI3K,

SRC AND MAPK PATHWAYS IN HUMAN BRONCHIAL EPITHELIAL CELLS...... 148

8.1 Summary.......................................................................................................................... 149

8.2 Introduction..................................................................................................................... 150

8.3 Results .............................................................................................................................. 151

8.3.1 PDBu-induced protein phosphorylation and translocation of PI3K/Akt related

signaling molecules............................................................................................................ 151

8.3.2 Effects of PDBu-stimulation on phosphorylation and distribution of Src and

MAPKs............................................................................................................................... 153

8.3.3 PI3K/Akt and Src regulates podosome assembly and proteolytic activity......... 155

8.3.4 ERK and JNK mediate proteolytic activity of PDBu-induced podosomes without

affecting podosome formation ......................................................................................... 160

xi

8.3.5 Crosstalk among PI3K/Akt, Src and MAPK pathways ....................................... 164

8.4 Discussion......................................................................................................................... 169

9. CHAPTER NINE. SUMMARY, GENERAL DISCUSSION AND PERSPECTIVES .. 172

9.1 Summary.......................................................................................................................... 173

9.2 Discussion......................................................................................................................... 175

9.2.1 The role of podosomes in normal bronchial epithelial cells in lung physiology and

pathophysiology ................................................................................................................ 175

9.2.2 PKCs and cell motility in lung physiology and pathophysiology ........................ 176

9.2.3 PKC isoforms and isoform redundancy ................................................................ 177

9.2.4 Cell migration vs. cell invasion ............................................................................... 177

9.2.5 Podosome formation vs. podosomal proteolytic function .................................... 178

9.2.6 MMP-9 gene expression vs. MMP-9 recruitment ................................................. 179

9.2.7 Study approaches and their limitations ................................................................. 182

9.2.7.1 Cell lines and primary cells.............................................................................. 182

9.2.7.2 Chemical inhibitor and siRNA ........................................................................ 183

9.2.7.3 In vitro vs. in vivo ............................................................................................. 183

9.3 Conclusions...................................................................................................................... 184

9.4 Future directions............................................................................................................. 186

10. REFERENCES.................................................................................................................... 188

xii

List of Figures

Figure 1-1 Epithelial cells in the lung 5

Figure 1-2 Lung epithelial repair after injury 7

Figure 1-3 Cellular mechanism of lung epithelial repair after injury 8

Figure 2-1 Steps of cell migration 19

Figure 2-2 Structure of focal adhesion 22

Figure 2-3 Morphology of lamellipodia, podosomes and invadopodia 26

Figure 2-4 Structure of podosome 27

Figure 2-5 PI3K/Akt signaling pathway 33

Figure 2-6 MAPKs signaling pathway 35

Figure 2-7 Molecular structure of MMPs 38

Figure 3-1 Molecular structure of PKCs 50

Figure 3-2 The serine/thronine phosphorylation sites of PKCs in the catalytic

domain

52

Figure 3-3 Molecular mechanism of PKCs activation 53

Figure 6-1 PDBu induces reorganization of cytoskeletal structure in BEAS2B

cells

84

Figure 6-2 Characterization of ventral podosomes and circular dorsal ruffles in

BEAS2B cells

87

Figure 6-3 Characterization of ventral podosomes and circular dorsal ruffles in

primary normal human bronchial epithelial cells

90

Figure 6-4 PDBu induces reorganization of cytoskeletal structure in A549 cells 92

xiii

Figure 6-5 Localization of actin associated proteins in podosomes and circular

dorsal ruffles

94

Figure 6-6 PDBu-induced redistribution of integrin and integrin associated

proteins

97

Figure 6-7 Lack of translocation of vinculin to actin-rich dots in normal human

airway epithelial cells after PDBu stimulation

100

Figure 6-8 Protein tyrosine kinases were recruited to PDBu-induced podosomes

and circular dorsal ruffles

103

Figure 6-9 Matrix metalloproteases are associated with podosomes and circular

dorsal ruffles

105

Figure 6-10 BEAS2B cells with podosomes can degrade matrix and are invasive 108

Figure 7-1 PDBu-induced PKC phosphorylation 120

Figure 7-2 PDBu-induced PKC translocation to podosomes 122

Figure 7-3 PDBu-induced podosome formation is mainly mediated through

classical PKC, but matrix degradation can be blocked by multiple

PKC inhibitors

125

Figure 7-4 PDBu-induced podosome formation is reduced by PKC siRNA but

matrix degradation can be blocked by multiple PKC siRNA

128

Figure 7-5 PKC regulates proteolytic activity of PDBu-induced podosomes

and MMP-9 translocation to podosomes

131

Figure 7-6 PKC regulates MMP-9 release and activation 134

Figure 7-7 Role of MMP-9 and MMP-14 in PDBu-induced proteolytic activity

of podosomes

137

xiv

Figure 7-8 Rottlerin reduced PDBu-induced translocation of PKC to

podosomes

140

Figure 7-9 PKC controls recruitment of PKC and MMP-9 to podosomes 142

Figure 8-1 PDBu-induced phosphorylation and translocation of PI3K/Akt

related proteins

152

Figure 8-2 PDBu-induced phosphorylation and translocation of Src and MAPK

signaling proteins

154

Figure 8-3 PI3K and Akt regulate podosome assembly and its proteolytic

activity through MMP-9 expression, release and activation

156

Figure 8-4 Src is involved in podosome formation and proteolytic activity 158

Figure 8-5 ERK and JNK mediate proteolytic activity of PDBu-induced

podosomes

161

Figure 8-6 Inhibitors for ERK and JNK pathways prevented MMP-9

translocation to PDBu-induced podosomes

163

Figure 8-7 Confirmation of ERK pathway in PDBu-induced gelatin matrix

degradation, MMP-9 translocation and gene expression with another

MEK inhibitor

166

Figure 8-8 Effects of multiple inhibitors on phosphorylation status of PI3K/Akt,

Src and MAPKs

168

Figure 9-1 Role of PKCs in podosome formation and podosomal proteolytic

function

174

Figure 9-2 MAPK signaling pathways regulate MMP-9 181

Figure 9-3 PKC, PI3K, Akt, Src and MAPK in podosome formation and 185

xv

podosomal proteolytic function

xvi

List of Tables

Table 1-1 Factors inhibit airway epithelial cell migration 11

Table 1-2 Factors promote airway epithelial cell migration 12

Table 1-3 Factors affect alveolar epithelial cell migration 14

Table 2-1 Comparison podosome and invadopodia in 2D vs. 3D culture 44

xvii

List of Abbreviations

ABR Actin binding repeat; cortacin domain containing the repeat region

ADAM A disintegrin and metalloprotease

ALI Acute lung injury

aPKC Atypical protein kinase C

ARDS Acute respiratory distress syndrome

Arp actin related protein

Arp2/3 Arp2/3 complex; complex of 7 proteins including Arp2 and Arp3, which

nucleate dendritic actin filaments

ATCC American type culture collection

ATP Adenosine triphosphate

BSA Bovine serum albumin

CCh Carbachol

cDNA Complementary DNA

CHO Chinese hamster ovarian cell line

cPKC Conventional protein C; classical PKC

CRIB Cdc42/Rac1 interacting binding domain

Csk Carboxyl-terminal Src kinase

DAG Diacylglycerol

DH Dbl homology domain

DMEM Dulbecco's modified Eagle's medium

DMSO Dimethyl sulfoxide

xviii

dsRNA Double stranded RNA

DTT dithiothreitol

Dyn2 Dynamin 2

E-cadherin Epithelial-cadherin

E. coli Escherichia coli

ECM Extracellular matrix

EDTA Ethylene diaminetetra acetic acid

EGF Epidermal growth factor

EGFP enhanced green fluorescent protein

EGFR Epithelial growth factor receptor

EGTA ethylene-bis (oxyethylenenitrilo) tetraacetic acid

EMT Epithelial-mesenchymal transition

ERK extracellular signal regulated kinase

F-actin Filamentous actin; a two-stranded polymer comprised of actin monomers

FAK Focal adhesion kinase

FBS Fetal bovine serum

FITC Fluorescein isothiocyanate

FRET Fluorescence resonance energy transfer

GABA -aminobutyric acid

G-actin Globular actin; monomeric actin

GAP GTPase-activating protein

GBD GTPase binding domain

G-CSF Granulocyte colony stimulating factor

xix

GDI Guanine nucleotide dissociation inhibitor

GEF Guanine nucleotide exchange factor

GFP Green fluorescent protein

GIT1 G-protein-coupled recptor kinase-interacting target 1

GITBD GIT1-binding domain of PIX

GPCR G-protein-coupled receptor

GST Glutathione S-transferase

HB-EGF Heparin-binding EGF-like growth factor

HEPES N-[2-hydroxyethyl] piperazine-N’-[2-ethanesulfonic acid]

HCl Hydrochloric acid

HDL High-density lipoproteins

HG-DMEM High glucose Dulbecco's modified Eagle's medium

HGF Hepatocyte growth factor

HIF-1 Hypoxia inducible factor-1

ICAM-1 Intercellular adhesion molecule-1

IGF Insulin growth factor

IKK I kappa B kinase

IL Interleukin

JNK c-Jun N-terminal kinase

kDa Kilo Dalton

LDL Low-density lipoproteins

LPS Lipopolysaccharide

MAPK Mitogen-activated protein kinase

xx

MARCKS Myristolated alanine-rich C kinase substrate

MMP Matrix metalloproteinases

MT1-MMP Membrane type 1-matrix metalloproteinases; MMP-14

NaCl Sodium chloride

NaF Sodium fluoride

NF- B Nuclear factor kappa B

NO Nitric oxide

nPKC Novel protein kinase C

N-terminal Amino terminal

PAGE Polyacrylamide gel electrophoresis

PAK p21 activated kinase

PBS Phosphate buffered saline

PCR Polymerase chain reaction

PDBu Phorbol-12,13-dibutyrate

PDGF Platelet derived growth factor

PDK1 Phosphoinositide-dependent kinase 1

PH domain Pleckstrin homology domain

PI3K Phosphatidylinositol 3-kinase

PICK1 Protein interacting with C kinase 1

PIX PAK-interacting exchange facyor

PKB Proteins kinase B

PKC Protein kinase C

PLC Phospholipase C

xxi

PMA Phorbol 12-myristate 13-aceteate

PPAR Peroxisome proliferators activated receptor

PS region Phosphtidylserine region

PTEN Phosphatase and tensin homolog deleted on chromosome ten

PTK Protein tyrosine kinase

pTyr Phosphorylated tyrosine

RACK Receptors for activated C kinase

RFP Red fluorescent protein

RICK Receptors for inactive C kinase

RNAi RNA interference

ROK Rho kinase

RPMI Roswell Park Memorial Institute medium

RT-PCR Real-time transcriptase chain reaction

SDS Sodium dodecyl sulfate

SDS-PAGE Sodium dodecyl sulphate polyacrylamide gel electrophoresis

SFK Src family kinase

SH2 Src homology domain 2

SH3 Src homology domain 3

siRNA Short interfering RNA

SSeCKS Src-suppressed C kinase substrate

TGF Transforming growth factor

TLR Toll-like receptor

Tris 2-amino-2-(hydroxymethyl)-1,3-propanediol

xxii

TRITC Tetramethylrhodamine isothiolcyanate

Triton X-100 t-octylphenoxypolyethoxyethanol

uPA Urokinase plasminogen activator

VCR Verprolin, central and acidic domain of the WASp family of NPFs

VSMC Vascular smooth muscle cells

WASp Wiskott Aldrich Syndrome protein

WIP WASp interacting protein

WT Wild type

xxiii

List of Publications

Publications included in PhD thesis:

Xiao H, Mura M, Li D, Liu M. Epithelial cell migration in respiratory diseases. (Review) (Submitted to American Journal of Physiology-Lung Cellular and Molecular Physiology)(Chapter 1)

Xiao H, Liu M. PKC in cell motility. (Review) (In preparation) (Chapter 3)

Xiao H, Eves R, Yeh C, Kan W, Xu F, Mak AS, Liu M. Phorbol ester-induced podosome formation in human bronchial epithelial cells. Journal of Cellular Physiology, 2009, 218(2): 366-375. (Chapter 6)

Xiao H, Bai XH, Kapus A, Mak AS, Lu WY, Liu M. PKC regulates recruitment of MMP-9 to podosomes, and its release and activation. (2nd revision of Molecular Biology of the Cell)(Chapter 7)

Xiao H, Bai XH, Mak AS, Liu M. Coordinated regulation of PKC activation-induced podosome formation and proteolytic activities of PI3K, Src and MAPK pathways in human bronchial epithelial cells. (In preparation for Journal of Cell Science) (Chapter 8)

Publications not included in PhD thesis:

Xiao H*, Cai G, Liu M. Fe2+-catalyzed nonenzymatic glycosylation alters collagen conformation during AGE-collagen formation in vitro. Archives of Biochemistry and Biophysics, 2007, 468(2):183-192. (* Corresponding author)

Xu J, Bai XH, Lodyga M, Han B, Xiao H, Liu M. XB130, a novel adaptor protein for signal transduction. Journal of Biological Chemistry, 2007, 282(22): 16401-16412.

Han B*, Xiao H*, Xu J, Bai XH, Liu M. Regulation of Src activity by AFAP leading to SRE/AP-1 transcriptional activation. (Under revision of Biochemical Journal, * contribute equally)

Xiao H, Liu M. XB130, Fish/Tks5 and Src interaction in podosome formation and proteolytic function. (In preparation)

1

1. Chapter One. General Background - Epithelial cell migration in human lung physiology,

pathophysiology and respiratory diseases

Part of the content of this chapter was prepared as a review article for American Journal of

Physiology-Lung Cellular and Molecular Physiology:

Xiao H, Mura M, Li D, Liu M. Epithelial cell migration in respiratory diseases. (Review)

(Submitted)

2

1. Chapter One. General background - Epithelial cell migration in human lung physiology,

pathophysiology and respiratory diseases

Respiratory epithelial cells are more than a structural component in the lung and may

contribute to normal lung physiology such as lung development and homoeostasis, and lung

repair after injury (1) through cell migration, differentiation and proliferation (2). These

processes are temporally strictly regulated by specific intracellular signaling pathways. My PhD

studies focus on the discovery of podosomes, a cellular structure responsible for cell invasion

and migration, formed by normal human bronchial epithelial cells. I have found that several

protein kinase C (PKC) isozymes are involved in podosome formation and proteolyic activities

in a coordinated fashion. I further found that PKC activated other signaling pathways, which also

contributed to the podosome functions. Firstly, in this chapter, I will review the role of epithelial

cell migration in human lung physiology, pathophysiology and various respiratory diseases.

Secondly, in Chapter two, I will introduce the cellular structures especially podosomes involved

in cell adhesion, migration and invasion. Thirdly, in Chapter three, I will review the role of

protein kinase C (PKC) family in cell motility.

In this chapter, I will focus on the role of epithelial cell migration in lung physiology,

pathophysiology and human lung diseases. Firstly, I will review the sub-types of epithelial cells

in the lungs and their specific physiological functions. Secondly, I will introduce the role of

airway epithelial cell motility and their regulatory factors under physiological and

pathophysiological conditions. Thirdly, I will summarize the role of alveolar epithelial cell

migration and their specific mediatory factors in normal lung physiology and respiratory

diseases.

3

1.1 Overview - Lung epithelial cell migration

Cell migration is a central process in the development and maintenance of multicellular

organisms. In the adult, cell migration is central to homeostatic processes such as mounting an

effective immune response and the repair of injured tissues. Cell migration has been described in

many organs and systems, contributing to many pathologic conditions, such as chronic

inflammatory diseases, vascular diseases, tumour formation and metastasis. In wound repair,

migration of epithelial cells and other tissue cells is required to heal the injury.

In the lung, epithelial cell migration plays a key role in both physiological and

pathophysiological conditions. Epithelial cell migration is involved in embryo and fetal lung

development, lung growth, maintenance of the air-blood barrier and homeostasis. When the

respiratory epithelium is injured by various insults, such as inhaled dusts, toxins, cigarette

smoking, infection of airborne micro-organisms, physical injuries (e.g. endotracheal intubation

and mechanical ventilation), the epithelial lining in the respiratory system can be seriously

damaged. In these pathophysiological conditions, cell migration as well as cell proliferation is

required for proper tissue repair, in order to restore proper barrier and gas exchange functions.

When the repair process is impaired, aberrant remodelling may occur, which plays an important

role in the pathogenesis of several lung disorders. However, in comparison with other cellular

and molecular processes in the respiratory system, our understanding on lung epithelial cell

migration is limited. The objective of this chapter is to promote interests in this fundamental

research area.

Although epithelial cell migration also plays a very important role in tumour cell invasion

and metastasis, in this chapter we will only focus on respiratory epithelial cell migration in lung

physiology and various non-tumour disorders. To facilitate the discussion, we will focus on the

4

role of cell migration in the repair process in the context of airways and lung parenchymal

diseases. Gaining a better understanding of the regulatory mechanisms of respiratory epithelial

cell migration, along with its clinical applications, may be of vital importance in promoting

normal respiratory function and healing, as well as treating many respiratory disorders that are

associated with great morbidity and mortality.

1.2 Respiratory epithelial cell migration is an early event in the repair process

1.2.1 Epithelial cells in the airway and lung parenchyma

The respiratory epithelium consists of highly differentiated epithelial cells exerting

different physiological functions at different pulmonary compartments (Figure 1-1). The

majority of the upper respiratory tree, excluding the vocal cords, is lined by pseudo-stratified, tall

columnar, ciliated epithelial cells. In the trachea and main bronchi, airway epithelial cells

provide physical protection and also participate in the innate immunity (3). Bronchial epithelial

cells provide a physical barrier with intercellular junctions against microorganisms; other

defensive mechanisms include secretion and clearance of mucus and the homeostasis of ion and

water transport (4). In the proximal bronchioles, epithelial cells take on a more cuboidal shape,

with both ciliated cells, and secretory nonciliated Clara cells. In the distal bronchioles, only Clara

cells can be identified.

Pseudostratified epithelium with gland

Pseudostratifiedepithelium

Airway smooth muscle cells

Fibroblasts

Vasculature

Columnar epithelium

Alveolar epithelium

Fig 1-1

5

Figure 1-1. Epithelial cells in the lung.

The respiratory epithelium is covered by highly differentiated epithelial cells. The upper respiratory tree is lined by pseudostratified, tall columnar, ciliated epithelial cells. In the trachea and main bronchi, bronchial epithelial cells provide a physical barrier against microorganisms. In the proximal bronchioles, epithelial cells take on a more cuboidal shape, with both ciliated cells, and secretory nonciliated Clara cells. In the distal bronchioles, both Clara and non Clara cells can be identified. Alveolar epithelial type I and type II cells cover the surface of alveoli. Type I pneumocytes are flattened squamous epithelial cells.

6

Alveolar epithelial cells cover the surface of alveoli. Type I pneumocytes are flattened

squamous epithelial cells, which interface with pulmonary capillaries and are highly permeable

to gases, thus perfectly suitable for gas exchange at the air-blood barrier. Type II pneumocytes

synthesize and secrete pulmonary surfactant, and transport sodium from apical to basolateral cell

surfaces to minimize alveolar fluid. Type II cells are involved in the innate host defense by

producing a variety of cytokines, chemokines and soluble mediators (5). Type II cells may act as

progenitor cells for type I alveolar cells (6).

1.2.2 Airway epithelial cell migration occurs after tissue injury as an early event

The respiratory epithelium is subjected to various chemical, physical, environmental and

inflammatory insults, which can vary in severity from temporary induction of surface epithelium

permeability due to destruction of tight junctions, to cell death and denudation of the epithelial

lining (Figure 1-2) (7).

The repair and regeneration of the epithelium is accomplished by three processes: the

neighbouring basal cells migrating over the denuded area, which is followed by cell

proliferation, active mitosis, squamous metaplasia, and finally, redifferentiation to pseudo-

stratified mucociliary epithelium (8) (Figure 1-3). These three steps play a distinct role at the

different stages of epithelium repair and regeneration.

Intactepithelium

DamagedepitheliumInjury Repair

Intactepithelium

Fig 1-2

7

Figure 1-2. Lung epithelial repair after injury.

The respiratory epithelium is subjected to various chemical, physical, environmental and inflammatory injuries, which can vary in severity from temporary induction of surface epithelium permeability due to destruction of tight junctions, to complete death and denudation of the epithelial cell lining.

Extracellular matrix



Fig 1-3

Surviving epithelial cells

Progenitor cell

Proliferation Cell migration

Differentiation

8

Figure 1-3. Cellular mechanism of lung epithelial repair after injury.

Lung epithelial repair mainly depends on three processes: 1) epithelial cell migration toward the wound area; 2) proliferation of the lung epithelial progenitor cells; 3) differentiation. Epithelial cell migration is an early event of the lung repair after injury. Both the airway epithelial cells and alveolar epithelial cells can migrate to cover the wound area in vitro and in vivo.

9

In order to understand the sequence of events that occur during wound healing in the

respiratory system, various experimental models have been developed, involving the induction of

injury to the respiratory epithelium and observation of the healing process and mechanisms. Cell

spreading and migration are the initial steps of wound repair in epithelial cells, as complete

wound closure has been noted in cell sheets as early as within 5-8 hours, long before the effects

of proliferation could be a contributing factor (7, 9-13). In fact, in one study, the repair process

was seen to begin almost immediately, with the remaining basal cells being seen to flatten and

migrate to cover the basement membrane within less than 20 minutes (14). Also, cell mitotic

activity typically does not peak until 24-48 hours after wounding, and mainly involved cells

located close to the wound edge (10, 12).

Interestingly, cells at the wound edge have been reported to travel 2.5 times more

distance than cells located far from the wound area (10), and migration speed was highest for

cells at the wound edge, at 35-45 µm/h (15). The reason for such variations in migration velocity

near the wound edge is likely due to the lack of contact inhibition by neighbouring cells (10, 15).

Erjefält et al studied deepithelialisation, reepithelialisation and associated events in

guinea-pig trachea after shedding-like epithelial denudation in vivo (12). They generated a

mechanical deepithelialisation of an 800 µm wide tracheal zone using an orotracheal steel probe

without bleeding or damage to the basement membrane. Immediately after epithelial removal,

secretory and ciliated and presumably basal epithelial cells at the wound margin dedifferentiated,

flattened and migrated rapidly about 2-3 µm/min over the denuded basement membrane. Within

8-15 h, a new, flattened epithelium covered the entire deepithelialised zone. At 30 h, a tight

epithelial barrier was established and after 5 days the epithelium was fully redifferentiated (16).

10

Similarly, Shimizu et al investigated epithelial regeneration in mechanically injured rat trachea

by using phenotypic markers that identify unique differentiated stages of epithelial cells (17).

Taken together, these studies demonstrate that airway epithelial cell migration is a critical

component of wound healing post injury in airways.

1.3 Regulation of airway epithelial cell migration

Numerous respiratory diseases are associated with abnormal or insufficient repair post

injury to heal the wound, leading to loss of epithelial integrity and function. For example,

chronic obstructive pulmonary disease (COPD) exacerbations may lead to dysregulation of

proinflammatory cytokines, matrix metalloproteinases (MMPs), tissue inhibitor of matrix

metalloproteinases (TIMPs), intercellular adhesion molecules, thus inducing remodelling of

airway mucosa and leakiness of epithelium (18). COPD exacerbations are often associated with

pollutants, viruses and bacteria as well. It is known that remodelling of epithelium during injury

and repair favours bacterial infections (18, 19).

We reviewed different groups of factors inhibiting or facilitating airway epithelial cell

migration during lung repair after injury. Bacterial and viral infection, inflammatory factors,

inhaled toxins, mechanical injury and alcohol abuse are major factors preventing airway

epithelial cell migration (Table 1-1), whereas certain secreted cytokines, inflammatory

mediators, growth factors, neuropeptides, extracellular matrix molecules are critical for

prompting airway epithelial cell migration (Table 1-2).

11

Table 1-1. Factors inhibit airway epithelial cell migration

Inhibitory factors Year References 1. Bovine herpesvirus-1 1995 (20) 2. Inflammatory mediators

ATP-mediated activation of dual oxidase 1 2007 (21) Endothelin-1 2003 (22) Nitric oxide (NO) 2008 (23) IL-4, IL-13 2001 (24)

3. Inhale factors Cigarette smoke 1995

2001(25)(26)

Hog barn dust 2007 (27) Feedlot dust 2007 (28) Arsenic 2008 (29) Air pollution and ozone exposure 2005 (30) Cold draught air 2001 (31) Ultrafine particles 2006 (32) High levels of ambient air pollution, particulate matter 2000 (33)

4. Mechanical ventilation 1998 (34) 5. Ethanol 2001

20052005

(31)(35)(36)

12

Table 1-2. Factors promote airway epithelial cell migration

Enhancers Year References 1. Cytokines

TNF 1996 (37) 1997 (38) IL-6 2008 (39) Chemokines to activate CXCR3 2006 (40) IFN 2001 (24)

2. Other inflammatory mediators Tachykinins 1995 (41) uPA 2001 Trefoil factor family (TFF)/EGF 2001 (42) TFF/PKC 2002 (43) Nitric oxide (NO) 2006

20072008

(23, 44, 45)

3. Growth factors Insulin and Insulin-like growth factor-I (IGF-I) 1990 (46) TGF- 2006 (47)

4. Receptor agonists Circulation hormone (i.e. catecholamine) 2002 (48)

-adrenergic receptor 2002 (49) adenosine receptors A2A 2006 (50)

5. Extracellular matrix molecules Fibronectin 1989 (51) Fibronectin via 5 1 integrin 1996 (52) ECM 1992 (53) very late adhesion integrins (VLA integrin) 1999 (54) ECM, MMP-9 1999 (55)

6. Signaling pathways PKC 1996 (37) 1997 (38) GSK3 / -catenin 2007 (56) GP130-STAT3 2008 (39)

13

1.4 Alveolar epithelial cell migration occurs after tissue injury

Lung alveolar cells have also been shown to be capable of migration to heal the wound

post-injury (57-59). Acute lung injury leads to type I alveolar epithelial cell death, denudation of

the alveolar basement membrane, and formation of an alveolar provisional matrix from

fibronectin, fibrinogen and type I collagen which provides a scaffold for alveolar repair (59). To

restore normal lung architecture, surviving type II alveolar epithelial cells reepithelialize

denuded alveoli (59). During reepithelialization, type II cells initially appear to migrate and

spread over a remodelled matrix; and then a secondary proliferative phase occurs (57).

Following acute lung injury, type II alveolar epithelial cells could migrate, spread over

and proliferate on a refaced matrix to repair the epithelium layer (60-62). It has been observed

that type II cells locomotion can be promoted by various growth factors, proinflammatory

cytokines and substrate adhesion molecules (Table 1-3). Urokinase-type plasminogen activator

(uPA) and plasminogen activator inhibitor-1 (PAI-1), inhaled particles, cytokines (IFN and IL-

6), and mechanical ventilation may inhibit alveolar epithelial cell migration (Table 1-3). Post

acute lung injury, multipotent molecules may act in different ways, either by cooperative or

counterbalancing activities, to modulate epithelial repair (57).

14

Table 1-3. Factors affect alveolar epithelial cell migration

Enhancers Year References 1. Growth Factors

PDGF, TGF- , IGF-I, KGF 1996 (57) TGF- 1994 (58, 63) TGF- 2008 (64) Hepatocyte growth factor (HGF) 1999

2003(65)(66)

Keratinocyte growth factor (KGF) 2003 (67) 2. Cytokines

IL-1 2000 (68) IFN /IL-2 combination 2004 (69) TNF- 1996 (57)

3. Extracellular matrix molecules Integrin/ECM 1997 (59) ECM/Collagenase 1999 (70) Osteopontin 2005 (71) Vitronectin/ v 1 integrin 2004 (72)

Inhibitors 1. uPA and PAI-1 2004 (72) 2. Inhale Particles

Ambient particle PM 102008 (73)

3. Cytokines IFN 1996 (57) IL-6 1996 (57)

15

1.5 Perspective: insight from epithelial cell migration to understanding of injury and repair

of lung diseases

Because respiratory epithelial injury and denudation/shedding is now known to be an

important and active component of many respiratory diseases, including inflammatory airway

diseases such as asthma and COPD, as well as remodelling diseases such as idiopathic

pulmonary fibrosis and bronchiolitis obliterans syndrome, further understanding of the

mechanisms governing epithelial repair and regeneration in vivo holds great promise in the

treatment of such diseases, which continue to hold great morbidity and mortality for its affected

patients.

Previous studies used in vivo models to study injury and repair. Later on, as cell culture

and microscopic techniques advanced, there was a trend towards employing in vitro studies to

determine the complex association of factors involved in these processes, allowing for

identification of molecular factors associated with the wound repair process. It will be necessary

to return to in vivo techniques once again, as the various limitations of cell culture models have

been described (74). The basement membrane appears to be a key player in epithelial repair post-

injury. Erjefalt et al. found the speed of cell migration in vivo to be in the range of several

µm/min, much faster than the speeds seen in cell culture experiments, pointing to the existence of

important in vivo factors that influence the repair process (12, 75). Furthermore, the clinical

features of cell migration and repair cannot be completely reproduced in cell culture models.

Performing more in vivo or ex vivo experiments with comtempory and laboratory technologies

such as real-time confocal microscopy, intravital imaging, two-photon microscopy, and real-time

multiphoton microscopy and so on, could lead to the uncovering of a wealth of knowledge with

direct clinical relevance in lung diseases.

16

Recent research in the area of tissue bioengineering and epithelial

regeneration/transplantation holds great potential as a treatment for disorders where epithelial

damage has already been incurred, with the goal to prevent abnormal healing/fibrosis and

promote functional reepithelialization. Additionally, interest in the possibility of using stem cells

to aid in repair and regeneration process of injured lung tissue has led to the discovery of adult

bone marrow-derived stem cells that may have the plasticity and ability to differentiate into

bronchial and alveolar epithelium (76-79). Circulating progenitor epithelial cells may also be

recruited to participate to repair after injury (80). Wong et al. demonstrated that targeted delivery

of short-term cultured bone marrow cells into a reversible airway injury milieu favored cell

engraftment, and may be used for cell based therapy (81). Kim et al isolated bronchioalveolar

stem cells (BASCs) from bronchioalveolar duct junction (82). BASCs are resistant to damage in

the naphthalene-induced airway denudation model. BASCs started to proliferate and self renew

at 30 h after nephelane treatment. The regeneration of intact airway epithelium is mediated by

the migration, proliferation and differentiation of BASCs in vivo, although there is some

evidence that BASCs are expaned in lung adnocarcinoma precusors and tumours (82, 83). The

migration of stem cells during tissue regeneration merit further investigation in the future (83).

In summary, in this chapter, I reviewed the importance of epithelial cell migration in lung

physiology, and pathophysiology and lung diseases. From this chapter, it is clear that the repair

of epithelium post-injury is crucial to restore barrier function in both normal physiology and to

prevent the initiation and progression of many chronic lung diseases. Research on the promotion

of epithelial repair and regeneration may lead to new therapeutic strategies, allowing for the

reconstitution of well-differentiated and functional airway epithelium.

17

2. Chapter Two. General background - Cellular structures for cell adhesion, migration and

invasion

18

2. Chapter Two. General background - Cellular structures for cell adhesion, migration and

invasion

In Chapter One, we discussed that epithelial cell migration plays an important role in

human lung physiology, pathophysiology and respiratory diseases. Cell migration results from

the coordination of motions generated in different parts of a cell and integrated with a directed

endocytic cycle. Nowadays advanced video microscopy reveals the process of cell migration in

details. In this chapter, firstly, I will review the molecular mechanisms in cell migration.

Secondly, I will summarize three types of cellular structures focal adhesion, podosomes and

invadopodia and their role in cell adhesion, cell migration and invasion. Thirdly, I will focus on

review the role of podosomal component proteins in cell motility. Fourthly, I will compare the

similarity and differences among focal adhesion, podosome and invadopodia.

2.1 Molecular mechanisms of cell migration

Cell migration involves many steps, and the ability of a cell to migrate is dependent upon

cell-cell interactions and cell-matrix interactions (84). Cell migration consists on a cyclical

process that can be divided into five phases: morphological polarization; membrane extension;

formation of cell-substratum attachments or adhesions; contractile force and traction; release or

breaking of cell attachments (Figure 2-1) (15, 85-87).






1. Morphological polarization

2. Membrane extension

3. Adhesion

4. Translocation

5. De-adhesion

Focal adhesionDirection of movement

Lamellipodium

New adhesion

Cell body movement

Detach old adhesion

Fig 2-1

19

Figure 2-1. Steps of cell migration.

Cell migration is a cyclical process that can be divided into five phases: 1) morphological polarization, which has been described as a clear distinction between cell front and rear; 2) membrane extension, which is an initial protrusion or extension of the plasma membrane at the leading edge of the cell, driven by polymerization of the cytoskeletal network of actin filaments, and stabilized by adhesive complexes; 3) adhesion, which is formation of cell-substratum attachments or adhesions at the leading edge; 4) translocation, movement of the cell body; 5) de-adhesion, also known as release or breaking of cell attachments, of adhesions located at the rear of the cell, thereby allowing for net movement in the forward direction.

20

The processes of cell migration can be turned on and off by quantitative changes in the

concentrations of various molecular components, including adhesion molecules, cytoskeletal-

linking proteins, and extracellular matrix (ECM) ligands, as well as by the changes of the

physicochemical properties surrounding the cells. The mechanisms that regulate these processes

into directional migration of the cell are still unclear, although many proteins have been

identified, such as the Rho/Rac family of signalling molecules, the integrin family of

transmembrane adhesion molecues linking the ECM to the cell’s cytoskeletal actin filaments and

microtubules (88-90).

Cdc42 and Rac are two members of the Rho family of GTPases that have been shown in

particular to be involved in regulating the formation of new cellular protrusions and focal

adhesions. Cdc42 has been shown to stimulate actin polymerization to form filopodia, and

activation of Rac stimulates lamellipodia formation. Additionally, these molecules have been

shown to regulate contractile forces at the leading edge of the cell by regulation of the myosin

light chain phosphorylation (91), via its interactions with PAK (p21 activated kinase) (92), and

via Rho kinase (93, 94).

Modification of proteins by tyrosine phosphorylation is involved in the formation of

adhesive structures. Focal adhesion kinase (FAK), paxillin, and tensin are among the

phosphoproteins known to constitute adhesive complexes (95-98). Tyrosine phosphorylation of

paxillin, tensin, and FAK creates recognition sites for proteins containing Src-homology 2 (SH2)

domains. Dysfunction in migration due to dysfunctional focal adhesion turnover has been

reported in cells lacking focal adhesion components: Src family kinases (99), FAK (100), and

calpain (101). Phosphatidylinositol 3-kinase (PI3K) and its down-stream signals, particularly

Akt (also called protein kinase B, PKB) play a central role in regulation of cell adhesion and

21

deadhesion, cell extension, migration and invasion (102). Other regulators of actin also localize

at or near the cell’s leading edge including PIP2 (103-105), WASP (106, 107), Scar (108, 109),

Arp2/3 (108); along with other molecules known to function in migration, such as integrins

(110), talin (111) and vinculin (112, 113).

2.2 Focal adhesion

Focal adhesions are small, dynamic protein complexes through which the cytoskeleton

connects to the extracellular matrix (Figure 2-2) (114). Due to the differences in their size and

shape, focal adhesions are commonly referred as focal complexes, focal contacts and fibrillar

adhesions (115). Many attempts have been made to classify these cell-substrate adhesion sites

using descriptive features such as shape, size, cellular location, GTPases dependency and protein

components (114, 116-120). The size of focal adhesion is usually within 15 nm to 60 nm (121).

The turnover rate of focal adhesion is within minutes to hours. Each cell may have many focal

adhesions in the cytoplasm and at the peripheral. The morphology of focal adhesions is arrow-

head shaped strctures as the two end of the stress fibers (122). Focal contacts are small initial

adhesions during cell attachment.


FAK

ILK

TensinPaxillin

Paxillin

Integrin

Talin

Actin-actinin

Vinculin

Actin

IAB

Cytosol

Membrane

Focal adhesion structure

Src

Fig 2-2

22

Figure 2-2. Structure of focal adhesion.

23

Focal adhesions serve both as the mechanical linkage to the extracellular matrix (123),

and as a biochemical signaling hub to concentrate and direct numerous signaling proteins,

kinases, adaptor proteins at sites of integrin binding and clustering (114). The dynamic assembly

and disassembly of focal adhesions plays a central role in cell migration. The smaller focal

adhesion, which also called “focal complexes”, is formed at the leading edge of a cell in

lamellipodia. Many of these focal complexes fail to mature and disassembly as the lamellipodia

withdraw. However, some of the focal complexes mature into larger and stable adhesions and

recruit more proteins such as zyxin (124, 125). Once in place, a focal adhesion remains

stationary with respect to the extracellular matrix, and the cell uses this as an anchor on which it

can push or pull itself over the extracellular matrix. Although focal adhesions are quite stable

under normal condition, in a motile cell, focal adhesions are being constantly assembly and

disassembly as the cell establishs new contacts at the leading edge and breaks old contacts at the

trailing edge of the cell. During cell attachment and spreading, initial adhesions evolve into small

focal complexes that further mature into focal adhesions connected to actin stress fibers.

As anchorage of the cell, focal adhesions connect with proteins of the extracellular matrix

generally through integrin (126). Integrin binds to extracellular proteins via short amino acid

sequences such as the R-G-D sequence motif in proteins such as fibronectin (127), laminin and

vitronectin (128), and the DGEA and GFOGER motifs in collagen (129). Integrins are

heterodimers which are formed from one and one subunit (130-134). Focal adhesions are

mostly composed of 1 and 3 integrins. The C-terminal PTB domain of tesins can interact

directly with the NPXY motif in the subunit of integrin (135). Within the cell, the intracellular

domain of integrin binds to the cytoskeleton via adaptor proteins such as talin, -actinin, filamin

24

and vinculin (136-140). Many signaling molecules such as FAK, Src, PKC and PI3K bind to and

associate with this integrin-adaptor protein-cytoskeleton complex (136-140).

2.3 Podosome

Podosomes are unique actin-rich structures which protrude into the extracellular matrix,

resulting in localized remodeling activities associated with enhanced invasiveness (Figure 2-3

and 2-4) (141, 142). Podosome, not only establishs close contact to the substratum, but can also

degrades components of extracellular matrix to assist motile cells to cross tissue boundaries,

thus, it has been called foot and mouth of the cell (141).

Although podosomes in different cell types share similar functions - promoting matrix

degradation and cell invasion, their morphologies appear to be diverse. For example, podosomes

in primary human macrophages are hundreds of small tiny dots (0.2-0.5 µm in size) distributed

at the ventral surface underneath the cytoplasma at the leading edge of lamellipodia (143-145).

In smooth muscle cells, podosomes appear as small dots (0.5-1 µm in size) which often

aggregate to form bands (2 µm in size) along the cortex of the cell (146). In bovine aorta

endothelial cells, podosome is one huge single ring (15 µm in size) beside the nucleus (147). In

c-Src Y527F transformed NIH 3T3 fibroblasts podosomes are numerous rosette-like structures

(2 µm in size) at the end of plasma membrane spike (148). In melanoma cells, podosomes are

couple of dots (1 µm in size) in the cytoplasm (149). In rat bladder carcinoma 804G cells,

podosome-like structures were identified around hemidesmosomes that make connections with

intermediate filaments, as small dots surrounded by rings (150, 151). Linder (152) summarized

that podosomes are dot-like structures attached to substrate, and containing actin regulators and

25

plaque proteins with the number of around 20-100 per cell with the maximum size of 1 µm in

diameter and 0.4 µm in depth.

So far nearly one hundred of proteins have been identified at the sites of podosomes. Six

groups of proteins were usually thought as the molecular components of podosomes (150): (i)

actin which is usually located at the core of podosomes and actin-associated proteins (e.g., F-

actin, cortactin, -actinin, gelsolin), and regulators (e.g., Arp2/3, WASP, N-WASP, Cdc42)

which work as actin linkers and serve as the architecture of “podosome bone”; (ii) integrin and

integrin-associated proteins (e.g., 1, 2, 3, v 2, v 3 -integrin, vinculin, paxilin, talin) which

form the cell-matrix anchor; (iii) protein kinases (e.g., Src, FAK, PKC, Pyk2, PI3K), which

regulate the actin cytoskeleton and membrane remodeling; (iv) signaling adaptor proteins (e.g.

Tks5/Fish, WIP, p130Cas, cofilin, AFAP110) which regulate podosome formation, dynamics

and maturation and lifespan; (v) motor proteins such as dynamin 2 and myosine which perform

as the actin-based motor; and (vi) MMPs (e.g. MMP-2, MMP-9, MMP-14), which mediate ECM

degradation. For individual podosome dot, it has an actin core enriched with F-actin, cortactin,

WASP/N-WASP, and Arp2/3 complex etc; and an ring structure composed of integrins, Src,

vinculin etc (150).

It is well known that podosomes function to degrade local extracellular matrix and

therefore promote invasion through underneath tissue boundaries. Less is known that podosomes

may also function to anchor a cell to the extracellular matrix, and to sense substrate rigidity and

transmit forces. Collin et al reported that motor protein myosin II formed circular structures at

the outside edges of the podosome actin ring to regulate podosome dynamics (153). They

proposed that podosomes are dynamic mechanosensors which interact with myosin tension and

actin network in living cells (153).

Fig 2-3


Podosomes Invadopodia

A B C

D

Cell body

Ruffles

InvadopodiaMatrix

26

Figure 2-3. Morphology of lamellipodia, podosomes and invadopodia.

(A) Lamellipodia is a broad actin-rich membrane extension at the leading edge of the cell; (B) Podosome is an actin-rich membrane inward protrusion of the ventral surface of the cell membrane; (C) Invadopodia is an actin-rich outward protrusion of the ventral surface of the cell membrane and it is usually formed at the peri-nucleus region; (D) Ruffle is an actin-rich outward membrane protrusion of the dorsal surface of the cell membrane and it is often located at the peripheral.


Cytosol

Membrane

Podosome structure

6 4 3 1 ezrin talin zyxin paxillin vinculin gelsolin N-WASP

Arp2/3 F-actin

Fig 2-4

27

Figure 2-4. Structure of podosome.

VASP Dynamin Cortactin

Podosomal ring

28

2.4 Invadopodia

Invadopodia are actin-based outward protrusions at the ventral surface of tumor cells and

transformed cells which mediate proteolysis of the extracellular matrix (Figure 2-3) (154). The

size of invadopodia varies from 0.1 µm to 0.8 µm in diameter and may reach 2-3 µm or greater in

length (155). Invadopida usually assemble into clusters around membrane invaginations

proximal to the Golgi complex in the cytoplasm (156). Invadopodia were first observed in the

oncogene v-Src transformed fibroblasts (157, 158). Invadopodia show stable once it formed and

prolonged protease secretion to degrade the extracellular matrix (159). Usually a cell can form

less number of invadopodia than podosomes.

Generally, invadopodia share similar molecular protein components with podosomes.

However, the same protein in different structure may play a distinct role. For example, cortactin

is commonly considered as a protein marker for podosome, although it does exist at invadopodia

as well. Cortactin as a regulator of the Arp2/3 complex is also of particular important in

invadopodia function (160). While most of studies have been on the possible role of cortactin in

actin assembly for direct formation of actin-rich invadopodia puncta (161), Emily et al found that

the primary role of cortactin in invadopodia is to promote protease secretion (160, 162). Thus

cortactin may link vesicular trafficking and dynamic branched actin assembly to regulate

protease secretion for invadopodia-associated ECM degradation (162).

2.5 Molecular components of podosomes

Podosomes are punctate actin-rich structures located at the ventral surface of the cell

membrane (163). A non-exhaustive list of component that has been identified in and around

podosomes as summarized above. Podosome formation is dependent on PKC, Src and

29

RhoGTPases (141, 142) and other kinases. I will introduce the roles of signal transduction

pathways involved in podosome assembly including Src, PI3k/Akt and MAPK in this chapter,

focusing their roles in cytoskeleton rearrangement, podosome/invadopodia assembly and cell

motility. The role of PKC in podosome formation and cell motility will be discussed in detail in

the next chapter.

2.5.1 Src

Src family kinases, as a family of cytoplasmic non-receptor protein tyrosine kinases, are

involved in various signaling pathways regulating diverse cell events such as cell adhesion,

migration, invasion, proliferation and cell cycle (164). The Src family consists of 8 members: c-

Src, Fyn, Lyn, Hck, c-Yes, Blk, Fgr and Lck. Many cell types express multiple Src family

kinases (165).

Src activation may lead to cytoskeleton reorganization. Podosomes are found as

organized F-actin rich structures on the ventral surface of v-Src-transformed cells (141, 166,

167). An inhibitory mutant of N-WASP impairs podosome formation as well as v-Src-induced

matrix degradation, indicating a probable link between control of actin assembly and the

formation of structures that mediate Src-induced matrix remodelling (168).

Src not only play a critical role in the process of podosome formation, but also regulate

podosomal dynamics, turnover rate, life span and proteolytic function. Podosome number and

the podosome-associated actin cloud were decreased in Src-/- osteoclasts (169). The life span of

podosomes in Src-/- osteoclasts was increased fourfold and that the rate of actin flux in the core

was decreased by 40%. Rescue of these cells with Src mutants showed that both the kinase

activity and either the SH2 or the SH3 domain are required for Src to restore normal podosome

30

organization and dynamics (169). When osteoclasts adhere onto bone surfaces, they condense

their actin-rich podosomes in tight belts to establish sealing zones (170). ARF (ADP-ribosylation

factor) GTPase-activating protein GIT2 (G protein-coupled receptor kinase-interactor 2), which

localizes to sealing zones, depends on Src phosphorylation for maintaining osteoclast polarity

(170). In smooth muscle A7r5 cells, the phosphorylation of Ser277 of actin filament associated

protein (AFAP) regulated podosome lifespan in a Src- and PKC -dependent manner (171).

Courtneidge et al showed that the adaptor protein tyrosine kinase substrate 5 or five SH3

containing protein (Tks5/Fish) was required for podosome/invadopodia formation, degradation

of ECM, and cancer cell invasion in vitro (148) and in vivo (172) . Recently they demonstrated

that Tks4, a protein contains an amino-terminal PX domain, four SH3 domains, and several

proline-rich motifs, is tyrosine phosphorylated and predominantly localized to rosettes of

podosomes in Src-transformed fibroblasts. In the absence of Tks4, MT1-MMP was not recruited

to the incomplete podosomes in these cells (173). Takenawa’s group reported that Src-expression

stimulated podosome formation at the sites of focal adhesion after PI(3,4)P2 accumulation in

NIH3T3 cells. The adaptor protein Tks5/Fish, which is essential for podosome formation, was

found to form a complex with Grb2 at adhesion sites in a Src-dependent manner. Further, it was

found that N-WASP bound to all SH3 domains of Tks5/Fish, which facilitated circular

podosome formation (174).

Src is involved in FGF induced activation of STAT5 during vascular endothelial

morphogenesis in mouse microvascular endothelial cell (175). The activated FAK forms a binary

complex with Src family kinases which can phosphorylate other substrates and trigger multiple

intracellular signaling pathways to regulate cancer cell migration, invasion, epithelial

mesenchymal transition (EMT) and angiogenesis (176). In Ras-mediated transformation, the

31

amoeboid movement of tumor cells in the three-dimensional matrix, and transmigration of tumor

cells through the mesothelial monolayer, mDia, is potentially linked to Rac activation and

membrane ruffle formation through c-Src-induced phosphorylation of focal adhesion proteins,

and ROCK antagonizes this mDia action (177). By activating Src-FAK,TGF integrates ErbB

receptor and integrin signaling to induce cell migration and survival during breast cancer

progression (178). By interacting with Src kinase, S100B, a Ca2+-binding protein of the EF-

hand type known as S100 that is abundantly expressed in astrocytes, contributes to reducing the

differentiation potential of cells and participate in the astrocyte activation in case of brain insult

and in invasive properties of glioma cells (179). In mouse embryonic fibroblast cells, Src-

Associated substrate during Mitosis of 68 kDa (Sam68) localizes near the plasma membrane

during cell attachment and serves as an adaptor protein to modulate Src activity for proper

signaling to small Rho GTPases during cell polarization and cell migration (180).

2.5.2 PI3K/Akt

The phosphoinositide 3-kinases (PI3K) are a family of enzymes that regulate diverse

biological functions in various cell types by generating lipid second messengers (181). PI3K can

phsophorylate the D3 position of the inositol ring of phosphoinositides. According to the

similarity of molecular structure, the PI3K family can be divided into three classes: (i) Class IA

PI3Ks, which can be activated by tyrosine-kinase-associated receptors, are heterodimeric

enzymes consisting of a regulatory subunit (p85 , p85 or p55 ) and a catalytic subunit (p110 ,

p110 or p110 ) (182), and Class IB PI3K, PI3K , which can be activated by G-protein-coupled

receptors, only contain one catalytic subunit p110 and one regulatory subunit p101 (182), (ii)

class II and (iii) class III (183). The class II PI3Ks harbor human PI3K-C2 , PI3K-C2 or

32

human HsC2-PI3K, and PI3K-C2 (184). The class III enzyme is a heterodimer of p150 protein

kinase and the mammalian homology of the yeast VSP34 PI3K (185).

PI3K is a major signaling component that can convert phosphatidylinositol-4,5-

biophosphate (PI (4,5)P2) to phosphatidylinositol-3,4,5-triphosphate (PI(3,4,5)P3) at the inner

leaflet of the plasma membrane (186). PI(3,4,5)P3 acts as a binding site for numerous

intracellular enzymes that contain pleckstrin homology (PH) domains (187), such as the

serine/threonine kinase Akt (188). In turn, PIP3 contributes to the recruitment and activation of a

wide range of downstream targets (189). Akt binds to PI(3,4,5)P3 via its N-terminal PH domain

(190, 191) and is activated by phosphorylation at T308 and S473 by PDK1 and putative PDK2,

respectively (192-194). Putative PDK2 were also identified as the mTOR/Rictor complex (195).

Activated Akt modulates a wide range of cellular functions, such as cell survival, proliferation,

polarity formation, and cortical actin regulation. Akt can modulate activities of multiple

downstream factors such as glycogen synthase kinase-3 (GSK3 ), BAD, caspase 9, and the

fork head transcription factors (196), through phosphorylation (197). Akt phosphorylates and

inhibits GSK3 activity, thereby leading to the stabilization of -catenin (198). The signaling

map for PI3K/Akt is shown in Figure 2-5.

PI3K/Akt signaling is critical for cytoskeleton rearrangement and podosomes. Interaction

of PI(4,5)P2 with gelsolin and WASP is critical for podosome assembly/disassembly and actin

ring formation in osteoclasts (199). PI(4,5)P2 enhances de novo actin polymerization by increase

Arp2/3 complex-induced actin nucleation through modulation of N-WASP (200). In fibroblast,

The first PH domain of AFAP-110 bound to PKC and further recruit c-Src to the site of

podosome, which was dependent on PI3K activitiy (201).

PI3K PI3Kp

PTENpp

pp

p

PIP2 PIP3

PDK1

PDK2

Akt

pp

p p

p

pp

p

T308

S473

Aktp

p

Cell growth Apoptosis Metabolism Proliferation

mTORp Bad

p

FKHRp

MDM2p

GSK3p

p21

p27

CyclinD1

Fig 2-5

33

Figure 2-5. PI3K/Akt signaling pathway.

34

2.5.3 MAPKs

Mitogen-activated protein kinase (MAPK) pathways constitute a large kinase network

that regulates a variety of physiological processes, such as cell differentiation, apoptosis and cell

adhesion, migration and invasion (202). Deregulation of MAPK activity has been implicated in

several pathological situations, including inflammation, oncogenic transformation, and tumor

cell invasion (202, 203). To date, several distinct groups of MAPK pathways have been

characterized in mammals (204): extracellular signal-regulated kinase (ERK)1/2, ERK3/4,

ERK5, ERK7/8, Jun N-terminal kinase (JNK)1/2/3 and the p38 isoforms / / (ERK6)/ (205-

208) (Figure 2-6).

The ERK pathway is activated by a large variety of mitogens and by phorbol esters,

whereas the JNK and p38 pathways are stimulated mainly by environmental stress and

inflammatory cytokines (203, 209, 210). MAPK cascades are organized as modular pathways in

which activation of upstream kinases by cell surface receptors leads to sequential activation of a

MAPK module (MAPKKK MAPKK MAPK) (Figure 2-6) (204). After MAPKs are activated

either in the cytoplasm or in the nucleus, they regulate transcription by modulating the function

of targeted transcription factors through serine/threonine phosphorylation (211). In addition to

the transcriptional effects, MAPKs regulate cell behavior also by phosphorylating cytoplasmic

target proteins, such as apoptotic or cytoskeletal proteins (212).

Fig 2-6

Stimulus

MAPKKK

MAPKK

MAPK

Transcription factors

Growthfactors

Stress, cytokines, growth factors

Stress, cytokines, growth factors

Stress, growth factors

A-Raf, B-Raf, c-Raf-1, Mos,

Tpl-2

MEK1/2

ERK1/2

Elk-1,Ets-2,RSK,MNK,

MSK,cPLA2

MEKK1/4,DLK,MLK1-4,LZK,

TAK1,ASK1,ZAK

MKK4/7

JNK1/2/3

Jun,ATF2,RNPK,p53,NFAT4, Shc

MLK3,TAK1,MEKK4,ASK1

MKK3/6

p38 / / /

Jun,ATF2,RNPK,p53,NFAT4, Shc

MEKK2/3

MEK5

ERK5

MEF2

35

Figure 2-6. MAPKs signaling pathway.

36

Accumulating data indicated that various signaling through activation of MAPKs to

control cell migration and invasion and MMPs. In vascular smooth muscle A7r5 cells, PKC-

mediated MEK/ERK/Caldesmon phosphorylation and translocation has been shown to remodel

actin stress fiber into F-actin-enriched podosome columns (213). MEK/ERK1/2/Caldesmon

signaling cascade regulated the lifetime and size of podosomes differentially (214). In murine

bone marrow-derived dendritic cells, the SH2 domain-containing leukocyte protein of 76 kDa

(SLP-76) adaptor protein mediated integrin engagement to facilitate podosome distribution

through ERK1/2 phosphorylation in the presence of chemokine (215). In bone marrow-derived

neutrophils, CDC42 regulated horizontal directional migration and vertical directional invasion

associated with podosome-like structures at the cell leading edge through ERK1/2 activity, while

CDC42GAP-induced p38 MAPK phosphorylation regulated directed migration by antagonizing

filopodia assembly (216). Big MAPK Erk5 may promote Src-induced podosome formation in

fibroblasts by RhoGAP7 and thereby limiting Rho activity (217). In murine fibroblast cell line

C3H10T1/2, oncogene v-Src transfection lead to MMP-2 expression through ERK signaling

pathway and transcription activation of Sp1 promoter (218). In Cos7 cells, concanavalin A

stimulated MMP-2 expression and activation through Src/ERK/p38 signaling pathway (219). In

human lung adenocarcinoma A549 cells, EGF stimulated Src kinase and JNK/ERK signaling

pathway through FAK at the cell membrane to increase MMP-9 expression and secretion (220).

2.5.4 Matrix metalloproteinases

Invadopodia and podosomes are able to degrade matrix proteins, such as fibronectin,

collagen and laminin (221). Matrix metalloproteinases such as membrane type-1 matrix

metalloproteinase (MT1-MMP, also called MMP-14), MMP-2 and -9, are thought to mediate

37

ECM degradation at sites of invadopodia extension (222, 223). In this section, firstly, I will

review the family members of MMPs, their molecular structures and the molecular mechanism

of regulation of MMP gene expression, activation and inhibition. Secondly, I will briefly

summarize the role of MT1-MMP, MMP-2/-9 in podosomes/invadopodia and cell motility.

The human MMP family includes 26 (and growing) members, including both secreted

and membrane-bound enzymes, characterized by their ability to degrade ECM and by their

dependence on Zn2+ binding for proteolytic activity (224, 225). MMPs are divided into five

major subclasses: collagenases (e.g. MMP-1,8,13), gelatinases (e.g., MMP-2, 9), stromelysins,

matrilysins, and membrane-type MMPs (e.g., MT1-MMP to MT6-MMP) (225, 226) (Figure 2-

7).

The activity of MMPs is regulated by three mechanims: transcription; secretion and

activation of the latent pro-enzymes; and inhibition by their endogenous inhibitors, the tissue

inhibitors of matrix metalloproteinases (TIMPs) (8, 226), which are able to inhibit

metalloproteinase activity by forming non-covalent complexes with active MMPs. Most MMPs

are secreted as inactive zymogens or pro-enzymes, and activation requires disruption of the Cys-

Zn2+ interaction (proteolytic removal of a prodomain) (227). Most pro-MMPs are thought to be

activated by tissue or plasma proteinases. Pro-MMP-2 activation is typically thought to take

place on the cell surface by MT1-MMP. Recently, it has been suggested that this activation

process requires both active MT1-MMP and the TIMP-2-bound MT1-MMP (228, 229).

Zn2+

Zn2+

Zn2+

Collagenase

MMP-1/8/13/18

Gelatinase

MMP-2/9

Membrane-Type

MMP-14/15/16/17

Fig 2-7

Signal/Prodomain Catalytic domain Hinge region Hemopexin

Transmembrane

Gelatin binding

PRCGVPD

RXKR

38

Figure 2-7. Molecular structure of MMPs.

39

The expression of many MMPs is transcriptionally regulated by growth factors,

hormones, cytokines, chemical agents (eg. phorbol esters, actin stress-fibre inducing drugs) and

cellular transformation (227). Phorbol esters have been shown to stimulate MMP- expression via

PKC in several systems (230-234). Up-regulation of PKC induces secretion of MMP-9 in

capillary endothelial cells (234), PKC-dependent NF- B activation is involved in MMP-9

induction in hepatocellular carcinoma cells (235), and inhibition of NF- B activity by synthetic

compounds inhibits MMP-9 secretion (236). Different isoforms of PKC has been implicated in

MMP-9 expression in various cell types (237-240). Phorbol 12-myristate 13-acetate (PMA)

induces MMP-9 expression via a PKC -dependent signaling cascade in BEAS2B human lung

epithelial cells (225).

MT1-MMP, MMP-2 and MMP-9 have been suggested to be the crucial MMP’s involved

in podosome/invadopodia matrix-degradation, and their regulation has been closely linked to

podosome formation and function (147, 241).

MT1-MMP (Membrane-Type 1- MMP) or MMP-14 is a membrane-bound

metalloproteinase with broad substrate specificity and multiple cellular functions (242). MT1-

MMP activates MMP-2 and possibly MMP-13, and may proteolytically modify CD44, V

integrin, and transglutaminase, therefore playing a regulatory role in cell migration (243).

Additionally, MT1-MMP itself has been shown to degrade multiple ECM components, including

collagen types I, II, and III; fibronectin; and laminins 1 and 5 (242-246). Protease inhibitor

studies have demonstrated that MT1-MMP is an enzyme crucial for gelatin matrix degradation in

the breast carcinoma cell line MDA-MB-231 (242). It has also been localized to invadopodia of

melanoma cells, and when it was overexpressed in RPM17951 human melanoma cells, cells

made contact with ECM, activated soluble and ECM-bound MMP-2, and degraded the ECM for

40

invasion (223). The localization of MT1-MMP to invadopodia is necessary for the invadopodia’s

degradative function, as in the absence of localization to invadopodia activated soluble MMP-2

failed to enable ECM degradation (223). Artym et al (159) suggested that 4 distinct invadopodial

stages in a stepwise model of invadopodia formation and function: (i) membrane-cortactin

aggregation at membranes adherent to matrix; (ii) MT1-MMP accumulation at region of

cortactin accumulation; (iii) matrix degradation at invadopodia region; and (iv) subsequent

contactin dissociation with foci of degraded matrix.

The gelatinase MMP-2 and 9 have gelatin-binding domains. Both have been implicated

in the angiogenesis and metastasis of cancer cells (247). MMP-2 (or Gelatinase A) and the

MMP-TIMP-2 complex have been found to bind and localize at invadopodia plasma membrane

of Src-transformed fibroblasts (248). MMP-2 is activated from its pro-MMP-2 form into active

MMP-2 by MT1-MMP at the cell surface (249). It has been suggested that invadopodia direct

localized degradation of the ECM by concentrating active membrane-associated collagenases at

sites of cellular invasion (248). It has been hypothesized that proteinase recruitment to

podosomes/invadopodia could involve trafficking of vesicles along microtubules (152), given

that, exocytosis of MMP-2 and MMP-9 in melanoma cells is microtubule-dependent (250).

Bound MMP-2 at invadopodia loses its propeptide, and thus becomes activated.

Previous reports suggest MMP-9 (or Gelatinase B) is overexpressed in response to injury,

and is linked to reepithelialization and early repair processes (87, 251, 252), whereas MMP-2 is

important during the extended remodeling phase (252, 253). In airway epithelium, MMP-9 is

expressed by human bronchial epithelial cells, and blocking MMP-9 activity results in inhibition

of the wound repair process (87, 251). Legrand et al found MMP-9 accumulated (via an actin-

dependent pathway) in the advancing lamellipodia of migrating cells at the leading edge of a

41

wound, and they speculated that the active form of MMP-9 was located at the digested area of

the coated extracellular matrix (87). MMP-9 has been implicated in migration of smooth muscle

cells (254), human eye corneal cells (255, 256), human skin epidermal cells (257), human

keratinocytes (258) and G8 mouse fetal myoblasts and KM201 and IM7.8 hybridomas (259).

MMP-9 has been shown to be involved in the migration of inflammatory cell types such

as macrophages, T-lymphocytes, and eosinophils (251, 260-264). MMP-9 is also a major factor

of human polymorphonuclear cell migration across the basement membrane and elastase

contributes to this process by activating pro-MMP-9 (262). Induction of keratinocyte migration

by EGF and HGF coincides with the induction of MMP-9 activity (264). Human bronchial

epithelial cell (HBEC) migration is a dynamic, stepwise process in which MMP-9 expression is

quickly and specifically regulated. Up-regulation of MMP-9 expression in migrating HBEC

might reflect a modification of the ECM in close contact with these cells. Additionally,

regulation of MMP-9 expression in migrating HBEC could be via cell-cell and cell-ECM

adhesion molecules and changes in cell shape (265). In migrating cells, cell-ECM adhesions are

provided by specialized regions of close contact, called primordial contacts (a type of cell-matrix

adhesion usually transiently formed at the advancing edge of a migrating cell), distributed at the

leading edge of the cell in the direction of migration (87, 266, 267). Primordial contacts of

migrating HBEC are characterized by a dense accumulation of actin filaments and the presence

of vinculin and type IV collagen (87, 261). These primordial contacts are very short-lived

structures. MMP-9 is localized and active at these cell-ECM contacts, blocking MMP-9

activation leads to cells remaining fixed on the previously established primordial contacts and

unable to migrate further (87).

42

2.6 Focal adhesion vs. podosome: what is the difference?

A variety of cell-matrix adhesions have been identified as focal adhesions, podosomes,

and invadopodia. These adhesion sites contain integrin clusters able to develop special structures,

which are different in their architecture and dynamics although they share almost the same group

of proteins (163). Here we compare the organization, dynamics and interplay between focal

adhesions and podosome, in order to understand how such subcellular sites - though closely

related in their composition - can be structurally and functionally different.

These two cell-ECM interactions both contain specific adhesion receptors named

integrins, cytoskeletal elements, and a wide variety of interconnecting adaptor proteins and

signaling proteins. Podosomes contain a ring of adhesive molecules centered on an actin column,

and their general orientation is perpendicular to the substrate and the plasma membrane. This

contrasts with the elongated structure of focal adhesions with a tangential orientation with

respect to the ECM. Dynamics and tension of both structures are also different, with podosomes

being more dynamic and instable as compared to focal adhesions. In all cases, alteration of their

dynamics results in modifications of cell differentiation and migration (268-270). These distinct

properties suggest specific functions: the most commonly proposed function is that podosomes

could be involved in matrix degradation and invasion, whereas focal adhesions are rather

associated with matrix remodeling such as fibronectin fibrillogenesis (271, 272). Podosomes

may be initiated from focal adhesions due to changes in the phosphorylation status of proteins

and in the composition of phosphoinositides on the plasma membrane (174).

2.7 Podosome vs. invadopodia: what is the difference?

43

Podosomes and invadopodia both are actin-rich membrane structures that form close

contact with the surrounding substrate, with dimensions ranging from 0.5 to several µm (273).

The similarities and distinctions between podosomes and invadopodia are under debate (Figure

2-3) (Table 2-1). Podosomes and invadopodia share many common features, including

appearance, components and similar proteolytic function (272). Many of the same signals

regulate both podosomes and invadopodia formation; for example, Src kinase, PKC, PI3K and

Rho GTPases (150). Moreover, the N-WASP-Cortactin-Dyn2-Arp2/3 complex is essential for

the formation of both structures and is believed to be the minimal machinery necessary for their

formation (141).

The three major differences between podosomes and invadopodia have been proposed as

location, depth and lifetime (272, 274). Podosomes most often form at the periphery of the cell,

while invadopodia are close to the nucleus and are associated with Golgi complex, allowing for

the prolonged secretion of MMPs. It is generally accepted that podosomes are typically 0.5-2 µm

(depth) inward invaginations into the cells, while invadopodia are 2-10 µm (depth) outward

protrusions into the ECM. Podosome have a shorter lifetime (approximately 2-10 min), while

invadopodia have a much longer lifetime (hours). Given the short lifetime of individual

podosomes, they are only able to degrade ECM within their immediate vicinity. Comparatively,

invadopodia are more persistent and are more likely to be found in cancer cells invading across

tissue boundaries. It has been suggested that the short-lived podosomes can probe the substrate

for sites of attachment before maturing into invadopodia (159, 274, 275). The morphology of

podosomes can be dots, rosettes and belts, which may indicate different stages of maturation.

44

Table 2-1. Comparison between podosomes and invadopodia in 2D vs. 3D cultures

2D-Glass Coverslip Conditions 3D- Matrix Conditions Podosomes Invadopodia Podosomes Invadopodia

Location Cell periphery, ventral surface

N/A Cell periphery, ventral surface

Close to nucleus, close to Golgi

Protrusiondirectionand depth

Necessarily inward invaginations; 0.5-2 µm (depth) inward

N/A Both inward invaginations and outwardprotrusions0.5-2 µm (depth)

Outwardprotrusions;2-10 µm(depth)outward

Lifetime 2-10 min N/A 2-10 min Hours

45

The current debate surrounds the issue of whether these differences are real or if they are

due to various culture and assay conditions. Some authors have suggested podosomes to be

artificial cellular structures due to cell culture conditions (272). Podosomes are inward

invaginations of the ventral surface cell membrane when cells are cultured on glass surfaces in

the common 2D culture system. In the 3D culture system with cells being cultured on matrix gel,

invadopodia are typically seen as outward protrusions of the ventral surface cell membrane into

the ECM. However, podosomes with both inward and outward protrusions were seen in a recent

study in leukocyte cells on tissue slides, imaged by electron microscopy (276-278).

Yamaguchi and Condeelis also described podosomes to be outward protrusions of the

ventral surface cell membrane into the ECM when cells are cultured in 3D conditions, and thus

may represent an immature form of invadopodia (274, 279). In our view, we consider podosomes

to be dynamic, transient precursors to invadopodia formation, on the “lower end” of the

continuum of ECM-degrading structures. Podosomes may be responsible for “tasting” the ECM

substrates available, and temporarily degrading matrix proteins. Should conditions call for

invadopodia formation, these podosomes may transform into full-fledged invadopodia, capable

of continuous and stronger activation of MMPs, more intense digestion of the ECM, and cellular

invasion. The molecular mechanisms that regulate the switch from podosomes further down the

continuum into invadopodia are largely unknown (280), and could play a significant role in the

initiation and regulation of tumor invasion and metastasis.

In summary, in this chapter, I reviewed the cellular structures involved in cell motility

such as focal adhesion, podosome and invadopodia for cell adhesion, cell migration and cell

invasion; I summarized the molecular component proteins and signaling transduction pathways

46

involved in podosome assembly such as Src, PI3K/Akt and MAPK. The importance of PKC in

podosome formation and cell motility is discussed in greater details in the next chapter.

47

3. Chapter Three. General background - Protein kinase C family in cytoskeleton

rearrangement and cell motility

Part of the content of this chapter has been prepared as a review article:

Xiao H, Liu M. PKC in cell motility. (In preparation)

48

3. Chapter Three. General background - Protein kinase C family in cytoskeleton

rearrangement and cell motility

Protein kinases C (PKC) play a vital role in cell migration and invasion in various types

of cells. Accumulating evidence show that PKCs are highly involved in the process of

podosomes and invadopodia formation. PKCs are also important in human lung physiology and

alteration of PKCs is linked to lung disorders. In this section, firstly, I will introduce PKC family

and review their subfamily members, molecular structure and the mechanism of PKC activation

and inactivation. Secondly, I will summarize the role of each PKC subfamily in cell migration

and invasion.

3.1 Introduction of PKC family

PKC is a family of serine/threonine kinases implicated in the transduction of signals

coupled to receptor-mediated hydrolysis of membrane phospholipids (281). These kinases

transduce signals involved in short-term processes (such as ion fluxes (282), neurotransmitter

release (283)), mid-term process (such as receptor modulation (284)), and long-term processes

(such as cell proliferation (285), synaptic remodeling (286, 287) and gene expression (288)).

Various external signals such as growth factors, neurotransmitter, and hormones, stimulate the

hydrolysis of inositol phospholipids and initiate formation of the second messenger

diacylglycerol (DAG) to mediate PKC activation (289, 290).

According to sequence homology and sensitivity to activators, at least 10 isoforms

encoded by 9 different genes have been described (Figure 3-1 and 3-2) (291). The various PKCs

are grouped into three subfamilies: (i) classical or conventional PKCs ( , I, II and ); (ii) novel

49

PKCs ( , , and ); and (iii) atypical PKCs ( and / ) (292). Except these three subfamilies,

there is PKCµ or protein kinase D1 (PKD1) (293).

3.1.1 Molecular structure of PKC family

The general structure of the different PKCs includes conserved domains (C1-C4)

separated by variable sequences (V1-V5) (Figure 3-1) (294). C1-C2 represents the regulatory

portion of each enzyme, where the specific activators interact with, while C3-C4 form the

catalytic region responsible for both the substrate binding and the activity (294). The catalytic

domain is characterized by a high degree of homology among the various kinases (295). The C1

domain bears one (in atypical) or two (in conventional and novel) cysteine-rich domains (294,

296), located near the amino-terminals region, which represent the docking-sites for

phosphatidylserine (PS) (297) and the physiological activator diacylglycerol (DAG) as well as

for the analogous phorbol esters (i.e. PMA, PDBu (phorbol-12,13-dibutyrate)) (298). The

atypical isoforms are sensitive and activated by PS and also ceramide and phosphatidylinositol-

3,4,5-trisphophate (295, 299). The C2 region contains the Ca2+ binding site (295, 300) and it is

present in the convetional isoenzymes. The nPKCs are not responsive to Ca2+ because of the

absence of amino acids residues essential for the Ca2+ binding in the C2-like domain; recruitment

to membrane is dependent on a C1 region with higher affinity for phospholipids in comparison

with cPKCs (295, 300, 301). C3 bears the ATP binding lobe, while C4 contains the substrate

docking sequence (294).

Structure of the PKC isozymes and PKC-related kinases

Classical PKCs

( , I, II, )

Novel PKCs

( , , , )

Atypical PKCs

( , / )

PKCµ/PKD

PS C1a C1b C2 C3 Kinase

DAG, phorbolester binding

Ca2+

bindingATP

binding

PS C1a C1bC2-like C3 Kinase

PS C1 C3 Kinase

TM C1a C1b PH C3 Kinase

N C

N

N

N

C

C

C

P P P

activation loop

turnmotif

hydrophobic motif

T500 T641S660

Regulatory domain Catalytic domain

P P P

T505 S643S676

P

T410

P P

T560E

P

S744

P P

S748 S916

Fig 3-1

PB1

50

Figure 3-1. Molecular structure of PKCs.

51

3.1.2 Mechanism of PKC activation

All known PKCs, at the N-terminal side, are characterized by a pseudosubstrate or

autoinhibitory region, adjacent to C1, which keeps each isoform in an inactive conformation

(Figure 3-3) (295, 302). It has been proposed that mature and phosphorylated cPKCs can be in

the cytosol and kept in an inactive conformation by intramolecular interactions between the N-

terminal pseudosubstrate region and the kinase domain. The interaction with the specific

activators allows the activation of the enzyme by opening its folded conformation: these

cofactors are in fact able to decrease the affinity of the pseudosubstrate domain for the catalytic

site that can exert its phosphorylating activity (295). The latent kinase can be recruited to the

membrane and activated by lipid ligands (such as phorbol ester), leading to a massive

conformational change that releases the pseudosubstrate domain from the substrate-binding site,

allowing for substrate binding, phosphorylation and activation of downstream signaling effectors

(302, 303). This activation mechanism is related to the translocation of PKCs to different

intracellular sites, and the subsequent phosphorylation of specific substrate by the translocated

enzyme (292). The translocation and activation of PKC from one to another subcellular

compartment was thought to reflect only the interaction between PKC and lipids (292, 304).

However, several proteins can interact with inactive and active PKCs, dictating enzyme location

in both basal and stimulated conditions, underlining the additional relevance of protein-protein

interactions in PKCs homeostasis (304).

PKC phosphorylation sites in the catalytic domain

C3 Kinase C

P P P

Ser579 FEGFEYVNSer574 FEGFEY I N

Thr560 VQLTPDDEThr555 VQLTPDDD

Thr410 TTSTFCGTPNYThr403 TTSTFCGTPNY

mPKChPKC /

Atypical

Ser662 FHGFSFVNSer729 FKGFSYFGSer674 FRNFSYVSSer695 FSNFSF I N

Ser643 PQLSFSDKThr710 P I LTVDEAThr555 PVLTP I DESer676 PRLSFADR

Thr505 PPSTFCGTPDYThr566 TTTTFCGTPDYThr512 TTATFCGTPDYThr538 KTNTFCGTPDY

mPKCmPKChPKCmPKC

Novel

Ser657 FEGFSYVNSer662 FAGFSYTNSer660 FEGFSFVNThr674 FQGFTYVN

Thr638 PVLTPPDQThr642 VELTPTDKThr641 PVLTPPDQThr655 PALTPPDR

Thr497 TTRTFCGTPDYThr500 TTKTFCGTPDYThr500 TTKTFCGTPDYThr514 TTRTFCGTPDY

mPKChPKC IhPKC IImPKC

Classical

Hydrophobic motifTurn motifActivation loop

Fig 3-2

52

Figure 3-2. The serine/thronine phosphorylation sites of PKCs in the catalytic domain.

- no- noAtypical

- noyesNovel

yesyesClassical

Ca2+DAGPKCs

C1

C3 C4C2Inactive

PKCs

C1 C3 C4C2Active PKCs

Cysteine-rich repeat

DAG PS Ca2+

Ca2+

DAG

Phospholipase C

A

B

Fig 3-3

53

Figure 3-3. Molecular mechanism of PKCs activation.

54

3.2 The role of PKCs in cell motility

3.2.1 Classical PKCs in cell motility

Classical or conventional subfamilies of PKC contain four members, , I, II, and

(289, 305). These Ca2+-dependent cPKC isozymes are activated by Ca2+, DAG, and

phosphatidylserine (PS) (289). Additional endogenous activators include cis-unsaturated fatty

acid (306) and lysophosphatidyl choline (lysoPC) (307).

In general, classical PKC activation seems to positively affect cell motility, invasion and

metastasis (292). Overexpression or stimulation of PKC generally plays a major role in

enhancement of cell migration in multiple cancer cell types. For example, early studies showed

that over-expression of PKC in MCF-7 breast cancer cells leads to increased anchorage-

independent growth, tumorigenicity and metastasis in mice (308). Introduction of the ERBB2

receptor into MDA-MB-435 breast carcinoma cells confers increased invasiveness through Src-

dependent activation of PKC (309). Overexpression of PKC II in rat intestinal epithelial cells

increased cell invasiveness, possibly through increased Ras and MEK activation (310). One

means through which classical PKC might enhance invasion and metastasis is the regulation of

integrins (292). PKC has been shown to directly interact with 1-integrin and increase

migration on 1-integrin substrates, possibly through regulation of internalization and recycling

of 1-integrin in human breast cancer MCF-7 and MDA-MB-231 cells (311). Furthermore,

disruption of the PKC - 1-integrin association blocks chemotactic migration in human breast

carcinoma cells (MCF-7 and MDA-MB-231) and in the Phoenix amphotropic retroviral

packaging cells (312). In human colon adenocarcinoma cell lines, a high level of PKC

expression together with a low level of E-cadherin was strongly related to a high migratory

55

activity (313). Similarly, The motility and invasion of low-metastatic rat prostate AT2.1 tumor

cells were increased by thymelea toxin, a selective activator of PKC over PKC (314).

Besides the above mentioned cancer cells, classical PKCs are also critical for cell

migration of other cell types. In GT1 hypothalamic neurons, PKC signaling pathway promoted

morphological differentiation via an increase in -catenin, a cell-cell adhesion molecule (315).

PKC promotes GT1 neuronal migration by activating focal adhesion complex proteins such as

p130Cas and FAK (315). In rat embryo fibroblasts (REF52 cells), PKC redistributed to the

leading lamellopodia of cells stimulated to migrate into an artificial wound (316). In human T

lymphocyte cell line HUT-78, and freshly isolated normal human T lymphocytes activated by

PMA or purified protein derivative prepared from mycobacterium tuberculosis, inhibition of

classical PKC activity by Gö6976 suppressed lymphocyte polarization and migration following

CD44 ligation, which suggested that PKC was important for lymphocyte motility (317).

However, there are also some evidences that PKC may negatively regulate cell

migration in certain conditions. In bronchial epithelial BEAS2B cells, hog barn dust added to the

entire culture medium slows airway epithelial cell migration through a PKC -dependent

mechanism in a wound healing assay (27). In MDA-MB-231 breast cancer cells, PKC activation

via PMA treatment inhibits EGF-induced cell spreading, the initial event of motility and

chemotaxis (318). Among the five PKC isoforms ( , , , and ) identified in this cell line,

PMA treatment only induced PKC translocation from the cytosol to the membrane, an event

that correlated with the development of the rounded morphology (318). One possible explanation

of these inhibitory effects is the asymmetrical stimulation of cells by phorbol ester added to the

cell culture medium. Therefore, the directioned cell migration is inhibited by enhanced

locomotion.

56

3.2.2 Novel PKCs in cell motility

Novel PKCs contain , , and (319). nPKCs also have twin C1 domains and a C2-like

domain which in the case of nPKCs, precedes the C1 domain in their N-terminal regulatory

regions (320, 321). The C2-like domain sequences of nPKCs lack calcium-co-ordinating acidic

residue side chains (320, 321). Hence nPKCs are activated by DAG/phorbol ester, without

requiring calcium (320, 321).

Similar to classical PKCs, novel PKCs, especially PKC , are also vital for cell migration

and invasion in many cancer cells. In renal carcinoma cell line CCF-RC1, PKC regulated tumor

cell migration by affecting the expression and activity of integrin 1 subunit and FAK (322).

PCPH/ENTPD5 (Pcph proto-oncogene protein or ectonucleoside triphosphate

diphosphohydrolase 5) is not expressed in normal prostate, but its expression increases along

cancer progression stages, being detectable in benign prostatic hyperplasia, highly expressed in

prostatic intraepithelial neoplasia, and remaining at high levels in prostate carcinoma. The

expression level and/or mutational status of PCPH/ENTPD5 contribute to the invasiveness of

prostate cancer cells through a mechanism involving PKC (323). KITENIN (KAI1 COOH-

terminal interacting tetraspanin) recruits Dishevelled (Dsh, a family of proteins involved in non-

canonical Wnt signaling pathways) and PKC to form a functional complex, which acts as an

executor in regard to cell motility, and thereby controls colorectal cancer cell invasion to

contribute to promoting metastasis (324). Quercetin (QUE; 3,5,7,3',4'-tetrahydroxyflavone)

inhibits tumor invasion via suppressing PKC /ERK/AP-1-dependent MMP-9 activation in breast

carcinoma cells (325). Sorafenib (Bay43-9006; Nexavar) is a Raf and VEGF receptor inhibitor

that blocks receptor phosphorylation and MAPK-mediated signaling (326). Coadministration of

Sorafenib with novel PKC inhibitor Rottlerin potently inhibits cell migration in human malignant

57

glioma cells (326). In EGFR-overexpressing invasive cells such as MDA468 breast cancer cells,

inhibition of MAPK activity with MEK inhibitor PD98059 blocks early stages of cell migration

(up to 4 h) (327). While inhibition of PKC activity with Rottlerin or dominant-negative PKC

expression blocks sustained cell migration after 4 h and up to 12 h, the combination of MAPK

and PKC inhibitors completely blocked TGF- -induced cell migration (327).

Novel PKCs are also important for cell motility of mesenchymal cells such as smooth

muscle cells (SMCs), fibroblasts, astrocytes and other cell types. In rat aortic SMCs, upon

treatment of PDGF-BB or TGF- 1, a fraction of PKC rapidly translocated from the cytosol to

the post-nuclear particulate fraction at 15 sec and reached an apparent maximum at 30 min,

which is important in the control of PDGF-BB or TGF- 1 stimulated vascular smooth muscle

cell migration (328). In response to mechanical stress, SMCs from PKC -/- mice showed an

abnormal cytoskeleton structure, which was related to a diminished phosphorylation of paxillin,

FAK, and vinculin (329). Mechanical stress enhanced SMC migration was diminished in these

PKC -/- SMC cells, which indicates that PKC is a key signal transducer between mechanical

stress and cell migration (329). In SMCs, PKC , and were expressed and they were activated

upon integrin engagement with different kinetics. PKC was activated early, whereas PKC and

PKC were activated later (330). Activation of PKC was necessary for cell attachment to

fibronectin (330).

In mouse fibroblasts, EGF stimulates myosin light chain phosphorylation, a marker for

contractile force, concomitant with PKC activation (331). Myosin-based cell contractile force is

considered to be a critical process in cell motility. The dominant-negative PKC construct or

PKC RNAi abrogated EGF-induced cell contractile force generation and motility through

dephosphorylation of myosin light chain (331). In primary human dermal fibroblasts, during cell

58

migration PDGF-BB strongly stimulates membrane translocation and leading edge clustering of

PKC (332).

In rat brain astrocytes, activation of ERK1/2 by a PKC -dependent event mediated

through Elk-1 pathway is essential for MMP-9 gene up-regulation and cell migration induced by

bradykinin (333). Phosphoprotein enriched in astrocytes-15 kDa (PEA-15), enriched in

astrocytes, also inhibits glioblastoma organotypic astrocyte cell migration in a PKC dependent

mechanism (334).

In human eosinophils, PKC mediates cell motility, CD11b expression and MMP-9

granule release during migration (335). Engagement of CD44 using immobilized mouse

antibodies or hyaluronan-enriched extracellular matrix lattices induces active migration in T

lymphocytes accompanied by cytoskeletal rearrangement and cell polarization. PKC inhibitor

Rottlerin reduced CD44-activated cell migration but did not completely ablate it (317).

In addition to PKC , other novel PKCs such as , and also play an important role in

cell migration. Sphingosine 1-phosphate (S1P) signals through S1P(1) and G(i) to activate PKC

and, subsequently, a PLD2-PKC -Rac1 cascade is necessary to stimulate the migration of human

lung endothelial cells during the angiogenic process (336). Expression of PS , a peptide inhibitor

based on the PKC pseudosubstrate sequence, slowed the rate of PMA induced endothelial cell

migration (337). In an in vitro three-dimensional assay in which endothelial cells organize into

capillary tubules, the endothelial cells that expressed PS formed fewer such tubules (337).

On the other hand, there are also some reports that PKC may inhibit cell migration

under certain conditions. PKC -mediated increased cell substrate adhesion as a limiting factor

for TNF- stimulated neutrophil transendothelial migration (338). In freshly isolated bovine

aorta endothelial cells, lysoPC, a major lipid constituent of oxidized low-density lipoprotein, can

59

activate PKC and inhibit cell migration in wound healing assay (339). Phosphorylated PKC

associated with and phosphorylated syndecan-4 (339), which increased -actinin binding to the

variable region of syndecan-4 and inhibited the ability of syndecan-4 to bind and activate PKC

(339). The combined effects led to dissociation of the focal adhesion complex and inhibition of

endothelial cell migration (339).

3.2.3 Atypical PKCs in cell motility

The atypical PKCs, PKC and PKC / , have four functional domains: a PB1 domain at

the N-terminus, a pseudosubstrate (PS) sequence, a C1 domain consisting of a single Cys-rich

zinc-finger motif, and a kinase domain in the C-terminus (340). Since the C1 domain of aPKC

isotypes lacks the repeat structure found in those of cPKCs and nPKCs, aPKCs do not respond to

DAG and phorbol esters directly (341). Thr410 in the activation loop of aPKCs is

phosphorylated by PDK1 which binds to the hydrophobic motif (340). aPKCs are associated

with Par-3, Par-6 and Cdc42 in a complex that plays an important role in cell polarization (342).

PKC can be tyrosine phosphorylated by the non-receptor tyrosine kinase c-Src in PC12 cells

(343). aPKC plays a critical role in human lung cancer cell growth (344) and is an oncogene in

human non-small cell lung cancer progression (345).

Interestingly, PKC plays a major role in cell motility in hematopoietic cell linage, stem

cells and progenitor cells. Sialomucin endolyn (CD164) is an adhesion receptor that regulates the

adhesion of CD34+ cells to bone marrow stroma and the recruitment of CD34+CD38(lo/-) cells

into cycle. It modulated CXCL12-mediated migration of umbilical cord blood CD133+ cells

through PKC and Akt signaling (346). SDF-1 and its receptor (CXCR4) play a major role in

migration, retention, and development of hematopoietic progenitors in the bone marrow.

60

Overexpression of PKC in leukemic pre-B acute lymphocytic leukemia G2 cells and U937 cells

led to increased directional motility to SDF-1 (347). SDF-1 triggered PKC phosphorylation,

translocation to the plasma membrane, and kinase activity (347). SDF-1-induced proliferation

and MMP-9 secretion also required PKC activation. PI3K was identified as an activator of

PKC , and Pyk-2 and ERK1/2 as downstream targets of PKC (347). In vivo engraftment of

human CD34+ enriched cells to the bone marrow of NOD/SCID mice was PKC dependent.

Injection of mice with inhibitory PKC pseudosubstrate peptides resulted in mobilization of

murine progenitors (347). Knockdown of PKC by small interference RNA (siRNA) impaired

CSF-1-induced chemotaxis of human acute monocytic leukemia THP-1 cells and impaired

migration of mouse peritoneal macrophages (348). In human peripheral monocytes, and murine

macrophage-like cell line J774.1, PKC is an essential for transducing the motility signal induced

by superoxide and a chemotactic peptide, fMLP. In these cells, PKC was activated to

phosphorylate RhoGDI-1, which liberated RhoGTPases, leading to their activation. These events

were inhibited by myristoylated PKC peptides in these cells (349).

PKC is also involved in cancer cell migration and invasion. In human breast cancer

MDA-MB-231, MCF-7 and T47D cells, EGF induced PKC translocation from the cytosol to the

plasma membrane and activation of PKC probably via PI3K. PKC is an essential component of

EGF-stimulated chemotactic signaling pathway in these human breast cancer cells (350). In

human pancreatic adenocarcinoma cells, PKC plays a critical role in maintaining a high linear

motility score in motile subclones. In motile cells, PKC is constitutively associated with the

plasma membrane, whereas in nonmotile cells, PKC is totally excluded from the plasma

membrane (351). In a preclinical astrocytome model, PTEN deficiency resulted in a marked

increase in cell invasiveness that was specifically suppressed by inhibitors of PKC (352).

61

PKC may also be critical for cell motility in many other cell types. In human pulmonary

artery endothelial cells, PKC play a critical role in sphingosine 1-phosphate potently stimulated

endothelial cell migration through a PLD2-PKC -Rac1 cascade (336). Human

immunodeficiency virus-1 (HIV-1) envelope glycoprotein gp120 induces toxicity and alters

expression of tight junction proteins in human brain microvascular endothelial cells (353). gp120

can also cause dysfunction of blood-brain barrier via PKC related signaling pathways and

mediated intracellular calcium release leading to cytoskeletal alterations and increased monocyte

migration (353). During cell migration of primarily mouse fibroblasts, human water channel

aquaporin-9 (AQP9) induced actin polymerization in the filopodia extension of lamellipodia is

augmented by activation of PKC (354). In MC3T3-E1 osteoblast-like cells, insulin-like growth

factor-1 stimulated cell migration in a PKC dependent manner (355). In primary human

mesangial cells, connective tissue growth factor (CTGF) stimulated cell migration is asscoaited

with a PKC -GSK3 signaling axis (356). CTGF induced cell migration and cytoskeletal

rearrangement through the phosphorylation and translocation of PKC to the leading edge of

migrating cells (357). Inhibition of CTGF-induced PKC activity with a myristolated PKC

inhibitor or transient transfection of human mesangial cells with a PKC kinase inactive mutant

(dominant negative) expression vector led to a decrease in CTGF-induced migration (357).

In terms of cell invasion, PKC mainly enhances cell invasion into the underneath matrix

through up-regulation of MMPs. PKC induced phenotypic alterations associated with malignant

transformation and tumor progression in mammary cells (358). The stable overexpression of

PKC in immortalized mammary epithelial NMuMG cells, activated the ERK pathway,

enhanced clonal cell growth and exerted profound effects on proteases secretion (358). PKC

overexpression markedly altered the adhesive, spreading, and migratory abilities of mammary

62

epithelial NMuMG cells (358). Ursolic acid (UA), a constant constituent of Rosmarinus

officinalis extracts, is a triterpenoid compound which has been shown to have antioxidant and

anticarcinogenic properties (359). UA was able to reduce IL-1 or TNF- -induced rat C6 glioma

cell invasion through suppressing the association PKC with ZIP/p62 and downregulating the

MMP-9 expression (359). During glioma cell invasion through the brain extracellular matrix,

PKC regulated transcription of the MMP-9 gene induced by IL-1 and TNF- in glioma cells via

NF- B (237).

Powell et al. found that PKC mRNA levels were reduced markedly in metastatic

Dunning R-3327 rat prostate tumors relative to the nonmetastatic Dunning H tumor and normal

rat prostate (360). They established stably transfected PKC in Dunning R-3327 MAT-LyLu rat

prostate tumor cells. Nine independent clones of PKC -expressing cells exhibited a lower

tendency to metastasize to lungs relative to vector-transfected cell clones, and the ability of four

PKC overexpressing MAT-LyLu cell clones to invade through Matrigel in a Boyden chamber

assay was greatly reduced (361). These results contradict with most observations of the role of

PKC in promoting cell motility. Whether this is specific to this type of prostate tumors in rats,

or indicating the different function of PKC in vivo need to be further investigated.

3.2.4 PKCµ in cell motility

PKCµ or PKD1 is a serine/threonine protein kinase, which contains two cysteine-rich

domains that bind diacylglycerol or phorbol esters, but it lacks the Ca2+ binding domain found in

cPKCs (362). PKCµ also has a pleckstrin homology (PH) domain that regulates its kinase

activity, but does not harbor the typical PKC autoinhibitory pseudosubstrate motif (362).

Accumulating evidences showed that PKCµ plays a very important role in cell cycle, cell

63

division, and cell proliferation. However, whether PKCµ is involved in cell migration is largely

unknown. Cortactin, paxillin and PKCµ were co-immunoprecipitated as a complex from

invadopodia-enriched membranes of invasive breast cancer MDA-MB-231 cells (363). In

contrast, this complex of proteins was not detected in lysates from non-invasive cells that do not

form invadopodia (363).

In summary, in this chapter, I reviewed the molecular structures of PKC family protein

kinases and the molecular mechanisms of their activation. I also summarized their distinct roles

in the regulation of cytoskeleton reorganization and cell motility. Most studies focused on the

role of single PKC isoform. Less evidences showed the interaction and crosstalk between

different PKC isoforms in cell motility. My thesis studies focus on coordinated activation of

PKC isoforms and relationship with other signaling pathways in podosome formation and

functions in normal human bronchial epithelial cells.

64

4. Chapter Four. Rationale, hypothesis and specific aims

65

4. Chapter Four. Rationale, hypothesis and specific aims

4.1 Rationale

The airway epithelial cells not only act as a protection from the external environment, but

also have a variety of biological functions including fluid and ion transport, mucus secretion and

ciliary transport, interaction with and/or recruitment of inflammatory cells, antimicrobial

activities, protection against oxidant and proteases, and modulation lung repair after injury (364,

365). Human lung bronchial epithelium is the primary target of lung diseases such as chronic

obstructive pulmonary disease, asthma, cystic fibrosis and lung cancer. Lung airway epithelial

cells are also important components of host defense (366). Epithelial cell movement over the

damaged matrix is a complex phenomenon including adhesion and deadhesion of cells to matrix

components and response to “migration-inducing” substances (364). Bronchial epithelial cells

express a variety of integrin and non-integrin receptors that can mediate their adhesion to the

extracellular matrix (367, 368). Cell migration is the most critical and first event occurring

during the epithelial repair process (7, 87).

Podosome is an actin-rich structures involved in cell migration and invasion across the

tissue boundaries (369). It was mainly found in mesenchymal cells and hematopoietic linage

cells, such as monocytes or macrophages (143-145), dendritic cells (370, 371), natural killer

cells (372), and osteoclasts (373, 374). It was also found in certain malignant metastatic invasive

cancer cells (148). Whether normal epithelial cells, in particular, normal human bronchial

epithelial cells can form podosome was largely unknown.

Phorbol esters such as PMA and PDBu, are often employed in biomedical research to

activate PKC family (375). PKC can mediate non-receptor tyrosine kinase c-Src activation and

subsequent podosomes formation in aorta endothelial cells (376) and vascular smooth muscle

66

cells (377). Several signaling proteins including PI3K/Akt and Src are described to serve as

downstream PKC effectors (375). However, which PKC isozyme(s) are involved in the process

of podosome formation and how do they regulate the proteolytic function to degrade

extracellular matrix are largely unknown. Whether Src tyrosine kinase and PI3K/Akt signaling

pathways are associated with PDBu-induced podosome assembly and how do these molecules

regulate the proteolytic function of podosomes in lung epithelial cells are still unclear. Phorbol

ester can induce dramatic cytoskeletal structure changes in lung epithelial cells (378). I speculate

that some of these structures are podosomes in lung epithelial cells and may be involved in

epithelial cell migration and invasion during lung physiological and pathophysiological

conditions.

4.2 Hypothesis

Normal human bronchial epithelial cell migration and invasion may be mediated by

podosome, and the formation and proteolytic function of podosomes are regulated by multiple

signal transduction pathways (including PKC, PI3K/Akt, Src and MAPKs) in a coordinated

fashion.

4.3 Specific aims

In my PhD thesis research work, I acquired primary normal human bronchial epithelial

cells and normal (non-cancerous) human lung bronchial epithelial BEAS2B cells and challenged

these cells with PDBu to activate PKCs. My hypothesis was tested in the following specific

aims:

67

1. To determine whether phorbol ester can induce podosome formation in primary normal (non-

cancerous) human lung bronchial epithelial cells and to characterize these molecular

structures (Chapter 6)

2. To identify which PKC isozyme(s) may contribute to podosome assembly and which PKC

isoform(s) control the proteolytic function in human lung bronchial epithelial cells (Chapter

7)

3. To determine whether PI3K/Akt/Src/MAPKs signaling pathway(s) are involved in regulating

podosome formation and proteolytic function in human lung bronchial epithelial cells

(Chapter 8).

68

5. Chapter Five. Materials and methods

69

5. Chapter Five. Materials and methods

5.1 Reagents and antibodies

Bisindolylmaleimide I (BIM I), Ro-31-8220, Gö6976, Rottlerin, BAPTA/AM,

myristoylated PKC pseudosubstrate (PS) and scrambled pseudosubstrate as its negative control

(PI), LY294002, Wortmannin, PP2, PP3, Su6656, GM6001, GM6001NC, Genistein, SP600125,

PD98059, SB20358 and Akt inhibitor II were from EMD Biosciences (Darmstadt, Germany).

Antibodies against different PKC isozymes and phosphorylated PKCs were from Cell Signaling

(Danvers, MA). siRNA targeting PKC , PKC , PKCµ, PKC , MMP-2, MMP-9, MMP-14,

control siRNA and fluorescein conjugated control siRNA, anti-integrin, anti-FAK, and mouse

monoclonal MMP-9 neutralization antibody were from Santa Cruz Biotechnology (Santa Cruz,

CA). Anti-vinculin, anti-actinin, anti-talin and anti-tubulin antibodies were from Sigma. Anti-

phospho Src Y416, anti-phospho PI3K p85 T458/p55 T199, anti-phospho PKD1 S241, anti-

phospho Akt S473, anti-phospho Akt T308, anti-phospho GSK3 S9, anti-phospho JNK2/3 p54

T183/Y185 and JNK1 p46 T183/Y185, anti-phospho p38 MAPK T180/Y182, anti- phospho

p44/p42 MAPK (ERK1/2) T202/Y204, anti-phospho ERK1/2 and anti-phospho PI3K p85

antibodies were from Cell Signaling (Danvers, MA). Anti-phospho-tyrosine (4G10), anti-PI3K

p85 and anti-cortactin (p80/85, clone 4F11) antibodies were from Upstate (Billerica, MA).

Anti-MT1-MMP, anti-MMP2, and anti-MMP9 antibodies were from Biomol (Plymouth

Meeting, PA). Anti-GAPDH antibody was from Upstate (Billerica, MA). Horseradish peroxidase

(HRP)-conjugated goat anti-mouse or anti-rabbit secondary antibodies were from Amersham

Pharmacia Biotech (Piscataway, NJ). Alexa Fluor 594 labeled secondary antibody, FTIC labeled

secondary antibody, Oregon green 488 phalloidin, rhodamine phalloidin, tetramethylrhodamine

isothiocyanate (TRITC), and Hoechst dye 33342 were from Molecular Probe (Eugene, OR).

70

pEGFP C2 vector, pEGFP-PKC and pEGFP-PKC plasmid constructs were gifts from Dr. Peter

J. Parker (London Research Institute, Cancer Research UK). mCherry-Tks5 plasmid and anti-

Tks5 polyclonal antibody were from Dr. Sara Courtneidge (Burnham Institute for Medical

Research, California, USA). PDBu, DMSO, fibronectin from bovine plasma, anti-vinculin

antibody, mouse IgG1, rabit IgG and other reagents without further mention were purchased

from Sigma (St. Louis, MO).

5.2 Cell culture

Primary normal human bronchial epithelial cells and Bronchial Epithelial Cell Medium

Bullet Kit were from Lonza Walkersville Inc. (Walkersville, MD). The Bronchial Epithelial Cell

Medium BulletKit (BECM BulletKit) contained Bronchial Epithelial Cell Basal Medium (BECB

Medium) and the following supplements, 0.4% BPE, 0.1% Hydrocortisone, 0.1% hEGF, 0.1%

Epinephrine, 0.1% Transferrin, 0.1% Insulin, 0.1% Retinoic Acid, 0.1% Triiodothyronine, and

0.1% GA-1000. These primary cells were freshly isolated from human bronchus (non-cancerous

and non-disease). They were confirmed by cytokeratin staining as the epithelial cell marker. The

primary cells were cultured according to the manufacture’s instruction. These primary cells were

only subject to 1-2 passages before use. Human bronchial epithelial BEAS2B cells, human lung

adenocarcinoma A549 cells from ATCC (Manassas, VA) and v-Src transformed NIH 3T3 cells

from Dr. G. Steven Martin (University of California at Berkeley, Berkeley, CA) were cultured in

Dulbecco’s modified eagle’s medium (DMEM, low glucose) (Life Technologies, Rockville,

MD) with 10% fetal bovine serum (FBS) (GIBCO, Carlsbad, CA), penicillin (1 mg/ml) and

streptomycin (1 mg/ml) (Life Technologies) at 37°C in a 5% CO2 humidified atmosphere

following standard procedures (379, 380).

71

5.3 siRNA transfection

siRNA was transfected into BEAS2B cells by Oligofectamine (Invitrogen, Carlsbad, CA)

following manufacturer’s protocol (381, 382). BEAS2B cells (10, 000 cells/well) were plated

into 6-well plates. siRNA (50 nM) and 10 µl Oligofectamine in OPTI-MEM (Invitrogen) was

added to each well at approximately 50% confluence of cells, and 48 h after incubation, cells

were stimulated with PDBu 500 nM for 30 min and followed by immunofluorescent staining.

5.4 Immunofluorescence staining

Cells were stained according to a standard staining protocol at room temperature (383).

BEAS2B cells (10, 000 cells/well) were cultured on glass coverslips (VWR, Mississauga,

Canada) in 6-well plates. Cells were pre-incubated with designated pharmacological inhibitors

and then challenged with PDBu 500 nM for 30 min. Cells were fixed with 4%

paraformaldehyde/PBS for 30 min at room temperature, permeabilized with 0.1% Triton X-

100/PBS for 10 min on ice and blocked with 1% BSA/PBS for 60 min for non-specific binding.

Cells were incubated with designated primary antibody for 2 h at room temperature and then

designated secondary antibody conjugated with Alexa Fluor 594 (Molecular Probe) for 1 h at

room temperature in dark. Then the cells were incubated with Oregon Green 488 Phalloidin

(Molecular Probe) for 1 h at room temperature and Hoechst dye 33342 for 10 min at room

temperature in dark. Between each step, cells were gently washed with PBS for three times.

After carefully washed with double distilled water, the coverslips were mounted on glass slides

with Dako fluorescence mounting medium (Dako, Mississauga, Canada).

For in situ zymography assay, cells were cultured on coated glass coverslips (VWR,

Mississauga, Canada) in 6-well plates. Cells were pre-treated with designated chemical inhibitors

72

and then stimulated with PDBu 500 nM for 8 h. Cells were fixed with 4%

paraformaldehyde/PBS for 30 min at room temperature and permeabilized with 0.1% Triton X-

100/PBS for 10 min on ice. Then cells were stained with Rhodamine Phalloidin (Molecular

Probe) for 1 h at room temperature and Hoechst dye 33342 for 10 min at room temperature in

dark. Cells were gently washed with PBS for three times between each step. The coverslips were

mounted on glass slides with Dako fluorescence mounting medium (Dako, Mississauga, Canada).

5.5 Microscopy and image analysis

The inverted laser scanning fluorescence confocal microscope (Olympus FluoView

Confocal, FV1000-ASW) equipped with acquisition and analysis software (Version 01.02.01),

and a 1 X 81 oil immersion objective (numerical aperture [NA], 1.40) was used. Cell images

triple stained with Hoechst dye 33342, Oregon Green 488 Phalloidin, and Alexa Fluor 594

labeled secondary antibody were obtained using selective laser excitation at 405 nm (Diode

Laser), 488 nm (Multiline Argon Laser), and 543 nm (Helium Neon Laser), respectively. Z-stack

scanning microscopy images were acquired at step of 0.1 µm from the bottom to the top of the

cell.

Phase-contrast and fluorescence images were also taken with a Nikon TE2000 inverted

fluorescence microscope (40X objective with an optical magnification lens of 10X and oil

immersion objectives of 40X [NA, 1.30] and 60X [NA, 1.40]), while digitally acquired images

were taken with a Nikon DXM 1200F camera (LUCIA 5.0 acquisition software, Nikon).

Fluorescence images were analyzed and processed with Simple PCI v6.0 (C-Image Inc.,

Burlington, MA). Quantification of PDBu-induced cytoskeletal structures was done by counting

at least 200 cells. Cells with dramatic cytoskeleton re-organization and at least 5 of the

73

podosome-like structures, either small dot (> 1 µm), rosette ring, or belt in cytoplasm were

considered to be podosome-positive cell. Cells with long and thin stress fibers were considered

as podosome negative cells.

We quantified the co-localization of the confocal images by Olympus FV10-ASW

Version 01.07.02 imaging software for Pearson’s correlation coefficient according to this

equation:

In this equation, S1 is the signal intensity of pixels in the first channel and S2 is the signal

intensity of pixels in the second channel. S1aver and S2aver indicate the average intensity of pixels

in the first and second channel, respectively. We only quantified the cytoplasm area of each cell

and excluded the peripheral edge of the cell.

5.6 Live cell imaging

BEAS2B cells (100, 000 cells/well) were seeded on round glass coverslips (diameter 50

mm) (Fisher Scientific, Ottawa, Canada) in 60-mm cell culture dish for live cell imaging. Cells

were grown to 20-30% confluence before treated with Lipofectamine 2000 transfection reagent

(Invitrogen), according to the manufacturer’s instructions (382, 384). Plasmids (2 µg) were

added to each 60-mm cell culture dish. After 48 h transfected cells were subjected to live cell

imaging. The time-lapse series of cells were taken at 37°C using the above mentioned confocal

microscope equipped with a computer-driven camera and autofocus system in humidified

chamber with 5% CO2. Differential Interference Contrast (DIC) and fluorescence images were

74

taken every 5 sec for up to 25 min. Original digital images were converted to movies by

Olympus FluoView FV10-ASW Version 01.07.02 software with 330 msec frames.

5.7 Fibronectin based in situ zymography

Fibronectin was conjugated with TRITC succinamidyl ester (Molecular Probes, Eugene,

OR) following manufacturer’s protocol with some modifications (385). Glass coverslips (12

mm) were pre-cleaned with 20% nitric acid for 30 min. Ethanol-sterilized coverslips were coated

with 50 µg/ml poly-L-lysine for 20 min, and fixed with 0.5% glutaraldehyde for 15 min. The

coverslips were inverted on an 80 µl drop of gelatin sucrose matrix for 1 h, and then incubated

with fibronectin matrix (0.2% TRITC-conjugated fibronectin) for 1 h in dark. The residual

reactive groups in the fibronectin-TRITC matrix were quenched with sodium borohydride at 5

mg/ml for 15 min and sterilized with 75% ethanol. The entire procedure was performed at room

temperature. Extensive washing with PBS was performed between steps.

To test the ability of cells to degrade matrix, cells were plated on coated coverslips in 24-

well plates and incubated at 37 ° for 4 h. Foci of degraded matrix were observed as dark holes

on the red fluorescent fibronectin matrix. A cell with at least one hole under the cell or near the

cell edge was considered as a positive cell in degrading matrix. As a positive control for the in

situ zymography technique, NIH3T3 cells were grown to 20-30% confluence, and transfected

with plasmid expressing constitutively active chicken SrcY527F (a kind gift from Dr. B. Elliot,

Queen’s University, Canada) with Lipofectamine PLUS reagent (Invitrogen), according to the

manufacturer’s instructions. Transfected cells were analyzed 24 h later using the same protocol

(385).

75

5.8 Gelatin based in situ zymography

Glass coverslips were pre-cleaned with nitric acid and sterilized with ethanol. After

treated with 50 µg/ml poly-L-lysine and fixed with 0.5% glutaraldehyde, the coverslips were

coated with a base of gelatin sucrose gel at the bottom and then a layer of 0.2% gelatin

conjugated with Oregon Green 488 (Molecular Probes, Eugene, OR) gel on top. The residual

reactive groups in the gelatin conjugated matrix were quenched with 5 mg/ml sodium

borohydride. Cells were plated on coated coverslips in 6-well plates and incubated at 37° for 8

h. Foci of degraded matrix were observed as dark holes (1-2 µm) on the green fluorescent gelatin

matrix. Cells with at least one hole (> 1 µm) under the cell or near the cell edge were counted in

a blinded fashion as positive to degrade matrix.

5.9 Cell invasion assay

Fibronectin gels were prepared in Transwell chambers inserted in 24-well culture plates,

with 100 µl of 2.5 mg/ml fibronectin in PBS with penicillin (1 mg/ml) and streptomycin (1

mg/ml), incubated at 37° for 48 h. Subsequently, 10,000 cells were added to each insert for 4 h

to allow for cell adhesion. PDBu was added to the Transwell insert and the lower well. After

overnight incubation, any cells remaining in the insert were removed using cotton Q-tips. The

nuclei of invaded cells were stained with Hoechst dye 33342 (146). Quantification of invaded

cells was performed for at least 20 fields per coverslip, by counting the number of nucleus

sprouts that had invaded through the fibronectin gels per field (20X objective with magnification

lens of 10X). Cells in the Transwell inserts were fixed with 4% formaldehyde, and stained for F-

actin using Oregon Geen 488 phalloidin. Confocal scanning microscopy was performed to

visualize cellular protrusions inside the pores.

76

5.10 Western blotting

Western blotting was performed according to a standard protocol as previously described

(382, 384). Cell lysates were harvested using protein lysis buffer (50 mM Tris-HCl pH 7.5, 150

mM NaCl, 2 mM EGTA, 2 mM EDTA, 1% Triton X-100, 100 µg/ml soybean trypsin inhibitor,

100 µg/ml benzamidine hydrochloride, 1 mM PMSF, 50 µg/ml aprotinin, 50 µg/ml leupeptin, 50

µg/ml pepstatin A, 50 µg/ml antipain, 50 mM sodium fluoride, 10 mM sodium pyrophosphate,

and 10 mM sodium orthovanadate). Protein (40 µg protein of each sample) was first separated by

SDS-PAGE, and transferred to nitrocellulose membranes by Semi-Dry Transfer Cell (BioRad,

Hercules, CA). After blocking non-specific binding by 5% BSA in TBST buffer (20 mM Tris pH

7.6, 137 mM NaCl, 0.1% Tween 20), the membranes were incubated in primary antibodies for

overnight at 4 °C and HRP-conjugated secondary antibodies for 1 h at room temperature. The

signals were tested with Enhanced Chemiluminescence detection solution (Amersham Pharmacia

Biotech, Piscataway, NJ) or SuperSignal Chemiluminescent substrate (PIERCE Chemical,

Rockford, IL). The membranes were stripped off with Restore Western Blot Stripping Buffer

(PIERCE Chemical, Rockford, IL) for 3h at room temperature and then re-probed with another

primary antibody.

5.11 Gelatinase zymography gel

Gelatin zymography was performed according to standard protocol (147). SDS-PAGE

gels (10%) containing gelatin 0.1% (1 mg/ml) were prepared according to a standard procedure

(385). Cell lysates and culture medium were freshly harvested and prepared with 6X

zymography sample buffer (12.5 mM Tris-HCl pH 6.8, 20% glycerol, 4% SDS (w/v), 0.5%

bromophenol blue). Cell lysates with 30 µg protein were added to each well and gels were run

77

with 1X Tris-Glycine SDS running buffer under standard running conditions (~90V, constant

voltage) for about 10 h (385). After running, the gels were incubated in renaturing buffer (2.5%

triton X-100 in H2O) and then incubated in developing buffer (50 mM Tris, 0.2 M NaCl, 5 mM

CaCl2, 0.02% Brij35) at 37° overnight. Gels were stained with 0.5% (w/v) Coomassie Blue R-

250 staining solution and destained with destaining solution (50% methanol, 10% acetic acid,

40% H2O).

5.12 Real-Time quantitative RT-PCR

The primers used for qRT-PCR for human MMP-2 were 5’-

TTGCTGGAGACAAATTCTGGAG-3’ (hMMP-2F) and 5’-GGGAAGCCAGGATCCATTTT-

3’ (hMMP-2R); for human MMP-9 were 5’-ACGCACGACGTCTTCCAGTAC-3’ (hMMP-9F)

and 5’-AAGCGGTCCTGGCAGAAATA-3’ (hMMP-9R); for human MMP-14 were 5’-

GGCCCAAAGCAGCAGCT-3’ (hMMP-14F) and 5’-AGCCGTAAAACTTCTGCATGG-3’

(hMMP-14R). Total RNA was extracted from BEAS2B cells with TRIZOL Reagent

(Invitrogen), according to the manufacturer’s instructions (386). cDNA was synthesized from

total RNA (1 µg ) using MuLV Reverse Transcriptase (Applied Biosystems, Foster City, CA)

and random hexamers from Taqman Reverse Transcription Reagent kit (Roche Applied Science,

Toronto, Canada) (386). qRT-PCR was performed using 2 × QuantiTect SYBR Green PCR kit

(Roche, Mannheim, Germany) on LightCycler480 (Roche, Mannheim, Germany) according to a

standard protocol (386). The amplification mixtures (10 µL) contained 2.5 µL of the 1:10 diluted

cDNA, 5 µL of SYBR Green I Master mix (Roche, Mannheim, Germany) and 300 nM forward

and reverse primers. Conditions for PCR included 95 °C for 5 min, and 40 cycles of 94 °C for 15

s, 60°C for 60 s. Each assay included a standard curve of five serial dilutions and a no-template

78

negative control. All assays were performed in triplicate. The MMP-2/-9/-14 gene expression

levels were normalized to the level of 18S ribosomal RNA (386).

5.13 Statistics

Each experiment was performed independently for at least three times. The quantification

of each value represents the mean of at least three independent measures + SD. Significance was

determined by ANOVA followed by Tukey's post-hoc analysis or by Student's t-test where

appropriate.

79

6. Chapter Six. Phorbol ester-induced podosomes in normal human bronchial epithelial

cells

The content of this chapter was published in Journal of Cellular Physiology, and the figures were

reprinted with the permission of the Wiley InterScience Wiley-Liss Inc. from:

Xiao H, Eves R, Yeh C, Kan W, Xu F, Mak AS, Liu M. Phorbol ester-induced podosomes in

normal human bronchial epithelial cells. Journal of Cellular Physiology. 2009; 218(2):366-375.

80

6. Chapter Six. Phorbol ester-induced podosomes in normal human bronchial epithelial

cells

6.1 Summary

Spreading and migration of the basal cells neighboring a wound is essential for airway

epithelial repair. To gain insight into the molecular mechanisms that govern these cellular

processes, we asked whether normal human airway epithelial cells can form podosomes, a

cellular structure discovered from cancer and mesenchymal cells that controls migration and

invasion. Herein, we report that phorbol-12, 13-dibutyrate (PDBu), a PKC activator, induced

reorganization of cytoskeletal structure in primary normal human bronchial epithelial cells, and

in normal human airway epithelial BEAS2B cells. Z-stack scanning confocal microscopy

showed that PDBu-induced podosome-like structures contain actin-rich columns that arise from

the ventral surface of the cell, and also revealed the presence of circular ruffles/waves at the

dorsal cell surface. The molecular components of these cytoskeletal structures were determined

with immunofluorescent staining. Using in situ zymography, we demonstrated that PDBu-

induced podosomes were capable of degrading fibronectin-gelatin-sucrose matrix. PDBu also

increased epithelial cell invasion across Transwell chamber. Podosomes and circular dorsal

ruffles may be important for epithelial cell migration and invasion, thus contributing to

respiratory epithelial repair and regeneration.

6.2 Introduction

The respiratory epithelium is frequently exposed to and damaged by infectious and non-

infectious environmental agents. A proper wound repair process is critical to maintain the

81

integrity of the epithelium. Airway and alveolar epithelial cells are essential components of host

defense system (387). Cell survival and repair also play important role in acute lung injury and

other lung diseases related to inflammatory responses (388). Regardless of the source of injury,

lesions in the airway epithelium can lead to cell death and shedding, and resulting in the loss of

surface epithelium (18, 87). After injury, the airway epithelium initiates a wound healing process

to restore its barrier integrity. The remaining viable epithelial cells at the edge of the wound

dedifferentiate, spread, and migrate over the denuded basement membrane to cover the de-

epithelialized zone (16, 389, 390). Cell migration is the most critical and first event occurring

during the epithelial repair process (7, 87).

Recently, many studies have implicated a specialized structure called the podosome as a

potential mechanism for cell migration and invasion (369). Podosomes are actin-rich cellular

structures which protrude into the extracellular matrix (ECM), resulting in localized remodeling

activities (141, 142). Accordingly, matrix degradation at podosomes is thought to contribute to

cellular invasiveness in physiological and pathological situations. Podosomes were first observed

as electron-dense actin-rich structures on the ventral surface of v-Src transformed fibroblasts

(369). Later, similar structures were found in mesenchymal cells, especially cells from

hematopoietic linage, such as monocytes or macrophages (143-145), dendritic cells (370, 371),

natural killer cells (372), and osteoclasts (373, 374). Podosomes have been found in certain

malignant metastatic invasive cancer cells (148). For examples, Spinardi et al. found dynamic

podosome-like structures in rat bladder carcinoma epithelial 804G cells stably transfected with

human 4 integrin subunit. Interestingly, they also showed similar structures in a few human

primary keratinocytes (151). However, whether normal human epithelial cells can form

podosomes either spontaneously or under stimulation is largely unknown.

82

Several stimuli are able to induce podosome formation in cell types that do not

spontaneously have podosomes, such as TGF- in aorta endothelial cells (147), phorbol ester in

vascular smooth muscle cells (146) and endothelial cells (376). Phorbol esters, e.g. phorbol 12-

myristate 13-acetate (PMA) and phorbol-12,13-dibutyrate (PDBu), are often used to activate

protein kinase C (PKC) (375). PKC activation regulates a diverse set of cellular processes

including cell proliferation, differentiation, migration, and invasion (375). PKC may mediate

non-receptor tyrosine kinase c-Src activation and subsequent podosome formation in fibroblasts

(377). Several signaling proteins including c-Src and focal adhesion kinase (FAK) often serve as

downstream PKC effectors (375). Phorbol ester induced dramatic changes in cytoskeletal

structure in murine lung epithelial cells (378).

We hypothesized that during airway repair, normal epithelial cells may form podosomes

in order for the basal cells neighboring the wound to digest ECM molecules for cell spreading

and transmigration. In the present study, we challenged BEAS2B cells (a cell line derived from

normal human bronchial epithelium obtained from autopsy of non-cancerous individuals) with

PDBu to determine podosome formation and function of podosome-like structures. Importantly,

we also found these structures in primary normal human bronchial epithelial cells with similar

functions.

6.3 Results

6.3.1 PDBu induces reorganization of cytoskeleton in normal human bronchial epithelial

cells

83

To determine the effects of PDBu on cytoskeletal structure in normal human bronchial

epithelial cells, BEAS2B cells were treated with PDBu, and the changes in cell morphology was

recorded with live cell imaging. After PDBu treatment, cell morphology changed immediately

with lamellipodia retraction and ruffling (data not shown). The filamentous (F)-actin was stained

and captured with confocal scanning microscopy. Untreated cells displayed long and thin stress

fibers in the cytoplasm (arrow in Figure 6-1A), and cortical filaments along the periphery (Figure

6-1A, arrowhead). PDBu treatment significantly reduced the number of stress fibers and induced

dramatic rearrangement of F-actin structures. The newly formed structures can be characterized

as actin-rich small dots (0.5 µm) (Figure 6-1B), rosette rings (Figure 6-1C-c1), and belts in the

cytoplasm (Figure 6-1C-c2), and membrane ruffles at the edge of peripheral membrane (Figure

6-1C-c3). The number of cells with cytoskeletal structure changes and the types of newly formed

structures are PBDu-dose dependent after 30 min of treatment (Figure 6-1D to 6-1H). We then

challenged cells with 500 nM of PDBu for different time periods. These newly formed

cytoskeletal structures appeared within 10 min (Figure 6-1I). The numbers of small dots (Figure

6-1J) and rings (Figure 6-1K) remained at the constant levels after 10 min, whereas the numbers

of belts (Figure 6-1L) and ruffles (Figure 6-1M) increased gradually. We also treated BEAS2B

cells with 500 nM PDBu for 1, 2, 4, 6, 8, 12, or 24 h. Reorganized cytoskeletal structures were

observed at all these time points (data not shown), which may be due to the continuous exposure

of cells to PBDu.

05

10152025303540

0 500 10001500 2000

A

B

C

E

F Rings # / Cell

H

PDBu (nM)

Ruffles # / Cell

G Belts # / Cell

D% Cells with Cytoskeleton Rearrangement

I

020406080100

0 10 20 30 40 50

% Cells with Cytoskeleton Rearrangement

K

M

L-1012345678

0 10 20 30 40 50

Rings # / Cell

01234567

0 10 20 30 40 50Time (min)

Ruffles # / Cell

-10123456

0 10 20 30 40 50

Belts # / Cell

0

510152025

0 10 20 30 40 50

Small Dots # / CellJ020406080100

0 500 1000 1500 2000Small Dots # / Cell

01234567

0 500 1000 1500 2000

012345

0 500 1000 1500 2000

012345

0 500 10001500 2000

b

c1

c2 c3

Fig 6-1

84

85

Figure 6-1. PDBu induces reorganization of cytoskeletal structure in BEAS2B cells.

Normal human lung bronchial epithelial BEAS2B cells were treated with different doses of PDBu for 30 min, or with 500 nM of PDBu for different time periods. F-actinwas stained with Oregon Green 488 Phalloidin (Green) and nuclei were counter-stained with Hoechst dye 33342 (blue). (A) Long stress fibers and cortical actin fibers were seen in control cells, indicated by an arrow and an arrowhead respectively. PDBu induced reorganization of cytoskeletal structures. The numbers of stress fibers was reduced, and F-actin staining shows newly formed structures: (B) small dots; (C-c1) ring, (C-c2) belt and (C-c3) membrane ruffles. Typical structures were enlarged. Dots, rings and belts are indicated with square boxes and membrane ruffles are shown with circles. Cytoskeletal rearrangement and new structures are quantified after 30 min of treatment with varying concentrations of PDBu, or after different time periods with 500 nM of PDBu. (D) and (I): percentage of cells with dramatic cytoskeletal rearrangement; (E) and (J): small dots; (F) and (K): rings; (G) and (L): belts; and (H) and (M): membrane ruffles.

86

6.3.2 Identification of ventral podosomes and circular dorsal ruffles

The transient and dynamic nature of these small dots, ring and belt structures indicate that

they could be podosomes or podosome-like structures. On the other hand, the ruffles appeared as

so called circular dorsal ruffles (or waves) (141). By definition, podosomes are actin-rich

columns that arise from the ventral surface (147). We performed immunofluorescent staining for

cortactin (an important component of podosomes) and F-actin. In untreated BEAS2B cells,

cortactin was evenly distributed in the cytoplasm (Figure 6-2A). After PDBu stimulation,

cortactin was redistributed to newly-formed small dots in the cytoplasm and ruffles at the edges

of cytoplasm membrane (Figure 6-2A). As shown in a series of digital images of Z-stack

scanning confocal microscopy (Figure 6-2B), the small dots in the cytoplasmic region are focal

inward invaginations of the ventral cell surface into the cytoplasm. Negative controls for

immunofluorescent staining were by using non-specific antibodoies mouse IgG1 and rabbit IgG

or missing primary antibody (data not shown).

Circular dorsal ruffles/waves are transient actin-based structures formed at the dorsal

membrane of cell peripheries (141). To determine the nature of ruffles induced by PDBu in

human airway epithelial BEAS2B cells, we performed scanning confocal microscopy along the

X-Z axis. The co-localization of cortactin (red) and F-actin (green) at the edge of the peripheral

membrane appeared to be swept back along the dorsal side of the cell (arrows in Figure 6-2C).

This distinct feature differentiates circular dorsal ruffle (waves) from linear ruffles (141).

B

F-actin Cortactin OverlayNucleusC

ontr

olPD

Bu

AF-

actin

Cor

tact

inN

ucle

us

0.0 µm 0.3 µm 0.6 µm 0.9 µm

1.2 µm 1.5 µm 1.8 µm 2.1 µm

Bottom

2.4 µm 2.8 µm 3.1 µm 3.4 µm

Top

Fig 6-2

87

C F-actin Cortactin NucleusFig 6-2

88

Figure 6-2. Characterization of ventral podosomes and circular dorsal ruffles in BEAS2B cells.

BEAS2B cells were treated with or without PDBu 500 nM for 30 min. F-actin stained with Oregon Green 488 Phalloidin (Green), cortactin was stained with Alexa Fluor 594 labeled secondary antibody (Red), and nucleus was stained with Hoechst dye 33342 (Blue). (A) PDBu-induced formation of podosomes (small boxes) and membrane ruffles (circles). (B) A series of digital images from Z-stack scanning confocal microscopy demonstrates that small dots are focal inward invaginations at the ventral surface into the cytoplasm. (C) Z-stack scanning confocal microscopy demonstrates that PDBu induced ruffles/waves are focal outward protrusions of the dorsal surface membrane (arrows).

89

6.3.3 Formation of podosomes in primary normal human bronchial epithelial cells

To further determine the physiological relevance of podosome formation, primary normal

human bronchial epithelial cells were stimulated with 500 nM PDBu. These cells showed long

and thin stress fibers in the cytoplasm (Figure 6-3A). After PDBu treatment, stress fibers

disappeared, and numerous podosomes featured as small dots were found in the cytoplasm, and

circular waves/membrane ruffles were found at the periphery of the cells (Figure 6-3A). The

podosome structures in primary normal human bronchial epithelial cells were confirmed with Z-

stack scanning confocal microscopy. Redistribution of cortactin from the cytoplasm of control

cells to the small dots was noted after PDBu stimulation (Figure 6-3B). These dots were

clustered at the ventral surface of the cell and co-localized with cortactin as focal inward

protrusions (Figure 6-3C square box). The membrane ruffles were identified as circular dorsal

ruffles/waves by Z-stack scanning technique as well (Figure 6-3C cricle). We also examined

effects of PBDu on A549 cells, a cell line derived from human lung cancer. PDBu also induced

podosomes in these cells. However, the doses of PDBu required were much lower (Figure 6-4).

No Treatment PDBu

F-actin Cortactin Overlay

Con

trol

PDB

u

0.0 µm 0.3 µm 0.6 µm 0.9 µm

1.2 µm 1.5 µm 1.8 µm 2.1 µm

Bottom

2.4 µm 2.8 µm 3.1 µm 3.4 µm

Top

B

C

A

F-ac

tinC

orta

ctin

Nuc

leus

Fig 6-3

90

91

Figure 6-3. Characterization of ventral podosomes and circular dorsal ruffles in primary normal human bronchial epithelial cells.

(A) PDBu induced podosome formation and circular dorsal ruffles. Normal human bronchial epithelial cells were treated with or without PDBu 500 nM for 30 min, and F-actin was stained with Rhodamine Phalloidin (Red). Podosomes are indicated within a box and membrane ruffles with a circle. (B) PDBu-induced podosomes (boxes) and membrane ruffles (circles) are co-localized with cortactin. F-actin was stained with Oregon Green 488 Phalloidin (Green), cortactin was stained with Alexa Fluor 594 labeled secondary antibody (Red), and nucleus was stained with Hoechst dye 33342 (Blue). (C) A series of Z-stack scanning microscopy images shows that PDBuinduced podosomes are localized at the ventral surface.

0 nM 10 nM 50 nM 100 nM 500 nM

PDBu 30 min

% Cells with Podosomes

020

4060

80100

0 100 200 300 400 500 600PDBu (nM)

A

B

Fig 6-4

92

Figure 6-4. PDBu induces reorganization of cytoskeletal structure in A549 cells.

(A) A549 cells were treated with different doses of PDBu for 30 min. F-actin was stained with Rhodamine Phalloidin (Red) and nuclei were counter stained with Hoechst dye 33342 (blue). Representative confocal microscopic images are used to show newly formed cytoskeletal structures. Typical structures were enlarged in the right column. Podosomes are indicated with box and membrane ruffles with circle. (B) Quantitative analysis of podosome formation after cells was treated with varying concentrations of PDBu for 30 min.

93

6.3.4 Accumulation of actin-associate proteins and regulators in podosome-like structures

We then used BEAS2B cells to further identify protein components of PDBu-induced

podosomes and circular dorsal ruffles. Immunofluorescent staining was performed using

antibodies targeting a number of proteins known to be localized in these structures. Four groups

of proteins were studied: (i) actin and actin-associated proteins (e.g., F-actin, cortactin, -

actinin), and regulators (e.g., Arp2/3, WASP, N-WASP); (ii) integrin and integrin-associated

proteins (e.g., 1-integrin, vinculin, paxilin, talin); (iii) protein kinases (e.g., Src and FAK),

which regulate the actin cytoskeleton and membrane remodeling; and (iv) matrix

metaloproteinases (MMPs), which mediate ECM degradation.

In addition to cortactin (Figure 6-2 and Figure 6-3), we also studied -actinin, which was

localized mainly in the cytoplasm and under surface membrane in untreated cells (Figure 6-5A).

Upon PDBu stimulation, it was translocated to membrane ruffles, dots, and rings (Figure 6-5A).

Actin regulators such as WASP, N-WASP and Arp2/3 (391) govern actin polymerization

and actin network attachment to the moving membranes at the leading edge (392). In BEAS2B

cells, Arp3 and WASP are mainly in the cytoplasm and weakly stained along actin filaments or

with cortical filaments (Figure 6-5B and 6-5C), whereas N-WASP was found only in the

cytoplasm and cell periphery (Figure 6-5D). Upon PDBu stimulation, Arp3 was translocated to

both small dots and ruffles (Figure 6-5B); whereas WASP and N-WASP were mainly

translocated to membrane ruffles with less distribution in small dots (Figure 6-5C and 6-5D).

F-actin Arp3

Con

trol

PDB

u

Overlay

F-actin -actinin

Con

trol

PDB

u

OverlayA

B

Fig 6-5

F-actin WASP

Con

trol

PDB

u

OverlayC

94

F-actin N-WASPC

ontr

olPD

Bu

OverlayD

Fig 6-5

95

Figure 6-5. Localization of actin associated proteins in podosomes and circular dorsal ruffles.

BEAS2B cells were treated with or without PDBu 500 nM for 30 min. F-actin was stained with Oregon Green 488 Phalloidin (Green) and nucleus was stained with Hoechst dye 33342 (Blue). -actinin, Arp3, WASP and N-WASP were stained with specific antibodies and revealed with Alexa Fluor 594 labeled secondary antibody (Red). Localization of podosomes was shown with boxes, and membrane ruffles with circles. (A): -actinin; (B): Arp3; (C): WASP and (D): N-WASP.

96

6.3.5 Recruitment of integrin and associated proteins in podosome-like structures

The integrin 1 subunit was distributed along the plasma membrane and also in the

cytoplasm weakly along the stress fibers in untreated cells. Upon PDBu treatment, 1 integrin

was translocated to the membrane ruffles and small dots (Figure 6-6A). In control cells, vinculin

(Figure 6-6B) and paxillin (Figure 6-6C) were mainly in the cytoplasm and were enriched in

prominent focal adhesions, at the ends of stress fibers. Upon PDBu-stimulation the punctate

structure of focal adhesions disappeared, and vinculin was distributed mainly in the cytoplasm

but not in podosome-like structures (Figure 6-6B). Paxillin was also mainly in the cytoplasmic

region, however, increased staining was also seen at the newly formed ruffles (Figure 6-6C). In

control cells, talin was found at the cortical filaments and also at focal adhesion sites; PDBu

treatment led to the translocation of talin to the membrane ruffles (Figure 6-6D). The different

distribution and translocation of these three integrin-associated proteins suggest that they may

play different role in cytoskeletal organization in human airway epithelial cells.

AF-actin Integrin 1 Overlay

Con

trol

PDB

u

F-actin Vinculin

Con

trol

PDB

u

B Overlay

Fig 6-6

F-actin Paxilin

Con

trol

PDB

u

OverlayC

97

F-actin TalinC

ontr

olPD

Bu

D Overlay

Fig 6-6

98

Figure 6-6. PDBu-induced redistribution of integrin and integrin associated proteins.

BEAS2B cells were treated with or without PDBu 500 nM for 30 min. F-actin was stained with Oregon Green 488 Phalloidin (Green) and nucleus was stained with Hoechst dye 33342 (Blue). Integrin 1 subunit, vinculin, paxilin and talin were stained with specific antibodies and revealed with Alexa Fluor 594 labeled secondary antibody (Red). Localization of podosomes was shown with boxes, and membrane ruffles with circles. (A): Integrin 1 subunit; (B): vinculin; (C): paxilin and (D): talin.

99

6.3.6 Lack of vinculin in PDBu-induced small dots in normal human lung epithelial cells

Vinculin is known to be an important component of podosome-like structures in many

cell types (272, 393). To determine the relationship between vinculin and actin-rich small dots,

BEAS2B cells, primary normal human bronchial epithelial cells (NHBE), and human lung

cancer A549 cells were treated with PDBu (500 nM for 30 min), followed by immunofluorescent

staining. We also stained v-Src transformed NIH3T3 cells for comparison because vinculin has

been found in so-called “rosettes” or ring-like structures in these cells (393). The F-actin staining

was switched from Oregon Green 488 Phalloidin (green) to Rhodamine-labeled Phalloidin (red).

Vinculin was revealed with FITC-labeled secondary antibody. Our results show that vinculin

was clearly found in ring-like structures in v-Src transformed NIH3T3 cells (Figure 6-7A),

indicating that the antibody and staining conditions we used were comparable to that of others. In

PDBu treated A549 cells, vinculin staining was found in many actin-rich dots (Figure. 6-7B).

The vinculin staining was also found in ring-like structures, surrounded by actin-rich rings

(Figure 6-7B). In primary NHBE cells, vinculin remained at the focal adhesion like sites, and

was not localized to the actin-rich dots (Figure 6-7C). In BEAS2B cells, again, vinculin was

found in belts and ruffles after PDBu stimulation, but not in small dots (Figure 6-7D).

A

B

No

Trea

tmen

tPD

Bu

A54

9 C

ell

Vinculin F-actin Overlay

No

Trea

tmen

t

v-Sr

cTr

ansf

orm

ed

NIH

3T3

Cel

l

Vinculin F-actin Overlay

Fig 6-7

No

Trea

tmen

tPD

BuPr

imar

y N

HB

E C

ell

C Vinculin F-actin Overlay

100

Vinculin F-actin OverlayN

o Tr

eatm

ent

PDB

uBEA

S2B

Cel

lD

Fig 6-7

101

Figure 6-7. Lack of translocation of vinculin to actin-rich dots in normal human airway epithelial cells after PDBu stimulation.

BEAS2B cells, primary NHBE cells, and human lung cancer A549 cells were treated with or without PDBu 500 nM for 30 min. Untreated v-Src transformed NIH3T3 cells were used as staining control. F-actin was stained with Rhodamine Phalloidin (Red) and nucleus was stained with Hoechst dye 33342 (Blue). Vinculin was revealed with FITC labeled secondary antibody (Green). Localization of podosomes was shown with boxes, and membrane ruffles with circles. (A): v-Src Transformed NIH3T3 cells; (B): A549 cells; (C): primary NHBE cells, and (D): BEAS2B cells.

102

6.3.7 Increased protein tyrosine kinases in podosome-like structures

Increased protein tyrosine phosphorylation is commonly observed in podosomes and

circular dorsal ruffles. In control BEAS2B cells, phosphotyrosine staining was localized

predominately along the cell periphery and in the cytoplasm at structures that appear to be the

focal adhesion sites (Figure 6-8A). PDBu stimulation enhanced phosphotyrosine staining at the

ruffles and small dots (Figure 6-8A). Phosphorylation of tyrosine-416 of c-Src is known to be a

sign of its activation (382, 384, 394, 395). PDBu treatment resulted in activation (increased

staining) and translocation of c-Src into actin-rich structures at the cell periphery and in small

dots (Figure 6-8B). In untreated cells, FAK was in cytoplasm and weakly along stress fibers

(Figure 6-8C). Upon PDBu treatment, FAK was translocated to membrane ruffles and to a few

small dots as well (Figure 6-8C).

6.3.8 Distribution of MMPs in podosome-like structures

We further examined the distribution patterns of MMPs. In untreated cells, MT1-MMP

was cytosolic, localized predominantly around the nucleus (Figure 6-9A); MMP-2 and MMP-9

were found both in the cytoplasm and along the cell periphery (Figure 6-9B and 6-9C). After

PDBu treatment, a portion of MT1-MMP remained in the peri-nuclear region, while enhanced

staining was found along the membrane ruffles and in some small dots (Figure 6-9A). MMP-2

and MMP-9 staining was increased around the nucleus after PDBu stimulation (Figure 6-9B and

6-9C). MMP-2 was found at the membrane ruffles and small dots (Figure 6-9B) whereas MMP-9

was only found along membrane ruffles (Figure 6-9C).

F-actin pTyrC

ontr

olPD

Bu

A

B

Overlay

F-actin pSrcY416 Overlay

Con

trol

PDB

u

Fig 6-8

103

C

Fig 6-8

F-actin FAK Overlay

Con

trol

PDB

u

104

Figure 6-8. Protein tyrosine kinases were recruited to PDBu-induced podosomes and circular dorsal ruffles.

BEAS2B cells were treated with or without PDBu 500 nM for 30 min. F-actin was stained with Oregon Green 488 Phalloidin (Green) and nucleus was stained with Hoechst dye 33342 (Blue). Protein tyrosine phosphorylation, phospho-SrcY416 and focal adhesion kinase (FAK) were stained with specific antibodies and revealed with Alexa Fluor 594 labeled secondary antibody (Red). Localization of podosomes was shown with boxes, and membrane ruffles with circles. (A): Tyrosine phosphorylatedproteins; (B): phospho-SrcY416; (C): FAK.

F-actin MT1-MMP OverlayC

ontr

olPD

Bu

F-actin MMP-2 Overlay

Con

trol

PDB

uA

B

Fig 6-9

105

F-actin MMP-9 Overlay

Con

trol

PDB

uC

Fig 6-9

106

Figure 6-9. Matrix metalloproteases are associated with podosomes and circular dorsal ruffles.

BEAS2B cells were treated with or without PDBu 500 nM for 30 min. F-actin was stained with Oregon Green 488 Phalloidin (Green) and nucleus was stained with Hoechst dye 33342 (Blue). MT1-MMP, MMP-2 and MMP-9 were stained with specific antibodies and revealed with Alexa Fluor 594 labeled secondary antibody (Red). (A): MT1-MMP; (B): MMP-2; and (C): MMP-9. Localization of podosomes was shown with boxes, and membrane ruffles with circles. MMP-9 was co-localized with belt-like actinstructures (indicated with arrows).

107

6.3.9 PDBu-induced podosome-like structures can degrade extracellular matrix

MMP-9 is crucial for human airway epithelial migration (87). MT1-MMP is necessary

for distal airway epithelial repair after injury (396). To determine if proteolytic activity is

associated with the PDBu-induced podosomes in epithelial cells, in situ zymography were

performed. BEAS2B cells were plated on a coverslip pre-coated with a layer of TRITC-

conjugated (a red fluorescent dye) fibronectin matrix and a base of gelatin and sucrose gel. ECM

substrate degradation was visualized as black areas devoid of red fluorescence. As shown in

Figure 6-10A, almost no matrix degradation was found under the control condition. In PDBu-

treated cells, the dark holes formed by degradation of the fibronectin-TRITC matrix were

generally found in the cytoplasm area with the sizes similar to small dots. We also transfected

NIH 3T3 cells with constitutively activated SrcY527F as a technical control to ensure the assay

works well, which showed stronger degradation of ECM as small dots (Figure 6-10A lower

panel). We also performed in situ zymography with primary normal human bronchial epithelial

cells using a similar method; digestion of the matrix was also observed (data not shown). Since

the in situ zymography was performed 4 h after PDBu stimulation, these dark holes represent the

consequence of ECM degradation. However, due to cell migration, this degradation did not

overlap well with actin-rich dots.

A FN-TRITCF-Actin OverlayC

ontr

olPD

Bu

SrcY

527F

No Treatment PDBuB C

0

12

34

56

7

0 nM 500 nM

PDBu (nM)

Fold

of C

ell I

nvas

ion *

Fig 6-10

108

109

Figure 6-10. BEAS2B cells with podosomes can degrade matrix and are invasive.

(A) Digestion of fibronectin-gelatin-sucrose matrix by PDBu-induced podosomes. BEAS2B cells were seeded on coverslips coated with TRITC conjugated fibronectin-gelatin-sucrose matrix. After treatment with 500 nM PDBu for 4 h, degradation of ECM was shown as dark dots and F-actin was stained to show cytoskeletalstructures. NIH 3T3 cells co-transfected with -actin-GFP and constitutively activated chicken SrcY527F constructs were included as a technical control. These experiments performed by Rob Eves (Queen’s University) and me. (B) BEAS2B cells with podosomes are invasive. BEAS2B cells were seeded into a Transwell insert with 8 µm pores coated with fibronectin gel. After 500 nM PDBu treatment, cells remaining in the Transwell insert were fixed and stained with Oregon Green 488 Phalloidin(Green) for F-actin. The cells crossing the insert were visualized by nucleus Hoechst dye 33342 staining (blue). (C) Results were quantified for the invasiveness of BEAS2B cells. *: P<0.05 compared with control group.

110

To explore the migration properties of BEAS2B cells bearing podosomes, we performed

a Transwell fibronectin gel invasion assay. After overnight incubation, Z-stack scanning

confocal microscopy was applied to visualize cells inside the fibronectin gel from the top of the

Transwell. BEAS2B cells with podosomes degraded fibronectin gel, were able to invade through

the gel to the bottom side of the well (Figure 6-10B). The number of cells crossed the Transwell

was quantified by counting the stained nuclei (Figure 6-10C). The results indicated that PDBu

treatment increased the invasiveness of normal human bronchial epithelial cells.

6.4 Discussion

The respiratory epithelium is known to undergo continuous damage and repair. Rapid re-

epithelialization by epithelial cell de-differentiation and migration are important features of

epithelial repair in vivo (75). Repair generally involves several steps, including migration and

spreading of epithelial cells at the margin of the injury into the damaged region, invading of

epithelial cells from the underneath layers to the surface of the epithelium, and proliferation of

epithelial cells (7). In the present study, we identified podosome and circular ruffle structures in

primary normal human bronchial epithelial cells and in BEAS2B cells stimulated with the

phorbol ester, PDBu. In most of our experiments, 500 nM of PDBu was used, which is lower

than doses typically used in podosome studies with other cell types. As shown in Figure 6-1,

podosome-like structures were found in lower doses of PDBu treated cells. These structures were

revealed by Z-stack scanning confocal microscopy with multiple molecular markers.

Importantly, we demonstrated digestion of ECM by podosomes and the invasive properties of

cells treated with PDBu.

111

Although podosomes in different cell types showed similar functions in promoting matrix

degradation and invasion, their morphologies, numbers and sizes appear to be diverse. For

example, podosomes in primary human macrophages look like hundreds of small tiny actin-rich

dots (0.2-0.5 µm) distributed at the ventral surface underneath the cytoplasm (143-145). In

smooth muscle cells, podosomes appear as small actin-rich dots (0.5-1 µm) which often

aggregate to form bands (2 µm) along the cortex of the cell (146). In bovine aorta endothelial

cells, podosomes form a single ring (15 µm) beside the nucleus (147). In v-Src transformed NIH

3T3 fibroblasts, podosomes are numerous rosette-like structures (2 µm) (148). In melanoma

cells, podosomes are fewer in number, and can be seen as small dots (approximately 1 µm) in the

cytoplasm (149). In rat bladder carcinoma 804G cells, podosome-like structures were identified

around hemidesmosomes that make connections with intermediate filaments, as small dots

surrounded by rings (150, 151). The features of PDBu-induced podosomes in human airway

epithelial cells are seen as numerous small dots in the cytoplasm, with occasional rosette rings

and belts. Linder described podosomes as dot-like structures attached to substrate, containing

actin regulators and plaque proteins, with the number of around 20-100 per cell, with the

maximum size of 1 µm X 0.4 µm (272). The sizes of the small dots in BEAS2B cells and normal

human bronchial epithelial cells are within this range.

The molecular component of rosettes of podosomes contain several groups of proteins

identified in aorta endothelial cells (147). In the present study, we identified some of these

proteins in podosome-like structures and some in circular dorsal ruffles. Spinardi et al. showed

the presence of vinculin in the small rings around the small dots in rat bladder carcinoma 804G

cells that were stably transfected with human 4 integrin subunit (150, 151). However, in the

present study, we did not find translocation of vinculin to small dots in BEAS2B cells, nor in

112

primary NHBE cells. In an early study, Dwyer-Nield et al. (378) reported PMA-induced

cytoskeletal reorganization in mouse lung epithelial E10 cells. They found vinculin dissociated

from the focal adhesion plaques and distributed throughout the cytoplasm (378). Our

observations were similar to what reported in that study. Interestingly, we found vinculin in

actin-rich dots in human lung cancer A549 cells, as well as in rosettes in v-Src transformed

NIH3T3 cells, which are tumorigenic. Thus, our results implicated that the composition and

function of podosome-like structures formed by normal epithelial cells may be different from

that of cancer and tumor cells. We could use vinvulin antiby to do immunoprecipitation in these

four cell types and identify the special proteins and kinases involved in vinculin recruitment to

podosomes. An alternative view is that these cells may form invadopodia rather than podosomes.

There is much discussion as to whether these two structures are related, which is beyond the

scope of this study. It has been suggested that the short-lived podosomes can probe the substrate

for sites of attachment before maturing into invadopodia (159, 274, 279). Gatesman A et al.

demonstrated that actin filament-associated protein (AFAP) is required to mediate PKC

activation of the nonreceptor tyrosine kinase c-Src and the subsequent formation of podosomes

in fibroblast cells upon PMA stimulation (377). Recently, Dorfleutner A et al. reported that the

phosphorylation and/or dephosphorylation of AFAP is involved in the regulation of podosome

stability and lifespan in A7r5 smooth muscle cells (171). In a separate study, we over-expressed

GFP tagged AFAP in BEAS2B cells and performed live cell imaging after PDBu treatment.

Small dots appeared within 2 min, with fast turnover, from 10 sec to 5 min (data not shown).

Therefore, we believe that the small dots in the cytoplasm of airway epithelial cells are likely to

be podosomes than invadopodia.

113

We observed translocation of the three MMPs to podosome-like structures in BEAS2B

cells after PDBu stimulation.. We found MT1-MMP and MMP-2 in actin-rich dots and MMP-9

in belt-like structures. All three MMPs were found in membrane ruffles upon PDBu stimulation

in BEAS2B cells. It has been proposed that the protease profiles in lamellipodia and in

invadopodia (podosomes) are similar, but the proteolytic activities of these structures are

different, determined by the balancing between proteases and their inhibitors (397). Tissue

inhibitor of matrix metalloproteinase-2, for example, has been found to regulate the ECM-

degrading activities of MT1-MMP and MMP-2 (397). The distribution and interaction between

TIMPs and MMPs should be further studied.

Podosomes and membrane ruffles in human airway epithelial cells comprise MMPs to

endow proteolytic properties. Interestingly, although PDBu induced formation of both

podosomes and circular dorsal ruffles, the digestion of matrix was only seen as small dots. We

further demonstrated the invasiveness of PDBu-treated cells using the Transwell system. PDBu-

induced formation of podosomes may play a critical role in normal epithelial cell invasion and

migration during airway repair. PKC activation could be an underlying mechanism of these

cellular processes.

Circular dorsal ruffles/waves, also called actin ribbons, are another transient cytoskeletal

structure that form in response to external stimuli. PMA-induced morphological changes of

keratinocytes have been reported as far as two decades ago (398), which have since been

described as circular dorsal ruffles (141). These ruffles were located at the edges of cells as

distortions of plasma membrane, very similar to what we reported in the present study. We used

Z-stack scanning confocal microscopy to demonstrate that these structures are indeed swept back

along the dorsal side of the cell. Moreover, in our time course study, the ruffles appeared

114

gradually after PDBu stimuli, which is in contrast to the rapid formation of peripheral ruffles

seen in lamellipodia (141). The molecular components of circular dorsal ruffles are similar to

podosomes in BEAS2B cells. The function of these circular dorsal ruffles is yet unknown. It has

been proposed that they might promote degradation of ECM during three-dimensional migration

(141).

Alteration of cytoskeletal structures can be induced by growth factors, cytokines and

many other biological mediators. For example, we have found dramatic depolymerization of F-

actin in rat lung epithelial cells after LPS stimulation (399-402), which are essential for TNF-

(399) and macrophage inflammatory protein-2 (402) expression in these cells. Whether these

stimuli can induce formation of podosomes and/or circular dorsal ruffles merits further

investigation.

In summary, PDBu as a PKC activator induced formation of podosomes and circular

dorsal ruffles in primary normal human bronchial epithelial cells and in BEAS2B cells. The

molecular components of these structures are similar but only podosomes showed evidence of

being able to digest matrix. These transient cytoskeletal structures may play important roles in

mediating airway epithelial cell migration and invasion, which could be important for tissue

repair, airway branching and lung development. Dysfunction of this system may contribute to

acute lung injury and chronic obstructive pulmonary disease, as well as other disorders of the

airway and the lung.

115

7. Chapter Seven. PKC regulates recruitment of MMP-9 to podosomes, and its release and

activation

The content of this chapter was prepared as an original research article in Molecular Biology of

the Cell:

Xiao H, Bai XH, Kapus A, Lu WY, Mak AS, Liu M. PKC regulates recruitment of MMP-9 to

podosomes, and its release and activation. Molecular Biology of the Cell. 2009. (Under revision)

116

7. Chapter Seven. PKC regulates recruitment of MMP-9 to podosomes, and its release and

activation

7.1 Summary

Podosomes are transient cell surface structures essential for degradation of extracellular

matrix during cell migration and invasion. Protein kinase C (PKC) is involved in the regulation

of podosome formation; however the roles of PKC isoforms in podosome formation and function

are largely unknown. In Chapter 6, we reported that phorbol-12, 13-dibutyrate (PDBu), a PKC

activator, induced the formation of podosome-like structures in normal human bronchial

epithelial cells. Here, we demonstrate that PDBu-induced podosome formation is mainly

mediated through redistribution of conventional PKCs, especially PKC , from the cytosol to the

podosomes. Interestingly, although blocking atypical PKC (PKC ) did not affect PDBu-induced

podosome formation, it significantly reduced gelatin matrix degradation at podosomes. Inhibition

of PKC reduced PDBu-induced gelatin matrix degradation at podosomes by reducing

recruitment of MMP-9 to podosomes, and its release and activation. Down-regulation of MMP-9

by siRNA and neutralization antibody also significantly reduced gelatin matrix degradation.

PDBu-induced recruitment of PKC and MMP-9 to podosomes was blocked by inhibition of

novel PKC with Rottlerin or PKC siRNA. Our data suggest that while the conventional PKC

controls podosome formation and dynamics, however, it is the atypical PKC that plays a

dominant role in the recruitment of MMP-9 to podosomes for gelatin matrix degradation.

117

7.2 Introduction

Podosomes, first described by Marchisio and colleagues (403), are short protrusions of

the ventral cell surface in direct contact with the substrate matrix. Recently, it has been

discovered that these cellular structures are responsible for cell invasion by degrading the

extracellular matrix (ECM) barriers (272, 369). These structures are composed of an F-actin

core, surrounded by a ring structure that is consisted of actin associated proteins, signal

transduction proteins, and matrix metalloproteases (MMPs), which include membrane-bounded

MMPs (such as MMP-14) and secreted MMPs (such as MMP-2 and MMP-9) (272). Several

exogenous signals can trigger the formation of podosomes, including phorbol ester, bombesin,

bradykinin, cytokines, growth factors, and growth hormones (404). These agonists act by

activating tyrosine and serine/threonine kinases, among which protein kinase C (PKC) appears to

be a key mediator of podosome formation (376, 405). PKC may lead to the activation of non-

receptor tyrosine kinase, c-Src, which in turn facilitates podosome formation (377).

PKC is a family consisting of 11 isozymes, which are classified into three subfamilies,

conventional/classical, novel, and atypical PKCs, based on their second messenger requirements

(375). The conventional PKCs (cPKCs) are , I, II, and , which require Ca2+, diacylglycerol

(DAG), and phospholipid for activation (375). The novel PKCs (nPKCs) include , , , and ,

which require DAG but not Ca2+ for activation (406). On the other hand, atypical PKCs (aPKCs),

including and / , require neither Ca2+ nor DAG for activation (375, 407). Due to their

structural similarity to DAG , phorbol esters, e.g. phorbol 12-myristate 13-acetate (PMA),

phorbol-12,13-dibutyrate (PDBu), are often employed to elicit PKC related signals (375, 407).

PKCµ, also called protein kinase D1, is another serine/threonine kinase, which has two C1

domains and a pleckstrin homology domain but no Ca2+ binding domain (292). It shares similar

118

functions with PKCs. Each PKC isozyme may have distinct roles in various cellular processes

(408).

PKC activation has been shown to remodel actin stress fibers into F-actin-enriched

podosome columns in cultured cells (213). Short-term exposure to PMA or PDBu induced

appearance of podosomes and rosettes in endothelial cells (376), vascular smooth muscle cells

(405, 409), and osteoclasts (410). Actin filament-associated protein AFAP-110 was required to

mediate PKC related activation of c-Src and subsequent formation of podosomes (377). PKCs

have been implicated in podosome formation in human umbilical vein endothelial cells (376),

and vascular smooth muscle cells (411). However, the contribution of different PKC isozymes in

these cellular processes is largely unknown.

In Chapter 6, we reported that PDBu-induced formation of podosome-like structures in

normal human bronchial epithelial cells. These structures are enriched in MMPs and endowed

with proteolytic activity to degrade fibronectin-gelatin-sucrose matrix (122). In the present study,

we asked whether distinct PKC isoenzymes are involved in podosome formation, and

recruitment of MMPs for ECM digestion in PDBu-stimulated human bronchial epithelial cells.

We found that cPKCs, especially PKC , is crucial for the formation of the podosomes, PKC

plays an important role to control the recruitment, release, and activation of MMP-9 at

podosomes, and PKC controls the recruitment of PKC to podosomes.

7.3 Results

7.3.1 PDBu-induced PKC phosphorylation and translocation to podosomes and circular

ruffles

119

Recently we have reported that phorbol ester (PDBu) could dramatically change cell

morphology and cytoskeletal structures in a dose- and time-dependent manner in primary normal

human bronchial epithelial cells and in human bronchial epithelial BEAS2B cells. Ventral

podosome-like structures and dorsal circular dorsal ruffles were identified with Z-stack scanning

confocal microscopy, together with immunostaining of a number of known marker proteins

(122). To determine which PKC isozymes are activated upon PDBu stimulation, we examined

their phosphorylation status after challenging BEAS2B cells with 500 nM PDBu from 0 to 240

min. The phosphorylation status of cPKCs was determined by western blotting with two

antibodies, one targets the phosphorylation sites of PKC at T638 and PKC II at T641 (Figure

7-1Aa1), and another pan-PKC antibody that targets amino acid sequence similar to the PKC II

S660 region (Figure 7-1Aa2). PDBu treatment did not induce significant alteration of

phosphorylation levels of cPKCs. On the other hand, after PDBu treatment, phosphorylation of

PKC T505 significantly increased after 1 min, peaked at 20-30 min and declined gradually

(Figure 7-1Aa4). The phosphorylation of PKC S643/676 increased gradually within the first 10-

20 min followed by a sharp increase at 30 min and plateaued for an hour, and then decreased

(Figure 7-1Aa5). The phosphorylation of PKC T538 also increased upon PDBu stimulation

within the first 10 min (Figure 7-1Aa6). PDBu also induced phosphorylation of aPKC within 1

min as detected by an antibody for PKC / T410/403 (Figure 7-1Aa8). The phosphorylation of

PKCµ S744/748 (Figure 7-1Aa10), and PKCµ S916 (Figure 7-1Aa11) dramatically increased

within 5 min of PDBu treatment. Total protein levels of PKC , PKC , PKC , PKCµ and

GAPDH remained unchanged during the eprimental period (Figure 7-1). These results suggest a

rapid activation of nPKC, aPKC, and PKC induced by PDBu in BEAS2B cells.

Fig 7-1

0 1 5 10 20 30 60 120 240 min

PDBu 500 nM

(p)PKC T505

(p)PKCµ S744/748

(p)PKCµ S916

(p)PKC T538

(p)PKC / T410/403

GAPDH

A

a1

a2

a4

a5

a6

a8

a10

a11

a13

(p)PKC pan II S660

PKC

PKC

PKC

PKCµ

a3

a7

a9

a12

(p)PKC / II T638/641

(p)PKC / S643/676

120

Figure 7-1. PDBu-induced PKC phosphorylation.

(A) PDBu-induced PKC phosphorylation. BEAS2B cells were treated with PDBu 500 nM for 0 min to 240 min. The total or phosphorylation of PKC isoforms was revealed by designated antibodies.

121

We then examined PDBu-induced distribution of PKC isoforms and phosphorylated

species. Under control conditions, PKC , , and , representative of cPKC, nPKC, and aPKC

subfamilies, respectively, as well as PKC were localized in the cytoplasm and perinuclear

region, but none co-localized with prominent actin stress fibers. PDBu treatment induced

disassembly of actin stress fibers and translocation of PKC isoforms and their phosphorylated

forms to small dots, rings, and circular ruffles (Figure 7-2A and 7-2B). The colocalization

coefficient of F-actin staining with PKC isoforms or their phosphorylated forms was measured

with multiple cells and expressed as Pearson’s correlation coefficient values in Figure 7-2C and

7-2D. We also did immunofluorescent staining for PKC , phosphorylated PKC / II T638/641,

and phosphorylated PKC pan II S660 in primary normal human bronchial epithelial cells with

similar results (data not shown). Therefore, although the phosphorylation status of cPKC

isoforms did not change drastically, they were clustered at the sites of podosomes and circular

ruffles in human bronchial epithelial cells upon PDBu stimulation.

Fig 7-2

PKC PKC PKC PKCµC

ontro

lP

DB

uA

(p)PKCa/ II T638/641 (p)PKC T505 (p)PKC / T410/403 (p)PKCµ S744/748

Con

trol

PD

Bu

B

122

0

0.2

0.4

0.6

0.8

1

PKC PKC PKC PKCµ

Pear

son'

s C

orre

latio

n C

oeffi

cien

t

(p)PKC (p)PKC (p)PKC (p)PKCµ

No TreatPDBu

0

0.2

0.4

0.6

0.8

1

Pear

son'

s C

orre

latio

n C

oeffi

cien

t

C D

Fig 7-2

PKCs (p)PKCs

123

********

Figure 7-2. PDBu-induced PKC translocation to podosomes.

PDBu induced translocation of PKC isoforms (A) and their phosphorylated forms (B) to podosomes. Cells were treated with or without PDBu 500 nM for 30 min. F-actinwas stained with Oregon Green 488 Phalloidin (Green). PKC isoforms and their phosphorylated forms were stained with specific antibodies and revealed with AlexaFluor 594 labeled secondary antibody (Red). Co-localization of PKC isoform with F-actin in small dots and ring structures was shown with small boxes and enlarged in the underneath panel. Circular ruffles were indicated with arrowheads. The overlapping of F-actin staining with PKC isoforms (C) and their phosphorylated forms (D) were quantified and expressed as Pearson’s correlation coefficients. *: P<0.05 compared with “no treatment” group.

124

7.3.2 PDBu-induced podosome assembly depends on cPKCs

To determine the proteolytic activity of PDBu-induced podosomes, we examined gelatin

matrix degradation by in situ zymography. After 60 min of stimulation with PDBu, enriched

podosome like small dots were revealed with F-actin staining (red) in the cytoplasm, which were

co-localized with or surrounded by digested area (shown as black holes) on gelatin pre-coated

matrix (green). The depth of the podosomes and digested matrix can be seen through X-Z and Y-

Z sections (Figure 7-3A).

To determine roles of PKC isoenzymes in PDBu-induced podosome formation and

gelatin matrix degradation, cells were pre-treated with chemical inhibitors for 1 h and then

challenged with PDBu (500 nM for 8 h). As shown in Figure 7-3B, pretreatment of cells with

BIM I and Ro-31-8220, which are blockers of both cPKCs and nPKCs (412, 413), effectively

inhibited PDBu-induced disassembly of stress fibers and formation of podosomes. Similarly,

Gö6976, a specific pharmacological inhibitor for Ca2+-dependent PKC and PKC I (413), and

BAPTA/AM, a membrane-permeable Ca2+ chelator effectively reduced PDBu-induced

podosomes. In contrast, Rottlerin, a cell-permeable PKC inhibitor that exhibits greater specificity

for PKC and PKC (414), had little effect (Figure 7-3B). Quantification of podosome number

per cell and the percentage of cells with podosomes confirmed the inhibitory effects of these

inhibitors (Figure 7-3C and 7-3D).

Fig 7-3

F-ac

tinO

verla

y

BIM IControl Gö6976 RottlerinB

020406080

100

% Cells Degrading Matrix

* * * *

*

AF-

actin

Mat

rixO

verla

y

10 µm

10 µm

10 µm

D

020406080

100

Con

trol

DM

SO

BIM

IR

o-31

-822

0B

AP

TA/A

MG

ö697

6R

ottle

rinC

ontro

lD

MS

OB

IM I

Ro-

31-8

220

BA

PTA

/AM

Gö6

976

Rot

tlerin

% Cells with Podosomes

No Treat PDBu

** *

*

E

0

10

20

30

40 Podosome # /Cell

C

**

**

125

126

Figure 7-3. PDBu-induced podosome formation is mainly mediated through classical PKC, but matrix degradation can be blocked by multiple PKC inhibitors.

(A) PDBu stimulation (500 nM, 60 min) induced podosomes (actin rich small dots, Red, indicated by arrows) were closely surrounded by degraded (Black) gelatin matrix (Green). (B) BIM I, Gö6976, but not Rottlerin, prevented PDBu-induced podosomeformation; however, all three inhibitors reduced matrix degradation. Cells cultured on gelatin matrix (Green) were pre-incubated with each inhibitor 1 µM for 1 h, and then stimulated with PDBu (500 nM for 8 h). F-actin was stained with RhodaminePhalloidin (Red). Digestion of gelatin matrix was revealed as black dots. (C) Quantitative analysis of podosome number per cell. (D) Quantitative analysis of number of cells with podosomes. (E) Quantitative analysis of matrix degradation. *: p < 0.05 in comparison with PDBu-treated control and DMSO (vehicle control) groups.

127

In control and DMSO (vehicle only) pre-treated groups, cells aggressively degraded the

gelatin matrix upon PDBu treatment (Figure 7-3B). Surprisingly, all the inhibitors tested

significantly prevented PDBu-induced matrix degradation compared to control and DMSO

groups (Figure 7-3B). Quantification of cells with gelatin degradation is shown in Figure 7-3E.

This observation is intriguing because it suggests that podosome formation is controlled by

cPKCs, but ECM degradation by the formed podosomes can be further regulated by nPKCs and

perhaps other PKC isozymes.

7.3.3 PDBu-induced matrix degradation can be blocked by siRNAs of multiple PKC

isoforms

Since chemical inhibitors may have non-specific effects (415), we then employed

siRNAs to knock down PKC isoforms to determine the roles of selected PKCs in podosome

formation and gelatin matrix degradation. As a negative control, a fluorescein conjugated control

siRNA that has no known homology to mammalian genes was used. The efficiency of siRNA

transfection was >90% as determined by the % of green fluorescent cells (data not shown).

PKC , , , and µ were chosen as representative of cPKC, nPKC, aPKC and PKC , respectively.

siRNAs reduced the expression of the targeted PKC protein without significant effects on other

PKCs, as shown by western blotting (Figure 7-4B). Only PKC siRNA significantly inhibited

PDBu-induced podosomes (Figure 7-4A and 7-4C and 7-4D), which supports the specific role of

cPKCs in controlling PDBu-induced podosome formation. Interestingly, siRNA-mediated

knockdown of PKC , PKC , PKC , or PKC significantly prevented PDBu-induced gelatin

degradation (Figure 7-4A and 7-4E) revealed by in situ zymography.

020406080

100


Fig 7-4

** * *

F-ac

tinO

verla

y

PKC siRNAControl siRNA PKC siRNAA

D % Cells with Podosomes

Con

trol

Con

trol s

iRN

A

PK

Csi

RN

A

PK

Csi

RN

A

PK

Csi

RN

A

PKC

µsi

RN

A

Con

trol

Con

trol s

iRN

A

PK

Csi

RN

A

PK

Csi

RN

A

PK

Csi

RN

A

PKC

µsi

RN

A

020406080

100

No Treat PDBu

*

E

B

PK

Csi

RN

A

PK

Csi

RN

A

PK

Cµ

siR

NA

PK

Csi

RN

A

PKCPKCPKCµPKCGAPDH

b1b2b3b4b5

Con

trol s

iRN

A

0

10

20

3035

25

15

5

C Podosome # /Cell

*

128

129

Figure 7-4. PDBu-induced podosome formation is reduced by PKC siRNA but matrix degradation can be blocked by multiple PKC siRNA.

BEAS2B cells cultured on gelatin matrix (Green) were transfected with siRNA for 48 h and then stimulated with PDBu (500 nM for 8 h). (A) Representative staining of control, PKC , and PKC siRNA-treated cells. (B) Western blots to show specificity of siRNAs. (C) Quantitative analysis showing that only PKC siRNA significantly reduced the number of podosomes per cell. (D) Quantitative analysis showing that PKC siRNA also significantly reduced the number of cells with podosomes. (E) Quantitative analysis showing that siRNA for PKC , , , and reduced PDBu-induced matrix degradation. *: p < 0.05 in comparison with PDBu-treated control and control siRNA groups.

130

These results further suggest that the presence of podosome like structures may not

necessarily lead to ECM degradation. The latter could be a process regulated by multiple PKC

isoforms.

7.3.4 PKC mediates PDBu-induced proteolytic activity of podosomes, MMP-9 recruitment

to podosome, and MMP-9 release and activation

We were particularly intrigued by the fact that siRNA targeting PKC reduced PDBu-

induced gelatin matrix degradation. This atypical PKC does not have lipid-binding region, hence,

it must be a down-stream target of cPKC and/or nPKC isozymes. Thus, we investigated the role

of PKC kinase activity in PDBu-induced podosome formation and ECM degradation.

BEAS2B cells were pre-incubated with myristoylated PKC pseudosubstrate PS (which

is cell permeable) to inhibit its activity, or the negative control PI (which is also myristoylated

and cell permeable), followed by treatment with 500 nM PDBu for 8 h. We confirmed the

specific inhibitory effect of PS but not PI on the phosphorylation of PKC by western blotting

(data not shown). These pre-treatment did not block PDBu-induced podosome formation.

However, inhibition of PKC kinase activity by its pseudosubstrate (PS) (416) significantly

prevented the matrix degradation (Figure 7-5A and 7-5B). The negative control peptide (PI) had

no such effect.

F-ac

tinO

verla

y

Control PS PI

Fig 7-5

A

B

Con

trol PS PI

Con

trol PS PI


No Treat PDBu

020406080

100

*

PI PS

F-ac

tinM

MP

-9

C D

00.20.40.60.8

1

PI PS

ControlPDBu

-0.20

0.20.40.60.8

1

00.20.40.60.8

1

Pear

son'

s C

orre

latio

n C

oeffi

cien

t

MMP-2

MMP-9

MMP-14

*

*

131

132

Figure 7-5. PKC regulates proteolytic activity of PDBu-induced podosomesand MMP-9 translocation to podosomes.

(A) PS, a specific pseudosubstrate for PKC (but not PI, the scrambled negative control peptide for PS) prevented the proteolytic activity of PDBu-induced podosomes. Cells cultured on gelatin matrix (Green) were pre-incubated with peptide (1 µM for 1 h), and then stimulated with PDBu (500 nM for 8 h). F-actin was stained with Rhodamine Phalloidin (Red). Digestion of gelatin matrix was revealed as black areas. (B) Quantitative analysis of percentage of cells showing matrix degradation. *: p < 0.05 in comparison with the PDBu-treated control group. (C) PS but not PI blocked MMP-9 translocation to podosomes. Note the MMP-9 (lower panel) was not clustered as actin rich small dots (higher panel). (D) PS but not PI reduced translocation of MMP-9 and MMP-14 to podosomes. *: p < 0.05 in comparison with the PDBu-treated PI control group.

133

To identify MMP isozymes regulated by PKC , whole cell lysates and culture medium

were collected for gelatin gel zymography assay. In the whole cell lysates, the expression and

activity of MMP-9 were higher than MMP-2, whereas in the cell culture medium, MMP-2 was

the major form revealed by the zymography (Figure 7-6A and 7-6B and 7-6C). The level of both

pro- and active-MMP-2 had no dramatic changes upon PDBu stimulation, while the level of pro-

and active-MMP-9 in culture medium were increased after PDBu challenge in a time-dependent

manner (Figure 7-6A). PKC pseudosubstrate PS, but not the negative control PI, inhibited

PDBu-induced secretion and activation of MMP-9 (Figure 7-6B). Similarly, compared to control

siRNA, PKC siRNA also inhibited PDBu-induced MMP-9 secretion and activation (Figure 7-

6C). Results from repeated experiments are quantified by densitometry (Figure 7-6D and 7-6E).

To investigate whether PKC pseudosubstrate affects on MMP gene expression,

BEAS2B cells were pre-incubated with PKC pseudosubstrate PS or the negative control PI.

Total RNA was extracted for qRT-PCR assay after PDBu stimulation (500 nM for 4 h). PDBu

stimulation increased MMP-9 mRNA approximately 30-35 folds, increased MMP-14 mRNA

around 2-3 folds, but had no effect on MMP-2 gene expression; however, PKC pseudosubstrate

PS did not inhibit PDBu-induced gene expression of MMP-9 or MMP-14 (Figure 7-6F). PKC

siRNA also did not inhibit PDBu-induced gene expression of these MMPs (data not shown).

We have previously shown recruitment of MMP-2, MMP-9, and MMP-14 to podosmes

after PDBu stimulation (122). To determine the role of PKC on these processes,

immunostaining together with F-actin staining was performed. The co-localization of MMPs in

actin-rich small dots was quantified. PKC pseudosubstrate PS but not PI significantly reduced

PDBu-induced translocation of MMP-9 and MMP-14 (but not MMP-2) to podosomes (Figure 7-

5C and 7-5D).

Control PS PI Control PS PI

No Treat PDBu

Pro-MMP-9

Pro-MMP-2

Active MMP-9

Active MMP-2

B C

ControlControl siRNA Control

No Treat PDBu

PKCsiRNA

Control siRNA

PKCsiRNAMedium

Pro-MMP-9

Pro-MMP-2

Active MMP-9

Active MMP-2

WCL

Pro-MMP-9

Pro-MMP-2

Active MMP-9

Active MMP-2

Medium 0 0.5 1 2 4 6 8 0 0.5 1 2 4 6 8 h

No Treat PDBu

9288

72

68

MW (KD)

92887268

MW (KD)

9288

7268

00.5

11.5

2

00.5

11.5

2

Con

trol

PS PI

PDBu

Fold

s of

Cha

nge

of M

MP

-9

Medium

WCL

*

*

00.5

11.5

2

00.5

11.5

2

Fold

s of

Cha

nge

of M

MP

-9

Medium

WCL

*

*

Con

trol

Con

trol s

iRN

A

PK

Csi

RN

A

PDBu

Fig 7-6A

D E

1020304001234

0

01234

Con

trol PS PI

Con

trol PS PI

MMP-2

MMP-9

MMP-14

No Treat PDBu

Fold

s of

Cha

nges

of G

ene

Exp

ress

ion

Leve

l

F mRNA

134

135

Figure 7-6. PKC regulates MMP-9 release and activation.

(A) PDBu induced MMP-9 secretion and enzymatic activity time-dependently. Cells were challenged with PDBu 500 nM from 0 to 8 h. The culture medium was examined with gelatin gel zymography. (B) PS reduced PDBu-induced MMP-9 secretion and enzymatic activity. Cells were pretreated with PS or PI (1 µM for 1 h) and challenged with PDBu 500 nM for 8 h. The culture medium and whole cell lysates (WCL) were examined with gelatin gel zymography. The MMP-2 and -9 bands were as indicated. (C) PKC siRNA reduced PDBu-induced MMP-9 secretion and enzymatic activity. Cells were transfected with control or PKC siRNA and challenged with PDBu 500 nM for 8 h. Whole cell lysates and culture medium were examined with gelatin gel zymography. (D) Quantitative analysis of MMP-9 bands. *: p < 0.05 in comparison with the PDBu-treated control and PI groups. (E) Quantitative analysis of MMP-9 bands. *: p < 0.05 in comparison with the PDBu-treated control and control siRNAgroups. (F) PS or PI did not affect PDBu-induced MMP gene expression. Cells were pretreated with PS or PI and challenged with PDBu for 4 h. RNA was extracted and qRT-PCR was performed. qRT-PCR was performed by Xiao-Hui Bai.

136

7.3.5 PDBu-induced MMP-9 is involved in degradation of gelatin matrix

To determine which MMP is responsible for the PDBu-induced proteolytic activity of

podosomes, we transfected cells with siRNAs for MMP-2, MMP-9, or MMP-14, and examined

the gelatin matrix degradation by in situ zymography. Cells were transfected with siRNA for 48

h and then challenged with 500 nM PDBu for 8 h. Each siRNA reduced the protein level of

targeted MMP and had no effects on other two MMPs examined (Figure 7-7B). In MMP-9 or

MMP-14 siRNA transfected group, although cells still formed podosomes upon PDBu

stimulation, the proteolytic activity was significantly inhibited compared to control siRNA and

MMP-2 siRNA groups. In contrast, the inhibitory effect of MMP-2 siRNA on gelatin matrix

degradation was not statistically significant (Figure 7-7A and 7-7C). To further confirm the role

of secreted MMP-9 in digesting gelatin matrix, we used neutralization antibody for MMP-9. In

comparison with mouse IgG as the negative control, MMP-9 antibody pre-incubation

significantly reduced the enzymatic activity without affecting podosome formation (Figure 7-7D

and 7-7E).

F-ac

tinO

verla

yControl siRNA MMP-2 siRNA MMP-9 siRNA MMP-14 siRNA

F-ac

tinO

verla

y

mIg G MMP-9 AB

020406080

100

Con

trol s

iRN

A

MM

P-2

siR

NA

MM

P-9

siR

NA

MM

P-1

4 si

RN

A

% C

ells

Deg

radi

ng M

atrix

mIg

G

MM

P-9

AB

mIg

G

MM

P-9

AB

% C

ells

Deg

radi

ng M

atrix

No Treat PDBu

No Treat PDBu

Fig 7-7

A

C

D

E

* *

020406080

100

*

Con

trol s

iRN

A

MM

P-2

siR

NA

MM

P-9

siR

NA

MM

P-1

4 si

RN

AMMP-2

MMP-9

GAPDH

MMP-14

Con

trol s

iRN

A

MM

P-2

siR

NA

MM

P-9

siR

NA

MM

P-1

4 si

RN

A

B

b1

b2

b3

b4

137

138

Figure 7-7. Role of MMP-9 and MMP-14 in PDBu-induced proteolytic activity of podosomes.

(A) siRNA for MMP-9 and MMP-14 (but not for MMP-2) prevented matrix degradationof PDBu-induced podosomes. BEAS2B cells cultured on gelatin matrix (Green) were transfected with siRNA for 48 h, and then stimulated with PDBu 500 nM for 8 h. F-actin was stained with Rhodamine Phalloidin (Red). Digestion of gelatin matrix was revealed as black areas. (B) The specificity of siRNAs for MMPs was determined with western blotting. (C) Quantitative analysis of percentage of cells showing matrix degradation. *: p < 0.05 in comparison with the PDBu-treated control siRNA group. (D) Neutralization antibody for MMP-9 prevented PDBu-induced matrix degradation of podosomes. Cells cultured on gelatin matrix (Green) were pre-treated with either MMP-9 neutralization antibody or mouse IgG (10 ng/ml for 8 h) and then stimulated with PDBu 500 nM for 8 h. (E) Quantitative analysis of percentage of cells showing matrix degradation. *: p < 0.05 in comparison with the PDBu-treated mIgG group.

139

7.3.6 Recruitment of PKC to PDBu-induced posodosmes depended on nPKC

To determine how PKC was recruited to podosomes in response to PDBu, we expressed

PKC -GFP in BEAS2B cells and performed live cell imaging. PKC -GFP appeared as small dots

in the cytoplasm and at the peripheral ruffles after PDBu stimulation, and similar as podosomes

these small dots showed fast turnover (Figure 7-9A upper panel). Tks5/Fish has been shown to

an important component of podosomes in invasive cancer cells (148). Indeed, we also found

Tks5/Fish in PDBu-induced small dots and rings in BEAS2B cells (Figure 7-8A). When PKC -

GFP and Tks5-mCherry were co-expressed in BEAS2B cells, they were well co-localized in the

cytoplasm as small dots and rings (Figure 7-8B). Interestingly, nPKC inhibitor Rottlerin pre-

treatment prevented PDBu-induced translocation of PKC -GFP to podosomes (Figure 7-8B and

7-9A lower panel). Data are quantified in Figure 7-9D. In PKC siRNA (but not in control

siRNA) transfected cells, PDBu-induced translocation of PKC (Figure 7-9B and 7-9E) or

MMP-9 (Figure 7-9C and 7-9F) to actin-rich small dots was significantly reduced. Rottlerin and

PKC siRNA also prevented PDBu-induced PKC phosphorylation (Figure 7-9G and 7-9H).

Rottlerin and PKC siRNA did not significantly inhibited MMP-9 mRNA expression (Figure 7-

9I and 7-9J). These results suggest that nPKC, especially PKC , is involved in the recruitment of

PKC to podosomes, and its subsequent regulatory effects on MMP-9 recruitment and gene

expression.

F-actin Tks5

00.20.40.60.8

1

No Treat PDBu

Pear

son'

s C

orre

latio

n C

oeffi

cien

t

A

C

Fig 7-8

PKC -GFP Tks5-mCherry Overlay DIC

Med

ium

+PD

BuR

ottle

rin+P

DBu

B

00.20.40.60.8

1

Medium+PDBu

Rottlerin+PDBu

Pear

son'

s C

orre

latio

n C

oeffi

cien

tD

**

Tks5 Tks5-mCherry vs PKC -GFP

140

141

Figure 7-8. Rottlerin reduced PDBu-induced translocation of PKC topodosomes.

(A) PDBu induced translocation of Tks5 to podosomes. BEAS2B cells were treated with or without PDBu 500 nM for 30 min. F-actin was stained with Oregon Green 488 Phalloidin (Green). Tks5 was stained with specific antibody and revealed with AlexaFluor 594 labeled secondary antibody (Red). Co-localization of Tks5 with F-actin in small dots was shown with small boxes, which were enlarged as single channel images in lower panel. The 3D structure of podosomes was also shown as Z-stacks of scanning confocal images on the right. (B) Rottlerin, an nPKC inhibitor reduced PDBu-induced colocalization between PKC -GFP and Tks5-mCherry (as a marker of podosomes). Cells were co-transfected with vectors expressing PKC -GFP and Tks5-mCherry, pre-treated with or without Rottlerin (1 µM for 1 h) and then stimulated with PDBu (500 nM for 30 min). (C) The overlap of F-actin staining with Tks5 was quantified and expressed as Pearson’s correlation coefficients. *: p < 0.05 in comparison with untreated control group. (D) Co-localization of PKC -GFP with Tks5-mCherry was quantified and expressed as Pearson’s correlation coefficient values. *: p < 0.05 in comparison with only PDBu-treated group.

Fig 7- 9

0 min 11 min 13 min 14 min 19 min

0 min 8 min 14 min 20 min 25 min

Med

ium

+PD

BuR

ottle

rin+P

DBu

Con

trol s

iRN

APK

Csi

RN

A

PKC -GFP

PKC -GFP

PKCF-actin MMP-9F-actin

A

B

No Treat

Con

trol

Con

trol s

iRN

AP

KC

siR

NA

PDBuC

ontro

lC

ontro

l siR

NA

PK

Csi

RN

A

(p)PKC

GAPDH

DNo Treat

Con

trol

PDBu

Con

trol

DM

SO

Rot

tlerin

DM

SO

Rot

tlerin

010203040

Control Rottlerin

No Treat

PDBu

0204060

MMP-9 mRNAMMP-9 mRNA

C

0

5

10

15

20

25

0 2 4 6 8 10 12 14 16 18 20 22 24 26Time (min)

# of

PK

C-G

FP S

mal

lDot

s Medium

Medium+PDBu

Rottlerin

Rottlerin+PDBu

E

F

G H

I J

ControlsiRNA

PKCsiRNA0

0.20.40.60.8

1

ControlsiRNA

PKC

No TreatPDBu

siRNA

PKC

Pear

son'

s C

oeffi

cien

t

00.20.40.60.8

1

Pear

son'

s C

oeffi

cien

t

ControlsiRNA

PKCsiRNA

MMP-9

* *

142

143

Figure 7-9. PKC controls recruitment of PKC and MMP-9 to podosomes.

(A) Rottlerin (1 µM), an nPKC inhibitor, prevented PDBu-induced PKC -GFP to form small dots in cytoplasm. BEAS2B cells transfected with pEGFP-PKC were subjected to PDBu (500 nM) stimulation. Images from time-lapse confocal microscopy show that PKC -GFP is clustered as small dots in cytoplasm and circular ruffles at the peripheral membrane. (B) PKC siRNA inhibited PKC translocation to PDBu-induced podosomes. Note the PKC (right panel) was not clustered at actin rich small dots (left panel). (C) PKC siRNA abolished MMP-9 translocation to PDBu-induced podosomes. Note the MMP-9 (right panel) was not clustered at actin rich small dots (left panel). (D) Quantitative analysis of the number of PKC -GFP small dots in BEAS2B cells from live cell imaging. Co-localization of F-actin with PKC (E) or with MMP-9 (F) was measured and expressed as Pearson’s correlation coefficients. *: p < 0.05 in comparison with the PDBu-treated control siRNA group. (G) Rottlerinprevented PDBu-induced phosphorylation of PKC . Cells were pre-incubated with Rottlerin (1, 5, 10 µM) for 1 h before challenged with PDBu 500 nM for 30 min. (H) PKC siRNA reduced phosphorylation of PKC . BAES2B cells were transfected with control siRNA or PKC siRNA for 48 h and then challenged with PDBu 500 nM for 30 min. (I) Rottlerin did not significantly reduced PDBu-induced MMP-9 mRNA expression. Cells were pretreated with Rottlerin 1 µM for 1 h and challenged with 500 nM PDBu for 4 h. (J) PKC siRNA did not significantly reduced PDBu-induced MMP-9 mRNA expression. Cells were transfected with control or PKC siRNA for 48 h and challenged with 500 nM PDBu for 4 h.

144

7.4 Discussion

The PKC family of serine-threonine kinases is critical signal transducers participating in

many agonist-induced signaling cascades (292), as well as in mechanotransduction in lung cells

(417). In the present study, we found that PDBu-induced formation of podosomes and the

degradation of ECM are two separate steps, and different PKC isoforms play distinct but

coordinated roles in these processes in normal human bronchial epithelial cells.

Our results demonstrated that inhibition of cPKCs, prevented podosome formation and

subsequently gelatin matrix degradation. It has been shown that both PKC and PKC were

necessary for podosome assembly in human endothelial cells; however only constitutively active

PKC A25E but not PKC A147E could mimic the effect of PMA to induce podosome formation

(376). It is possible that in phorbol ester-induced podosomes, cPKC subfamily is the most direct

effector of these lipid ligands. The phosphorylation status of cPKCs was not significantly

changed during PDBu stimulation; however, cPKCs were found in podosomes and circular

ruffles detected by antibodies against PKC , and phosphorylated cPKCs. It has been proposed

that mature and phosphorylated cPKCs can be kept in the cytosol in an inactive conformation by

intramolecular interactions between the N-terminal pseudosubstrate region and the kinase

domain. The latent kinase can be recruited to the membrane and activated by lipid ligands (such

as phorbol ester), leading to a massive conformational change that releases the pseudosubstrate

domain from the substrate-binding site, allowing for substrate binding, phosphorylation and

activation of downstream signaling effectors (302, 303). Therefore, although the phosphorylation

status of cPKCs is not significantly altered upon PDBu stimulation, redistribution of latent

phosphorylated cPKC may lead to their activation and subsequent initiation of podosome

145

formation and function. Although we do not have direct evidence to support the existence of

latent cPKCs, our results can be explained by this concept well.

It has been proposed that the distribution of PKC isoforms in different cellular

compartments determines their functions by presenting them to different substrates and

interaction partners (418). We were surprised to see that the phosphorylation status of novel and

atypical subfamilies of PKCs and PKCµ increased dramatically upon PDBu stimulation, and

multiple phosphorylated PKCs were recruited to the sites of podosomes and circular ruffles,

instead of to different compartments in the cell. Down-regulation of PKC , , , and with

siRNA prevented PDBu-induced gelatin matrix degradation. Clearly, these PKC family members

were not only clustered together but also involved in the function of podosomes. Indeed,

Rottlerin (nPKC inhibitor) and PKC siRNA prevented PKC and MMP-9 recruitment to

podosomes. These results suggest that the function of podosomes requires the synergy of

multiple PKC family members.

One of the interesting findings of the present study is that the formation of podosomes

and the proteolytic degradation function of podosomes can be uncoupled. Rottlerin, siRNAs for

PKC , , and , siRNAs for MMP-9 and MMP-14, and PKC pseudosubstrate, reduced PDBu-

induced gelatin matrix degradation without affecting podosome formation. Our data are in

agreement with the recent report that MMP-14 deficient mouse bone marrow cells and spleen

dendritic cells produced podosomes but lost the ability to degrade extracellular matrix after TLR

signaling (419). It is possible that actin-rich podosome structures only provide a platform to

recruit other proteins, and the activities of MMPs and other enzymes are further regulated locally

at the sites of these structures.

146

Interestingly, we found that PKC is involved in controlling the proteolytic function of

PDBu-induced podosomes, mainly through MMP-9. Both PKC siRNA and PKC

pseudosubstrate reduced PDBu-induced secretion of MMP-9 to the culture medium and its

conversion from pro- to active form. In rat C6 glioma cells, IL-1 and TNF induced gene

expression of MMP-9 via PKC through NF B (237). In contrast, we found that in BEAS2B

cells PKC pseudosubstrate did not block PDBu-induced MMP-9 gene expression. On the other

hand, PKC pseudosubstrate blocked PDBu-induced translocation of MMP-9 and to less extend

of MMP-14 to podosomes. This is a novel finding in podosome studies.

Our results suggest that MMP-9 is the major MMP in PDBu-induced gelatin matrix

degradation in BEAS2B cells. The expression of MMP-14 is also up-regulated by PDBu, and

MMP-14 siRNA partially reduced PDBu-induced gelatin matrix degradation. The MMP-2 level

in the cell lysates is relatively lower than that of MMP-9, but higher in the culture medium,

suggesting that it is constitutively released. Our data also show that MMP-2 is not significantly

involved in PDBu-induced gelatin matrix degradation. Of course, we cannot exclude the

possibility that MMP-2 may be important for the degradation of other types of matrix molecules.

IL-1 induced an MMP activation cascade (MMP-14 MMP-13 MMP-9) in hepatic stellate

cells in 3D matrix (420). In non-malignant monkey kidney epithelial cells, pro-MMP-9

activation was accomplished via a cascade of zymogen activation initiated by MMP-14 and

mediated by MMP-2 (421). The proteolytic activity of PMA-induced podosomes in endothelial

cells depended on MMP-14 mediated activation of MMP-2 but not MMP-9 (422). The

interaction among MMP molecules is another interesting area of research.

Taken together, our results suggest that cPKC (particularly PKC ) can regulate rapid

podosome formation in normal human bronchial epithelial cells as an upstream signal; atypical

147

PKC may be the direct effector at the site of podosomes that modulates the recruitment, release,

and activation of MMP-9. The coordinated actions of PKC isozymes, of course with different

regimen, may also exist in podosomes formed by other cell types under different conditions.

148

8. Chapter Eight. Coordinated regulation of PKC activation-induced podosome formation

and proteolytic activities of PI3K, Src and MAPK pathways in human bronchial epithelial

cells

The content of this chapter was prepared for an original research article in Journal of Cell

Science:

Xiao H, Kagzi W, Bai XH, Mak AS, Liu M. Coordinated regulation of PKC activation-induced

podosome formation and proteolytic activities of PI3K, Src and MAPK pathways in human

bronchial epithelial cells. Journal of Cell Science. 2009.

149

8. Chapter Eight. Coordinated regulation of PKC activation-induced podosome formation

and proteolytic activities through PI3K, Src and MAPK pathways in human bronchial

epithelial cells

8.1 Summary

Podosomes are cellular structures, which migratory and actively invading cells rely on to

break through the extracellular matrix. In chapter 7, we reported that phorbol 12, 13-dibutyrate

(PDBu), a protein kinase C (PKC) activator, induced podosome formation in normal human

bronchial epithelial cells, through PKC to recruit and activate MMP-9 at the podosomes. The

objective of this chapter of study was to explore signaling pathways that are involved in PKC

activation-induced podosome formation and matrix degradation. Herein, we reported that PDBu

increased phosphorylation of PI3K p85, PDK1, Akt, GSK3 , Src, ERK1/2 and JNK. PI3K p85,

phosphorylated Akt, Src, ERK and JNK were recruited to podosomes. Inhibitors for PI3K, Akt

and Src suppressed PDBu-induced podosome formation and matrix degradation. In contrast,

blockers for MEK/ERK or JNK did not inhibit podosome formation but reduced proteolytic

activity of podosomes, decreased MMP-9 recruitment to podosomes, and its release and

activation. Further inhibition studies demonstrated that PI3K/Akt and Src are upstream of

ERK1/2 and JNK. These results indicate that podosome assembly and podosomal proteolytic

activity are regulated by multiple signal transduction pathways at different levels in a

coordinated fashion.

150

8.2 Introduction

Cell migration plays important role in lung development, growth, and repair after

injuries. Podosomes are actin-rich structures located at the ventral surface of the cell membrane

that migratory and actively invading cells rely on to break through the extracellular matrix (163).

Podosomes have been mainly reported in mesenchymal cells (such as fibroblasts, osteoclasts,

smooth muscles cells and endothelial cells) (150) and cancer cells (148, 172). Recently, we

reported that phorbol 12,13-dibutyrate (PDBu), a protein kinase C (PKC) activator, induced

podosome formation in normal human bronchial epithelial cells, which may be important for

airway development and repair (122). We further found that while the conventional PKCs

(especially PKC ) control PDBu-induced podosome assembly, it is the atypical PKC that

regulates the gelatin matrix degradation by influencing the recruitment of MMP-9 at the

podosomes and its release and activation (423). PKC activation also induced podosomes in

smooth muscle cells (424), endothelial cells (147, 425), and fibroblasts (426).

It has been shown that Src, PI3K and MAPK (mitogen-activated protein kinases)

pathways can be activated by PKC and involved in podosome formation (214, 376). The class Ia

PI3K has a regulatory subunit (e.g. p85) and a catalytic subunit (e.g. p110). Phosphorylation of

the p85 subunit can lead to the activation of the catalytic subunit for PI(3,4,5)P3 formation (200,

427), which can bind PH domain containing proteins, such as Akt (190, 191). Activated Akt can

modulate cell motility, cell polarity and cortical actin structures (102). PDK1 can phosphorylate

Akt for its activation (192-194). Non-receptor Src family tyrosine kinases play a central role in

podosome regulation (154). It has been shown in fibroblasts phorbol ester PMA can recruit

AFAP (actin filament associated protein) to PKC , and AFAP can then recruit and activate c-Src

for podosome formation (201, 377). PI3K activation is required for this PKC-directed activation

151

of c-Src by AFAP (201). MAPK signaling pathways are also important for gene expression, cell

polarity (428), and cell invasion (429). In vascular smooth muscle cells (213, 214, 430), murine

bone marrow-derived dendritic cells (215), or bone marrow-derived neutrophils (216), ERK1/2

have been reported to mediate podosome formation. Big MAPK Erk5 promoted Src-induced

podosome formation in fibroblasts (217).

Most of these previous studies focused on podosome formation. Recent studies suggest

that the proteolytic activity of podosomes can be further regulated by multiple mechanisms after

podosome formation (419, 431). The objective of the present study was to determine the

involvement of PI3K/Akt, Src and MAPK pathways in PKC-induced proteolytic activity, as well

as podosome formation, in normal human bronchial epithelial cells.

8.3 Results

8.3.1 PDBu-induced protein phosphorylation and translocation of PI3K/Akt related

signaling molecules

To determine whether PI3K/Akt pathway is activated upon PDBu stimulation, we

examined the phosphorylation status of signaling proteins after challenging human bronchial

epithelial BEAS2B cells with 500 nM PDBu from 0 to 120 min. The phosphorylation of PI3K

p85 subunit was increased and reached the maximum at 60 min. The phosphorylation of PDK1

was also increased gradually. The phosphorylation of AktS473 increased to the plateau at 30

min. The phosphorylation of GSK3 , a down-stream signal of Akt, was also gradually enhanced

after PDBu treatment (Figure 8-1A).

(p)PI3Kp85T458

PDBu 500 nM

0 1 5 10 20 30 60 120 min

(p)AktS473

(p)PDK1S241

(p)GSK3 S9

A

B

GAPDH

Fig 8-1

PI3Kp85 (p)PDK1S241

Con

trol

PD

Bu

(p)AktS473 (p)GSK3 S9

a1

a2

a3

a4

a5p85 (p)PDK1 (p)Akt (p)GSK3

No TreatPDBu

C

00.20.40.60.8

1

Pearson's Correlation Coefficient

152

* *

Figure 8-1. PDBu-induced phosphorylation and translocation of PI3K/Akt related proteins.

(A) PDBu-induced protein phosphorylation. BEAS2B cells were treated with PDBu500 nM from 0 min to 120 min. The phosphorylation of PI3K p85, PDK1, Akt and GSk3 was examined with western blotting. (B) PDBu-induced translocation of PI3K/Akt related proteins to podosomes and circular ruffles. Cells were treated without or with PDBu 500 nM for 30 min. F-actin was stained with Oregon Green 488 Phalloidin (Green). The target proteins were stained with specific antibodies and revealed with Alexa Fluor 594 labeled secondary antibody (Red). Co-localization of proteins with F-actin in small dots and ring structures is shown with small boxes andenlarged in the underneath panel. Circular ruffles were indicated with arrowheads. (C) Quantitative analysis of target proteins overlapping with F-actin staining by Pearson’s correlation coefficients. *: P<0.05 compared with “no treatment” group.

153

Next, we investigated the effects of PDBu-stimulation on distribution of PI3K/Akt related

signaling proteins. Under control condition, PI3K p85 , phosphorylated PDK1, Akt and GSK3

were localized in the cytoplasm and peri-nucleus region, but not overlapped with actin stress

fibers (Figure 8-1B). PDBu stimulation resulted in dramatic cytoskeleton reorganization, that is,

disassembly of actin stress fibers and formation of actin-rich small dots, rings and belts in the

cytoplasm and circular membrane ruffles at the peripheral (Figure 8-1B). The molecular

components and the 3-D structure of these podosomes and circular ruffles have been

characterized previously (122). PDBu treatment induced translocation of PI3K p85, and

phosphorylated Akt to the newly formed podosomes and circular ruffles (Figure 8-1B). The co-

localization of F-actin staining with these proteins were quantified and expressed as Pearson’s

correlation coefficient values (Figure 8-1C).

8.3.2 Effects of PDBu-stimulation on phosphorylation and distribution of Src and MAPKs

To determine whether c-Src and MAPKs are activated upon PDBu treatment, we

examined the phosphorylation of c-Src, ERK1/2, JNK and p38 after challenging the BEAS2B

cells with 500 nM from 0 to 120 min. The phosphorylation of SrcY416, a sign of c-Src

activation, was increased dramatically at 30 min. The phosphorylation of ERK1/2 was increased

during the first 10 minutes and slightly decreased thereafter. The phosphorylation of JNK was

increased dramatically within the first 30 min followed by a rapid decrease to the basal level at

the end of 120 min. The phosphorylation of p38 was not dramatically altered by PDBu treatment,

in comparison with other MAPK molecules examined (Figure 8-2A).

PDBu 500 nM

0 1 5 10 20 30 60 120 min

(p)p38T180/Y182

(p)ERKT202/Y204

GADPH

(p)JNKT183/Y185

(p)SrcY416

Fig 8-2

A

a1

a2

a3

a4

a5

BSrcY416 (p)ERK

Con

trol

PD

Bu

(p)JNK (p)p38

SrcY416 (p)ERK (p)JNK (p)p38

No Treat

PDBu

C

00.20.40.60.8

1


154

***

Figure 8-2. PDBu-induced phosphorylation and translocation of Src and MAPK signaling proteins.

(A) PDBu-induced phosphorylation of Src and MAPK signaling proteins. (B) PDBuinduced translocation of phosphorylated Src and MAPK molecules to podosomes. (C) Quantitative analysis of these proteins overlapping with podosomes by Pearson’s correlation coefficients. *: P<0.05 compared with “no treatment” group. See Figure 8-1 legend for experimental design.

155

Then, we examined the PDBu-stimulated re-distribution of c-Src and MAPKs.

Phosphorylated SrcY416, ERK1/2, JNK and p38 were mainly distributed in the cytoplasm and

peri-nucleus area under control condition. After PDBu stimulation, phosphorylated SrcY416 was

co-localized with podosomes and circular ruffles. Phosphorylated ERK1/2 and JNK (but not

p38) were also accumulated in PDBu-induced podosomes (Figure 8-2B). Quantification of the

co-localization of F-actin and these proteins showed significant increased phosphorylated ERK

and JNK in podosome after PDBu stimulation (Figure 8-2C).

8.3.3 PI3K/Akt and Src regulates podosome assembly and proteolytic activity

To determine the role of PI3K/Akt signaling in PDBu-induced podosome, we pre-

incubated cells with inhibitors (1 µM for 1 h) for PI3K (LY294002 or Wortmannin) or Akt (Akt

inhibitor II) before challenging them with PDBu (500 nM for 8 h). The effects of these inhibitors

on podosome formation and gelatin matrix degradation were determined with in situ zymography

assay with experimental conditions optimized (423). These inhibitors prevented PDBu-induced

disruption of actin stress fiber, reduced podosome formation (Figure 8-3A and 8-3B), and

blocked gelatin matrix degradation of PDBu-induced podosomes examined with in situ

zymography (Figure 8-3A and 8-3C).

A B Fig 8-3F-actin Overlay

Con

trol

LY29

4002

Wor

tman

nin

Akt

II

020406080

100

Con

trol

DM

SO

LY29

4002

Wor

tman

in

Akt

II

Con

trol

DM

SO

LY29

4002

Wor

tman

in

Akt

II

% Cells Degrade Matrix

No Treatment PDBu

0

20406080

100

% Cells With Podosomes

0

10

20

30

40

Con

trol

LY29

4002

PP

2A

ktII

Con

trol

LY29

4002

PP

2

Akt

II

MMP-9 mRNA

No Treatment PDBu

D

WCL

Medium

Pro-MMP-9

Pro-MMP-2

Active MMP-9

Active MMP-2

Pro-MMP-9

Pro-MMP-2

Active MMP-9

Active MMP-2

* * *

* * *

**

*

C

E

F

156

157

Figure 8-3. PI3K and Akt regulate podosome assembly and its proteolyticactivity through MMP-9 expression, release and activation.

(A) LY294002 and Wortmannin, inhibitors for PI3K, and Akt inhibitor II (Akt II), prevented PDBu-induced podosome formation and proteolytic activity of podosomes. Cells cultured on gelatin matrix (Green) were pre-incubated with each inhibitor (1 µMfor 1 h), and then stimulated with PDBu (500 nM for 8 h). F-actin was stained with Rhodamine Phalloidin (Red). Digestion of gelatin matrix was revealed as black areas.(B) Quantitative analysis of percentage of cells showing podosomes. (C) Quantitative analysis of percentage of cells showing matrix degradation. (D) LY294002, Akt II and PP2 (inhibitor for Src) attenuated PDBu-induced MMP-9 mRNA gene expression. Cells were pretreated with each inhibitor (1 µM for 1 h) and challenged with PDBu for 4 h. RNA was extracted and qRT-PCR was performed. *: p < 0.05 in comparison with PDBu-stimulated control group. (E and F) LY294002, Akt II and PP2 reduced PDBu-induced MMP-9 protein expression and enzymatic activity. Cells were pretreated with each inhibitor (1 µM for 1 h) and challenged with PDBu 500 nM for 8 h. The culture medium (F) and whole cell lysates (WCL) (E) were examined with gelatin gel zymography.

A BFig 8-4

F-actin Overlay

Con

trol

PP

2 020406080

100

Con

trol

DM

SO

Gen

iste

in

PP

2

PP

3

Su66

56

Con

trol

DM

SO

Gen

iste

in

PP

2

PP

3

Su66

56


No Treatment PDBu

020406080

100% Cells With Podosomes

**

*

* * *

C

158

Figure 8-4. Src is involved in podosome formation and proteolytic activity.

(A) PP2, inhibitor for Src, prevented PDBu-induced podosome formation and proteolytic activity of podosomes. See Figure 8-3 legend for experimental design. Similarly, Genistein (general inhibitor for tyrosine kinases), Su6656 (Src inhibitor), and PP3 (negative control for PP2) (1 µM 1 h each) were studied. (B) Quantitative analysis of percentage of cells showing podosomes. (C) Quantitative analysis of percentage of cells showing matrix degradation. *: p < 0.05 in comparison with PDBu-stimulated control group.

159

We have shown that MMP-9 expression, recruitment to podosomes, release and

activation are crucial for PDBu-induced proteolytic activities of podosomes (423). To examine

whether PI3K/Akt pathway affects MMP-9 gene expression, cells were pre-incubated with the

inhibitors (1 µM for 1 h), and total RNA was extracted for qRT-PCR assay after PDBu treatment

(500 nM for 4 h) (423). PDBu stimulation increased MMP-9 mRNA approximately 35-40 folds.

LY294002, Wortmannin, or Akt II almost completely blocked PDBu-induced MMP-9 mRNA

expression (Figure 8-3D). To investigate MMP isozymes regulated by PI3K/Akt pathway,

whole cell lysates and cell culture medium were collected for gelatin gel zymography assay. In

cell lysates, the expression and activity of MMP-9 were higher than MMP-2, while MMP-2 was

more prominent in the cell culture medium (Figure 8-3E and 8-3F). The expression and

activation of MMP-2 were not significantly altered upon PDBu treatment both in the cell lysates

and cell culture medium (Figure 8-3E and 8-3F). The latent and active forms of MMP-9 were

increased after PDBu challenge in both cell lysates and medium. LY294002 or Akt II

dramatically prevented the protein expression, activation and secretion of MMP-9 (Figure 8-3E

and 8-3F). The inhibitory effects are statistically significant (P<0.05) as quantified by

densitometry (data not shown).

To identify the role of Src in PDBu-induced podosome, we pre-incubated cells with

Genistein (a general inhibitor for tyrosine kinases), PP2 or Su6656 (specific inhibitors for Src) (1

µM for 1 h). Each of these inhibitors prevented PDBu-induced podosome assembly and gelatin

matrix degradation, whereas PP3, a non-functional analogue of PP2, did not show such

inhibitory effects (Figure 8-4). PP2 also significantly reduced PDBu-induced MMP-9 gene

expression (Figure 8-3D), protein synthesis, activation and release (Figure 8-3E and 8-3F). The

160

inhibitory effects are statistically significant (P<0.05) as quantified by densitometry (data not

shown).

8.3.4 ERK and JNK mediate proteolytic activity of PDBu-induced podosomes without

affecting podosome formation

To investigate whether MAPKs are involved in PDBu-induced podosomes, cells were

pre-treated with PD98059 (inhibitor for MEK, up-stream kinase of ERK), SP600125 (inhibitor

for JNK) and SB20358 (inhibitor for p38) (1 µM for 1 h) prior to the PDBu-stimulation. None of

these inhibitors prevented podosome formation upon PDBu challenge (Figure 8-5A and 8-5B).

Interestingly, PD98059 and SP600125 (but not SB20358) blocked the gelatin matrix degradation

(Figure 8-5A and 8-5C). Pre-treatment of cells with PD98059, SP600125 and SB20358

significantly inhibited PDBu-stimulated MMP-9 mRNA expression (Figure 8-5D). Pretreatment

of cells with PD98059 or SP600125 (to less extend with SB20358) also reduced PDBu-induced

MMP-9 protein expression and activation (Figure 8-5E and 8-5F).

To determine whether MAPKs play a role in MMP-9 recruitment to PDBu-induced

podosomes, we pre-incubated cells with these inhibitors and performed immunofluorescent

staining for MMP-9. The co-localization of MMP-9 in actin-rich small dots was quantified.

PD98059 and SP600125 but not SB20358 significantly reduced PDBu-induced translocation of

MMP-9 to podosomes (Figure 8-6).

AF-actin Overlay

Con

trol

PD98

059

SP60

0125

SB20

358

Fig 8-5

010203040

Con

trol

PD98

059

SP60

0125

SB20

358

Con

trol

PD98

059

SP60

0125

SB20

358

MMP-9 mRNA

No Treatment PDBu

WCL

Medium

D

Pro-MMP-9

Pro-MMP-2

Active MMP-9

Active MMP-2

Pro-MMP-9Active MMP-9

Active MMP-2Pro-MMP-2

020406080

100

% Cells With Podosomes

020406080

100

Con

trol

DM

SO

PD98

059

SP60

0125

SB20

358

Con

trol

DM

SO

PD98

059

SP60

0125

SB20

358


No Treatment PDBu

B

* *

* **

C

E

F

161

162

Figure 8-5. ERK and JNK mediate proteolytic activity of PDBu-induced podosomes.

(A) PD98059 (inhibitor for MEK, upstream kinase of ERK) and SP600125 (inhibitor for JNK) but not SB20358 (inhibitor for p38) (1 µM each for 1h), prevented proteolyticactivity of PDBu-induced podosomes, but not affect podosome formation. See Figure 8-3 legend for experimental design. (B) Quantitative analysis of percentage of cells showing podosome assembly. (C) Quantitative analysis of percentage of cells showing matrix degradation. (D) PD98059, SP600125 and SB20358 attenuated PDBu-induced MMP-9 mRNA gene expression. See Figure 8-3 legend for experimental design. *: p < 0.05 in comparison with PDBu-stimulated control group. (E) PD98059, SP600125 and SB20358 (to less extend) reduced PDBu-induced MMP-9 protein expression and enzymatic activity.

AF-actin MMP-9

Con

trol

PD98

059

SP60

0125

SB20

358

Fig 8-6B

Con

trol

PD98

059

SP60

0125

SB20

358

Con

trol

PD98

059

SP60

0125

SB20

358

No Treatment PDBu

0

0.2

0.4

0.6

0.8

1


* *

163

Figure 8-6. Inhibitors for ERK and JNK pathways prevented MMP-9 translocation to PDBu-induced podosomes.

(A) Cells were pre-incubated with PD98059, SP600125, or SB20358 (1 µM for 1 h), and then stimulated with PDBu (500 nM for 30 min). F-actin was stained with Oregon Green 488 Phalloidin (Green). MMP-9 was revealed by Alexa Fluor 549 labeled secondary antibody (Red). (B) Quantitative analysis of percentage of MMP-9 overlapping F-actin staining by Pearson’s correlation coefficients. *: p < 0.05 in comparison with PDBu-stimulated control group.

164

It has been shown that U0126, a specific inhibitor for MEK, blocked PMA-induced

podosome formation in smooth muscle cells (214), and in fibroblasts (217). In contrast, we found

that PD98059 only blocked proteolytic activity without affecting podosome formation in human

bronchial epithelial cells (Figure 8-5). Thus, we also pre-incubated BEAS2B cells with U0126 or

its negative control U0124 (1 µM for 1 h). Compared to U0124, U0126 greatly attenuated gelatin

matrix degradation of PDBu-stimulated podosomes without affecting podosome assembly

(Figure 8-7A, 8-7B and 8-7C). U0126, but not U0124, also dramatically limited the translocation

of MMP-9 to the sites of PDBu-induced podosomes (Figure 8-7D and 8-7E). These interesting

data indicated that podosome assembly and matrix degradation are regulated differently in

different cell types. It may be due to the distinct expression level and activation status of certain

unknown proteins or kinases in these models.

8.3.5 Crosstalk among PI3K/Akt, Src and MAPK pathways

Since the major effect of PDBu is to activate PKCs, we investigated whether PDBu-

induced phosphorylations of signal molecules examined in the present studies are mediated via

PKCs. As expected, pre-treatment of BEAS2B cells with BIM I (a pan PKC blocker) suppressed

PDBu-induced phosphorylation of PI3K p85, Akt, Src, ERK and JNK in a dose-dependent

manner (Figure 8-8A). Inhibitor for PI3K (LY294002) (Figure 8-8B) and Akt (Akt II) (Figure 8-

8C) attenuated not only the phosphorylation of p85 and Akt but also the phosphorylation of

SrcY416 and JNK. Src inhibitor (PP2) reduced phosphorylation of SrcY416, p85, ERK and JNK

(Figure 8-8D). Blocker for MEK (PD98059) and JNK (SP600125) prevented the PDBu-induced

phosphorylation of ERK1/2 and JNK, respectively (data not shown). Taken together, these data

165

suggest that PI3K/Akt pathway is upstream of Src, and Src may affect the phosphorylation

(activation of ERK1/2 and JNK).

A B Fig 8-7F-actin Overlay

U01

24U

0126

No Treatment PDBu

020406080

100

U0124 U0126 U0124 U0126

% Cell With Podosomes

0

20

40

60

80

100% Cell Degradating Matrix

C

F-actin MMP-9D E

U01

24U

0126

No Treatment PDBu

U0124 U0126 U0124 U0126

*

*

0

0.2

0.4

0.6

0.8

1


166

167

Figure 8-7. Confirmation of ERK pathway in PDBu-induced gelatin matrix degradation, MMP-9 translocation and gene expression with another MEK inhibitor.

(A) Cells cultured on gelatin matrix (Green) were pre-incubated with U0126 (inhibitor for MEK) or its analogue control U0124 (1 µM for 1 h), and then stimulated with PDBu(500 nM for 8 h). F-actin was stained with Rhodamine Phalloidin (Red). Digestion of gelatin matrix was revealed as black areas. (B) Quantitative analysis of percentage of cells showing podosomes. (C) Quantitative analysis of percentage of cells showing matrix degradation. (D) U0126 prevented MMP-9 translocation to PDBu-induced podosomes. F-actin was stained with Oregon Green 488 Phalloidin (Green). MMP-9 was revealed by Alexa Fluor 549 labeled secondary antibody (Red). (E) Quantitative analysis of percentage of MMP-9 overlapping F-actin staining by Pearson’s correlation coefficients. *: p < 0.05 in comparison with PDBu-stimulated control group.

168

No Treat

No

Trea

t

DM

SO

10µ

MB

IM 1

µMB

IM 5

µMB

IM 1

0µM

PDBu

No

Trea

tD

MS

O 1

0µM

BIM

1µM

BIM

5µM

BIM

10µ

M

a1

a2

a3

a5

a6

a7

No Treat

No

Trea

t

LY 1

µM

LY 5

µM

LY 1

0µM

No

Trea

t

LY 1

µM

LY 5

µM

LY 1

0µM

PDBu

b1

b2

b3

b5

b6

b7

(p)PI3K p85

(p)AktS473

(p)SrcY416

(p)JNK

(p)p38

GAPDH

No Treat

No

Trea

t

AktI

I 1µM

AktI

I 5µM

AktI

I 10µ

M

No

Trea

t

AktI

I 1µM

AktI

I 5µM

AktI

I 10µ

M

PDBu

c1

c2

c3

c5

c6

c7

A B

CNo Treat

No

Trea

t

PP

2 1µ

M

PP

2 5µ

M

PP

2 10

µM

No

Trea

t

PP

2 1µ

M

PP

2 5µ

M

PP

2 10

µM

PDBu

d1

d2

d3

d5

d6

d7

D

(p)PI3K p85

(p)AktS473

(p)SrcY416

(p)JNK

(p)p38

GAPDH

(p)ERKa4

(p)ERK

b4

c4 d4

Fig 8-8

Figure 8-8. Effects of multiple inhibitors on phosphorylation status of PI3K/Akt, Src and MAPKs.

Cells were pretreated with different doses of (A) BIM I (inhibitor for PKCs), (B) LY294002 (PI3K inhibitor), (C) Akt inhibitor II (Akt inhibitor) and (D) PP2 (Srcinhibitor) (1, 5, 10 µM for 1 h), and then stimulated with PDBu (500 nM, 30 min). Phosphorylation status of multiple proteins was examined with western blotting.

169

8.4 Discussion

In the present study, we demonstrated that in human bronchial epithelial cells, multiple

signal transduction pathways (PI3K/Akt, Src and MAPKs) are involved in PKC activation-

induced podosome formation and gelatin matrix degradation in a coordinated fashion. PI3K/Akt

and Src appear to be the upstream signals to participate in the regulation of podosome formation,

whereas ERK and JNK are the downstream signals to participate in the regulation of gelatin

matrix degradation. Interestingly, p38 MAPK was neither associated with PDBu-stimulated

podosome formation nor matrix degradation.

In the present study, we showed increased phosphorylation of SrcY416 and multiple

molecules in PI3K/Akt pathway. PI3K p85 , phosphorylated Akt, and c-Src were clustered at the

PDBu-induced F-actin rich small dots, rings, and belts in the cytoplasm and membrane ruffles.

Blocking PI3K or Akt activity with chemical inhibitors significantly prevented podosome

formation and proteolytic activity of PDBu-induced podosomes. These inhibitors also reduced

PDBu-induced phosphorylation of SrcY416, a sign of activation of c-Src (432). It has been

shown that PI3K activity was required for PMA-induced activation of c-Src through AFAP for

podosome formation and cell migration (201). Our results support that PI3K/Akt signaling

cascade and c-Src are involved in PKC activation-related podosome formation in normal human

bronchial epithelial cells as upstream event.

In contrast, our results also suggest that JNK and ERK1/2 are down-stream signaling

molecules, acting as direct effectors to regulate gelatin matrix degradation of podosomes in

human bronchial epithelial cells. Inhibitors for JNK and ERK pathways did not block podosome

assembly, but inhibited gelatin matrix degradation. This finding is very interesting, because

ERK1/2 has been implemented in PKC-mediated podosome formation in vascular smooth

170

muscle A7r5 cells (213, 214), and in primary aorta smooth muscle cells (430). In murine bone

marrow-derived dendritic cells (215) and bone marrow-derived neutrophils ERK1/2 is also

involved in cell migration and invasion, associated with podosome-like structures at the cell

leading edge (216). Big MAPK Erk5 promoted Src-induced podosome formation in fibroblasts

(217). We cannot explain why ERK1/2 and JNK are only involved in gelatin matrix degradation

in human bronchial epithelial cells. However, it has been recently shown that TGF- induced

formation of F-actin cores (podosomes) requires signaling through PI3K and Src, whereas TGF-

-induced matrix degradation requires ERK in human breast cancer cells (433). The formation

and proteolytic activity of podosomes should be considered as two highly related by distinct

cellular events. It should point out that our results could be the first report to indicate the

involvement of JNK in podosome related functions. Interestingly, phosphorylated ERK and JNK

were not significantly co-localized with podosomes but inhibition of these pathways reduced

translocation of MMP-9 to podosomes. The intermediate mechanisms of these cellular processes

merit further investigation.

In our recent (122, 423) and the present study, we found that MMP-9 is the most

dominant MMP isozyme in PDBu-induced gelatin matrix degradation. In human lung

adenocarcinoma A549 cells, EGF stimulated Src and JNK/ERK signaling pathway through FAK

at the cell membrane to increase MMP-9 expression and secretion (220). In human melanoma

cells, the homophilic interactions of CD151 (PETA-3/SFA-1), a member of the tetraspanins,

stimulated intergrin-dependent signaling to c-Jun through FAK-Src-MAPKs pathways to

enhance cell migration and MMP-9 expression (434). In human breast adenocarcinoma MCF-7

and MDA-MB-435 cells, CD99, a cell surface glycoprotein, enhanced cell migration and MMP-

9 expression through Akt-, ERK-, and JNK-dependent AP-1 activation (435). In the present

171

study, inhibitors for PI3K, Akt, Src, MEK and JNK also reduced MMP-9 gene expression. The

relationships of these pathways in podosome formation, matrix degradation and MMP-9 gene

expression should be further determined.

Taken together, our results suggested that PKC-activation induced podosome formation

and gelatin matrix degradation requires PI3K/Akt, Src and MAPKs in a coordinated fashion.

Since human bronchial epithelial cells are important for maintenance of airway epithelial

integrity and repair after tissue injury, these results may contribute to molecular mechanisms

related to these physiological and pathophysiological processes.

172

9. Chapter Nine. Summary, general discussion and perspectives

173

9. Chapter Nine. Summary, general discussion and perspective

9.1 Summary

Lung airway and bronchial epithelial cells are the major player in host defense and are

the primary target of lung diseases such as chronic obstructive pulmonary disease, asthma, cystic

fibrosis (364, 365). Epithelial cell migration plays an important role in the lung physiology, lung

repair after injury, and lung homeostasis. The serine/threonine PKC family is highly involved in

cell proliferation, differentiation, cytoskeleton reorganization, cell migration and invasion, and

the progression of tumorigenesis (292). Phorbol ester can induce dramatic cytoskeletal structure

changes in lung epithelial cells (378). I hypothesized that some of these structures are podosomes

in lung epithelial cells and may be involved in epithelial cell migration and invasion under

physiological and/or pathophysiological conditions. In my PhD thesis studies, I acquired primary

normal human bronchial epithelial cells and normal (non-cancerous) human lung bronchial

epithelial BEAS2B cells and challenged them with phorbol ester PDBu to activate PKCs. My

PhD thesis has three major original findings:

1. I discovered that phorbol ester may induce podosome formation in human normal (non-

cancerous) bronchial epithelial cells. I characterized these cellular structures and revealed the

potential function of these podosomes in normal bronchial epithelial cells

2. I demonstrated that PDBu-induced podosome assembly was mainly mediated through

redistribution of conventional PKCs, especially PKC , from the cytosol to the podosomes,

while atypical PKC locally played a dominant role in the regulation of proteolytic activity of

PDBu-induced podosomes through recruitment, release and activation of MMP-9 as

summarized in Figure 9-1

Fig

9-1

PD

Bu

Pro

-MM

P-2

Pro

-MM

P-9

Act

ive

MM

P-9

MM

P-1

4

PKC

Extra

cellu

larM

atrix

PKC

PKC

PKC

µ

Nuc

leus

MM

P-9

mR

NA

PD

Bu

Act

ive

PKC

(late

nt)

Activ

e PK

C

PKCPKCPK

Cµ

Gen

e Ex

pres

sion

Prot

ein

Synt

hesi

sSe

cret

ion

Act

ivat

ion

PD

Bu

cPKC

s(P

KC

)

Podo

som

eFo

rmat

ion

nPKC

sR

ecru

it PK

Cto

Po

doso

me

MM

P G

ene

Expr

essi

on

Pro

tein

Sy

nthe

sis

Secr

etio

n

Activ

atio

n

PKC

Reg

ulat

e

Podo

som

e

Form

atio

n

Podo

som

e

Func

tion

PKC

PKC

PKCPKCµ

PKC

Phos

phor

ylat

ion

MM

P-9

Rec

ruitm

ent

Oth

er P

KCs

??

174

Figu

re 9

-1. R

ole

of P

KC

sin

pod

osom

efo

rmat

ion

and

podo

som

alpr

oteo

lytic

func

tion.

PDBu

-indu

ced

podo

som

efo

rmat

ion

is m

ainl

y m

edia

ted

thro

ugh

clas

sica

l PKC

ses

peci

ally

PKC

.Th

e re

crui

tmen

t, se

cret

ion

and

activ

atio

n of

MM

P-9

and

pro

teol

ytic

activ

ity o

f pod

osom

esm

ainl

y de

pend

on

atyp

ical

PKC

spa

rticu

larly

PKC

. Nov

el P

KCis

invo

lved

in re

crui

tmen

t of P

KC

toth

e si

te o

f pod

osom

es.

175

3. I investigated that PI3K/Akt/Src were involved in regulating PDBu-induced podosome

formation as the upstream signals; whereas ERK1/2 and JNK MAPKs regulated the

proteolytic activity of PDBu-induced podosomes through recruitment of MMP-9 to

podosome and MMP-9 expression, secretion and activation.

These results will contribute to the understanding of the molecular mechanism of normal

epithelial cell migration and invasion in lung physiology and lung diseases.

9.2 Discussion

Several new findings were elucidated in my thesis. In this section, I will focus on the

topics never mentioned in previous chapters.

9.2.1 The role of podosomes in normal bronchial epithelial cells in lung physiology and

pathophysiology

Podosome is a newly discovered cellular structure that plays a critical role in cell

migration and invasion. Carman et al. found that lymphocytes used podosomes and extended

“invasive podosomes” to palpate the surface of, and ultimately form transcellular pores through,

the endothelium in diapedesis at ex vivo tissue slides by electronic microscopy. These

podosome-like structures were dependent on Src kinase and the actin regulatory protein WASP

(276). I speculate that similar cellular structures may exist when airway epithelial cells are trying

to across tissue boundaries. When epithelium is damaged, the surviving epithelial cells may

migrate towards the wounded area, or invade from the underneath to cover the denuded

epithelium as showed in Figure 1-2. Alternatively, when cells migrate over a two-dimensional

176

surface, matrix degradation of podosomes may also promote deadhesion of the cell. My work

that primary normal bronchial epithelial cells can form podosomes and membrane ruffles/waves

upon PKC activation, suggests that normal cells have the potential to gain an aggressive motility

under stimulation.

9.2.2 PKCs and cell motility in lung physiology and pathophysiology

PKCs are important in many cellular responses in the lung physiology and

pathophysiology. Dempsey et al. summarized the potential exogenous stimuli for PKC isozymes

in the lung, such as oxygen tension (hypoxia and hyperoxia), mechanical forces (pressure and

shear stress), inhaled irritants (cold air, oxidants and other noxious chemicals), inhaled dust

(asbestos, silica and beryllium) and certain inhaled allergens causing hypersensitivity

pneumonitis (436). These diverse agents are known to induce many different cellular responses

such as vessel contraction, increased vascular permeability, cell migration, cell invasion and cell

proliferation. In human lung cancer epithelial H460 and H1299 cells, nicotine induced the

activation of PKC and increased cell migration and cell invasion through phosphorylation of µ-

and m-Calpains (437). In human lung cancer NCI-H69 cells, tobacco nitrosamine 4-

(methylnitrosamino)-1-(3-pyridyl)-1-butanone (NNK) enhanced cell migration and invasion

through induction of the influx of Ca2+ and activation of PKC/Raf/MEK/ERK1/2 signaling

cascade through 7nAChR, which induced phosphorylation of both µ- and m-calpain leading to

calpain activation and secretion (438). In normal human lung bronchial epithelial BEAS2B cells,

hog barn dust slowed airway epithelial cell migration in a wound healing assay in vitro through a

PKC -dependent mechanism (439). Mechanisms that regulate PKC activity, including signaling

177

cascades, phosphorylation and interaction with isozyme-specific binding proteins, are potential

therapeutic targets for lung diseases.

9.2.3 PKC isoforms and isoform redundancy

For each PKC subfamily, I only focused on one representative isoform to study in my

thesis. Thus negative results in any one assay may simply refelect the fact that other isoforms of

the same class may compensate the deficiency. For example, Rodriguez et al found that /

isoform aPKC was the major contributor to podosome assembly and invasion, with playing a

subsidiary role in NIH 3T3 cells transformed by v-Src and by activated by c-Src (SrcY527F)

(440). Inhibition of both isoforms had a greater effect than inhibiting either one alone. In Chapter

8, I used a myristoylated pseudosubstrate inhibitor PS for PKC . The sequence of PS is Myr-

SIYRRGARRWRKL-OH. It not only targets human PKC , but also PKC . Therefore, there may

be an alternative interpretation of the data.

9.2.4 Cell migration vs. cell invasion

From my point of view, cell migration is a process that cells move along the surface,

whereas cell invasion is a situation that cells transmigrate to across different cell layers and/or

extracellular matrix. To explore the migration properties of BEAS2B cells bearing podosomes,

we performed a wound healing cell migration assay. In control group, cells formed a broad

lamellipodia at the leading edge toward the wound. In PDBu treated group, cells appeared

membrane ruffles all around. Cells were not polarized toward the wound and PDBu reduced cell

migration. Similarly, Slager et al. found that agricultural dust exposure activated PKC to slow

down BEAS2B cell migration during wound repair (439). In contrast, in Boyden chamber

178

invasion assay, BEAS2B cells with podosomes degraded the coated fibronectin gel in the

Transwell and invaded through the gel to the bottom side of the well. PDBu stimulation

enhanced the invasiveness of normal epithelial cells in vitro. I suspect that podosomes may have

similar function for cell motility in vivo. To prove this concept is difficult, however, with

advanced technologies and awareness of cell movement in 3D, it may be explored in the future.

9.2.5 Podosome formation vs. podosomal proteolytic function

With in situ zymography assay, I found that Rottlerin, PKC /µ/ siRNA, PKC

pseudosubstrate inhibitor, MMP-9/-14 siRNA, PD98059, U0126, SP600125 can inhibit matrix

degradation of PDBu-stimulated podosomes without affecting podosome assembly. Several

groups have also demonstrated that podosome assembly and podosomal matrix degradation can

be uncoupled and regulated by distinct signaling pathways. West et al. found that MMP-14

deficient mouse bone marrow cells and spleen dendritic cells produced podosomes but lost the

ability to degrade extracellular matrix after TLR signaling (419). In CA1D cells, a Ras

transformed invasive variant of the MCF10A human breast cell line, activation of PI3K and Src

kinases upon TGF- treatment is required for the formation of the actin core of podosomes,

whereas ERK activation through Smad2/3 signaling is for activating the protease MMP-9 and

degradation of extracellular matrix (433). In v-Src transformed NIH3T3 cells, Tks5 was required

for podosome formation, while Tks4 is required for MT1-MMP recruitment and extracellular

matrix degradation (431). Therefore, both data from us and other groups signify that podosome

assembly and proteolytic function are qualitative ly distinct, separately regulated processes.

These studies are important because understanding the biology of podosomes and regulation of

179

their proteolytic function will aid in identification of novel targets for anti-invasive or anti-

metastasis therapy.

In addition, my data could also be explained in an alternative way. The distinction

between podosome formation and proteolytic function is quantitative rather than qualitertive.

Accumulation of MMPs and matrix degradation increases as podosome assembly processes, but

the assays for MMP recruitment and matrix degradation are in general less sensitive than those

for detection of abundant cytoskeletal proteins such as actin or cortactin. Thus matrix

degradation can only be observed when podosome assembly and MMP recruitment has

processed beyond a certain threshold size. In this quantative model, weak inhibitors of podosome

formation could appear to specifically block MMPs recruitment and matrix degradation, because

the recruitment of cytoskeletal components such as actin or cortactin could still be detected when

podosome assembly is only partially inhibited. In Chapter 7 and 8, one interesting phenomenon

is we found that certain inhibitors of MMP recruitment and matrix degradation in thi system are

inhibitors of podosome formation in other systems. This might be due to cell type specific. It

could be also interpreted by this “quantitative model” mentioned above. Certain inhibitor may be

more or less effective in different cell types.

9.2.6 MMP-9 gene expression vs. MMP-9 recruitment

In BEAS2B cells, the expression and activity of MMP-9 are inducible by PDBu. MMP-9

is dominant in gelatin matrix degradation in our model. I found that PKC pseudosubstrate

inhibitor attenuated MMP-9 enzymatic activity and translocation without interrupting the mRNA

level, but PD98059 and SP600125 inhibited mRNA expression, as well as translocation and

enzymatic activity of MMP-9. It indicated that PKC might locally regulate MMP-9 secretion

180

and activity, and ERK1/2 and JNK signaling can manipulate both MMP-9 gene expression and

activity.

The activity of MMPs is generally regulated by several steps, gene transcription (249);

mRNA stability (441); secretion and activation of the latent pro-enzymes (442-445); and

inhibition by their endogenous inhibitors, the tissue inhibitors of matrix metalloproteinases

(TIMPs), which are able to inhibit metalloproteinase activity by forming non-covalent complexes

with active MMPs (8, 226). Recently, it is found that MMP intracellular trafficking is an

indispensable step for MMP regulation (446). In N2a neuroblastoma and primary neuronal cells,

MMP-2 and MMP-9 and their inhibitor TIMP-1 were secreted in 160–200 nm vesicles in a

Golgi-dependent pathway. These vesicles distributed along microtubules and microfilaments, co-

localized differentially with the molecular motors kinesin and myosin Va and undergo both

anterograde and retrograde trafficking. MMP-9 retrograde transport involved in the

dynein/dynactin molecular motor (446). In hippocampal neurons, MMP-2 and MMP-9 vesicles

were preferentially distributed in the somato-dendritic compartment and were found in dendritic

spines. Non-transfected hippocampal neurons also demonstrated vesicular secretion of MMP-2

in both its pro- and active forms and gelatinolytic activity localized within dendritic spines (446).

MAPK pathway signaling molecules are highly involved in both direct and indirect

regulation of MMP-9, through gene expression and protein-protein interaction through integrin

1 as shown in Figure 9-2. Accumulated data indicated that JNK can regulate MMP-9. In mouse

fibroblast cells, JNK1 increased the expression of MMP-9 protein (447). In human osteosarcoma

U-2 OS cells, JNK1 is necessary for expression of MMP-9 mRNA (448). I found that in addition

to decrease MMP-9 mRNA level, JNK pan inhibitor SP600125 also reduced MMP-9 protein

expression and activation and secretion in BEAS2B cells.

Fig 9-2

181

Figure 9-2. MAPK signaling pathways regulate MMP-9.

Network of how MAPK signaling pathways regulate MMP-9 generated by Ingenuity Pathway Analysis software.

182

Increasing evidence showed that MAPKs can regulate MMP-9 through integrin 1.

Thrombin is a known procoagulatory serine protease to induced invasion and metastasis in

various cancers. Thrombin-induced invasion of human osteosarcoma US-OS cells through

Matrigel was mediated by PI3K and ERK signaling pathways through the induction and

association of MMP-9 and integrin 1 on the cell surface (449). Accumulation and depletion of

GM3 gangliosides in epithelial cells cultured on fibronectin correlated inversely with MMP-9

expression and activation, EGFR and MAPK phosphorylation and Jun expression (450).

Ganglioside depletion facilitated the interaction of MMP-9 and integrin 5 1 (450), suggesting

that dissociation of integrin and MMP-9 as the potential mechanisms for the GM3-induced

effects on MMP-9 function and cell migration (450).

3 1 integrin played a critical role in MMP-9 gene expression, activation and production.

In immortalized mouse keratinocytes, 3 1 integrin not only regulated MMP-9 mRNA stability

in response to activation of MEK/ERK pathway (441), but also was directly essential for MMP-

9 secretion (442). In human ovarian cancer cell line MDAH 2774, 3 1 integrin mediated MMP-

9 activity through transcriptional regulation of its endogenous inhibitor TIMP-1 via c-fos binding

to the AP-1 up region of the TIMP-1 promoter (451). We also found that 1 integrin was well

colocalized with PDBu-induced podosomes in BEAS2B cells. It would be interesting to

investigate whether integrin is involved in MMP-9 expression and production through interaction

with PKCs.

9.2.7 Study approaches and their limitations

9.2.7.1 Cell lines and primary cells

183

In my PhD thesis work, I used primary normal human bronchial epithelial cells and

normal human bronchial epithelial cell line BEAS2B. Different from primary cells that die in a

few weeks in culture, cell lines are modified cells, with the immortal growing property. The

modification could cause changes that may affect the nature of the original cells. In my thesis,

although most studies were using cell lines, I did confirm my results with primary cells. The

primary cells even form better podosomes than cell lines. I also confirmed that PKC play a

dominant role in podosome assembly and the proteolytic function in primary cells by multiple

approaches. Due to the limited availability and the difficulties to transfect the primary cells, cell

lines are still important tools for biochemical studies.

9.2.7.2 Chemical inhibitor and siRNA

I used variety of chemical inhibitors. One potential limitation for pharmaceutical inhibitor

is the specificity of these chemicals. I also used specific siRNA to knockdown targeted proteins

in my work. siRNA is more specific than chemical inhibitor. However, the potential off-target

effects should be kept in mind. Combining multiple methods is necessary for conclusive studies.

9.2.7.3 In vitro vs. in vivo

I mainly used in vitro cell culture model to study the role of epithelial cell migration and

invasion with the advanced cell culture and microscopy techniques. There are various limitations

of employing cell culture to study the molecular mechanism of epithelial cell migration and

invasion in lung repair and regeneration (74). Now, armed with the new technology available

such as intravital confocal microscopy, two-photon and multi-photon microscopy, real-time

confocal microscopy and so on (452, 453), it is time to return to in vivo model once again. For

184

example, we could generate rat lung epithelial injury model by naphthalene (81) and study the

process of surviving epithelial cell migration and invasion to recover the denude area by

immunofluorescent staining. To get high resolution of bronchus epithelium, we could also use

GFP-mouse and the “stick objective”, a new product from Olympus (453), to investigate the

process of surviving epithelial cell migration and invasion to the injured area by live cell

imaging.

9.3 Conclusions

In my PhD thesis, studies provide evidence for the molecular mechanism of podosome

formation and their proteolytic function in normal human bronchial epithelial cells. PKC, PI3K,

Akt, Src and MAPKs play a coordinated role in phorbol ester-induced podosome formation and

their proteolytic function as summarized in Figure 9-3. The upstream signaling pathways

classical PKC , PI3K, Akt and Src control podosome formation. Novel PKC facilitated the

translocation of atypical PKC and MMP-9 to the sites of podosomes. Atypical PKC , ERK and

JNK work as the downstream signals to locally control podosomal proteolytic function. All these

findings may be important for normal epithelial cell migration and invasion during physiological

or pathophysiological conditions

Fig 9-3

cPKC

nPKC

aPKC

PI3K/Akt/Src

ERK/JNK

Podosomeassembly

Proteolyticactivity

PDBu

Cell migration

and

Cell invasion

Figure 9-3. PKC, PI3K, Akt, Src and MAPK in podosome formation and podosomal proteolytic function.

Signaling cascades PKC, PI3K, Akt, Src, and MAPK especially ERK and JNK, play a central role in podosome formation and the podosomal proteolytic function. The upstream signaling pathways classical PKCs, PI3K, Akt and Src control podosomeformation. Novel PKC facilitated the translocation of atypical PKCs to the sites of podosomes. Atypical PKC , ERK, and JNK work as the downstream signals to locally control podosomal proteolytic function. These may be important for normal bronchial epithelial cell migration and invasion during physiological and pathophysiologicalconditions.

185

186

9.4 Future directions

In my PhD thesis, I have used a cellular model of normal human bronchial epithelial cells

to study the molecular mechanism of podosome formation and podosomal proteolytic function in

vitro. To further continue my work, we design the following potential research topics to carry on.

These future studies will aid in elucidation of the mechanism of cell migration and invasion in

physiological and pathophysiological conditions.

o To identify the potential interaction between PKC and and MMP-9, we may perform

immunoprecipatation and co-immunoprecipitation. This will reveal whether PKC , and

MMP-9 can bind to each other directly or indirectly.

o To further investigate the signaling cascade of PKCs, PI3K, Src, MAPKs and MMP-9, we

need to identify the binding partners of each molecule, upstream activators and downstream

substrates.

o To examine whether PKC need kinase activity to activate MMP-9 and facilitate MMP-9

secretion, we need to make dominant negative and dominant active constructs to test our

idea.

o To investigate whether these cellular structures- podosome, invadopodia, and membrane

ruffles- really exist in vivo, we may optimize the staining protocol to stain podosomal

markers, cytoskeleton proteins and MMPd, using frozen section of human lung bronchus

tissues. We can also use electronic microscope to visualize the structure of lung bronchus

epithelium tissue sample to identify the membrane invaginations and protrusions of cells

across tissue boundaries.

187

o To determine the role of PKCs in podosomes and cell migration and invasion in vivo, we can

stain the native and phosphorylated species of PKCs especially and on the above

mentioned tissue slides to reveal whether they are co-localized with podosomes.

o To investigate the role of PKCs in lung repair and regeneration after injury, we may inject

PKC chemical inhibitors (BIM I, Gö6976, Rottlerin, PKC pseudosubstrate inhibitor) or

deliver PKC siRNA to the lungs in the rat naphthalene epithelium regeneration model (81).

We can test whether these PKC blockers or PKC siRNA prevent the surviving epithelium

repair after denudation by naphthalene.

o Similarly, we can also use PI3K, Src and MAPKs inhibitors to test the role of these

molecules in epithelial repair and regeneration after injury in the naphthalene model.

188

10. References

189

References

1. Vignola, A.M., La Grutta, S., Chiappara, G., Benkeder, A., Bellia, V., and Bonsignore, G. 2002. Cellular network in airways inflammation and remodelling. Paediatr Respir Rev3: 41-46.

2. Selman, M., and Pardo, A. 2006. Role of epithelial cells in idiopathic pulmonary fibrosis: from innocent targets to serial killers. Proc Am Thorac Soc 3: 364-372.

3. Zabner, J., Seiler, M.P., Launspach, J.L., Karp, P.H., Kearney, W.R., Look, D.C., Smith, J.J., and Welsh, M.J. 2000. The osmolyte xylitol reduces the salt concentration of airway surface liquid and may enhance bacterial killing. Proc Natl Acad Sci U S A 97: 11614-11619.

4. Ivanov, A.I., Nusrat, A., and Parkos, C.A. 2004. The epithelium in inflammatory bowel disease: potential role of endocytosis of junctional proteins in barrier disruption. Novartis Found Symp 263: 115-124; 124-132, 211-118.

5. He, X., Han, B., Mura, M., Li, L., Cypel, M., Soderman, A., Picha, K., Yang, J., and Liu, M. 2008. Anti-human tissue factor antibody ameliorated intestinal ischemia reperfusion-induced acute lung injury in human tissue factor knock-in mice. PLoS ONE 3: e1527.

6. Adamson, I.Y., Bowden, D.H. 1974. The type 2 cell as progenitor of alveolar epithelial regeneration. A cytodynamic study in mice after exposure to oxygen. Lab Invest 30: 35-42.

7. Keenan, K.P., Combs, J.W., McDowell, E.M. 1982. Regeneration of hamster tracheal epithelium after mechanical injury. I. Focal lesions: Quantitative morphologic study of cell proliferation. Virchows Arch [Cell Pathol] 41: 193-214.

8. Coraux, C., Martinella-Catusse, C., Nawrocki-Raby, B., Hajj, R., Burlet, H., Escotte, S., Laplace, V., Birembaut, P., Puchelle, E. 2005. Differential expression of matrix metalloproteinases and interleukin-8 during regeneration of human airway epithelium in vivo. J Pathol 206: 160-169.

9. Zahm, J.m., Chevillard, M., Puchelle, E. 1991. Wound repair of human respiratory epithelium. Am J Respir Cell Mol Biol 5: 242-248.

10. Zahm, J.M., Pierrot, D., Chevillard, M., Puchelle, E. 1992. Dynamics of cell movement during the wound repair of human surface respiratory epithelium. Biorheology 29: 459-465.

11. Zahm, J.M., Kaplan, H., Herard, A.L., Doriot, F., Pierrot, D., Somelette, P., et al. 1997. Cell migration and proliferation during the in vitro wound repair of the respiratory epithelium. Cell Motil Cytoskeleton 37: 33-43.

12. Erjefalt, J.S., Erjefalt, I., Sundler, F., Persson, C.G.A. 1995. In vivo restitution of airway epithelium. Cell Tissue Res 281: 305-316.

190

13. Wilhelm, D.L. 1953. Regeneration of tracheal epithelium. J Pathol Bacteriol 5025: 543-550.

14. Erjefalt, J.S., Sundler, F., Persson, C.G.A. 1997. Airway basal cells may promptly assume a barrier structure. Thorax 52: 213-217.

15. Zahm, J.M., Kaplan, H., Herard, A.L., Doriot, F., Pierrot, D., Somelette, P., et al. 1997. Cell migration and proliferation during the in vitro wound repair of the respiratory epithelium. Cell Motil Cytoskeleton 37: 33-43.

16. Erjefalt, J.S., Erjefalt, I., Sundler, F., and Persson, C.G. 1995. In vivo restitution of airway epithelium. Cell Tissue Res 281: 305-316.

17. Shimizu, T., Nishihara, M., Kawaguchi, S., Sakakura, Y. 1994. Expression of phenotypic markers during regeneration of rat tracheal epithelium following mechanical injury. Am J Respir Cell Mol Biol 11: 85-94.

18. Puchelle, E., Zahm, J.M., Tournier, J.M., and Coraux, C. 2006. Airway epithelial repair, regeneration, and remodeling after injury in chronic obstructive pulmonary disease. Proc Am Thorac Soc 3: 726-733.

19. Goldman, D.L., Davis, J., Bommarito, F., Shao, X., and Casadevall, A. 2006. Enhanced allergic inflammation and airway responsiveness in rats with chronic Cryptococcus neoformans infection: potential role for fungal pulmonary infection in the pathogenesis of asthma. J Infect Dis 193: 1178-1186.

20. Spurzem, J.R., Raz, M., Ito, H., Kelling, C., Stine, L.C., Romberger, D.J., Rennard, S.I. 1995. Bovine herpesvirus-1 infection reduces bronchial epithelial cell migration to extracellular matrix proteins. Am J Physiol Lung Cell Mol Physiol 268: L214-L220.

21. Wesley, U.V., Bove, P.F., Hristova, M., McCarthy, S., and van der Vliet, A. 2007. Airway epithelial cell migration and wound repair by ATP-mediated activation of dual oxidase 1. J Biol Chem 282: 3213-3220.

22. Dosanjh, A., and Zuraw, B. 2003. Endothelin-1 (ET-1) decreases human bronchial epithelial cell migration and proliferation: implications for airway remodeling in asthma. J Asthma 40: 883-886.

23. Bove, P.F., Hristova, M., Wesley, U.V., Olson, N., Lounsbury, K.M., and van der Vliet, A. 2008. Inflammatory levels of nitric oxide inhibit airway epithelial cell migration by inhibition of the kinase ERK1/2 and activation of hypoxia-inducible factor-1 alpha. J Biol Chem 283: 17919-17928.

24. Ahdieh, M., Vandenbos, T., and Youakim, A. 2001. Lung epithelial barrier function and wound healing are decreased by IL-4 and IL-13 and enhanced by IFN-gamma. Am J Physiol Cell Physiol 281: C2029-2038.

191

25. Cantral, D.E., Sisson, J.H., Veys, T., Rennard, S.I., Spurzem, J.R. 1995. Effects of cigarette smoke extract on bovine bronchial epithelial cell attachment and migration. AmJ Physiol 268: L723-L728.

26. Wang, H., Liu, X., Umino, T., Skold, C.M., Zhu, Y., Kohyama, T., et al. 2001. Cigarette smoke inhibits human bronchial epithelial cell repair processes. Am J Respir Cell Mol Biol 25: 772-779.

27. Slager, R.E., Allen-Gipson, D.S., Sammut, A., Heires, A., DeVasure, J., Von Essen, S., et al. 2007. Hog barn dust slows airway epithelial cell migration in vitro through a PKCalpha-dependent mechanism. Am J Physiol Lung Cell Mol Physiol 293: L1469-L1474.

28. Wyatt, T.A., Slager, R.E., Devasure, J., Auvermann, B.W., Mulhern, M.L., Von Essen, S., Mathisen, T., Floreani, A.A., and Romberger, D.J. 2007. Feedlot dust stimulation of interleukin-6 and -8 requires protein kinase Cepsilon in human bronchial epithelial cells. Am J Physiol Lung Cell Mol Physiol 293: L1163-1170.

29. Olsen, C.E., Liguori, A.E., Zong, Y., Lantz, R.C., Burgess, J.L., and Boitano, S. 2008. Arsenic upregulates MMP-9 and inhibits wound repair in human airway epithelial cells. Am J Physiol Lung Cell Mol Physiol 295: L293-302.

30. Ahmad, S., Ahmad, A., McConville, G., Schneider, B.K., Allen, C.B., Manzer, R., Mason, R.J., and White, C.W. 2005. Lung epithelial cells release ATP during ozone exposure: signaling for cell survival. Free Radic Biol Med 39: 213-226.

31. Suadicani, P., Hein, H.O., Meyer, H.W., and Gyntelberg, F. 2001. Exposure to cold and draught, alcohol consumption, and the NS-phenotype are associated with chronic bronchitis: an epidemiological investigation of 3387 men aged 53-75 years: the Copenhagen Male Study. Occup Environ Med 58:160-164.

32. Elder, A., and Oberdorster, G. 2006. Translocation and effects of ultrafine particles outside of the lung. Clin Occup Environ Med 5: 785-796.

33. Shukla, A., Timblin, C., BeruBe, K., Gordon, T., McKinney, W., Driscoll, K., Vacek, P., and Mossman, B.T. 2000. Inhaled particulate matter causes expression of nuclear factor (NF)-kappaB-related genes and oxidant-dependent NF-kappaB activation in vitro. Am J Respir Cell Mol Biol 23:182-187.

34. Savla, U., Waters, C.M. 1998. Mechanical strain inhibits repair of airway epithelium in vitro. Am J Physiol 274: L883-892.

35. Spurzem, J.R., Veys, T., Devasure, J., Sisson, J.H., and Wyatt, T.A. 2005. Ethanol treatment reduces bovine bronchial epithelial cell migration. Alcohol Clin Exp Res 29: 485-492.

192

36. Wyatt, T.A., Kharbanda, K.K., Tuma, D.J., Sisson, J.H., and Spurzem, J.R. 2005. Malondialdehyde-acetaldehyde adducts decrease bronchial epithelial wound repair. Alcohol 36: 31-40.

37. Ito, H., Rennard, S.I., and Spurzem, J.R. 1996. Mononuclear cell conditioned medium enhances bronchial epithelial cell migration but inhibits attachment to fibronectin. J Lab Clin Med 127: 494-503.

38. Wyatt, T.A., Ito, H., Veys, T.J., and Spurzem, J.R. 1997. Stimulation of protein kinase C activity by tumor necrosis factor-alpha in bovine bronchial epithelial cells. Am J Physiol273: L1007-1012.

39. Kida, H., Mucenski, M.L., Thitoff, A.R., Le Cras, T.D., Park, K.S., Ikegami, M., Muller, W., and Whitsett, J.A. 2008. GP130-STAT3 regulates epithelial cell migration and is required for repair of the bronchiolar epithelium. Am J Pathol 172: 1542-1554.

40. Shahabuddin, S., Ji, R., Wang, P., Brailoiu, E., Dun, N., Yang, Y., Aksoy, M.O., and Kelsen, S.G. 2006. CXCR3 chemokine receptor-induced chemotaxis in human airway epithelial cells: role of p38 MAPK and PI3K signaling pathways. Am J Physiol Cell Physiol 291: C34-39.

41. Kim, J.S., Rabe, K.F., Magnussen, H., Green, J.M., and White, S.R. 1995. Migration and proliferation of guinea pig and human airway epithelial cells in response to tachykinins. Am J Physiol 269: L119-126.

42. Oertel, M., Graness, A., Thim, L., Buhling, F., Kalbacher, H., and Hoffmann, W. 2001. Trefoil factor family-peptides promote migration of human bronchial epithelial cells: synergistic effect with epidermal growth factor. Am J Respir Cell Mol Biol 25: 418-424.

43. Graness, A., Chwieralski, C.E., Reinhold, D., Thim, L., and Hoffmann, W. 2002. Protein kinase C and ERK activation are required for TFF-peptide-stimulated bronchial epithelial cell migration and tumor necrosis factor-alpha-induced interleukin-6 (IL-6) and IL-8 secretion. J Biol Chem 277: 18440-18446.

44. Bove, P.F., and van der Vliet, A. 2006. Nitric oxide and reactive nitrogen species in airway epithelial signaling and inflammation. Free Radic Biol Med 41: 515-527.

45. Bove, P.F., Wesley, U.V., Greul, A.K., Hristova, M., Dostmann, W.R., and van der Vliet, A. 2007. Nitric oxide promotes airway epithelial wound repair through enhanced activation of MMP-9. Am J Respir Cell Mol Biol 36: 138-146.

46. Shoji, S., Ertl, R.F., Linder, J., Koizumi, S., Duckworth, W.C., and Rennard, S.I. 1990. Bronchial epithelial cells respond to insulin and insulin-like growth factor-I as a chemoattractant. Am J Respir Cell Mol Biol 2: 553-557.

47. Keshamouni, V.G., Michailidis, G., Grasso, C.S., Anthwal, S., Strahler, J.R., Walker, A., Arenberg, D.A., Reddy, R.C., Akulapalli, S., Thannickal, V.J., et al. 2006. Differential protein expression profiling by iTRAQ-2DLC-MS/MS of lung cancer cells undergoing

193

epithelial-mesenchymal transition reveals a migratory/invasive phenotype. J Proteome Res 5: 1143-1154.

48. Spurzem, J.R., Gupta, J., Veys, T., Kneifl, K.R., Rennard, S.I., and Wyatt, T.A. 2002. Activation of protein kinase A accelerates bovine bronchial epithelial cell migration. AmJ Physiol Lung Cell Mol Physiol 282: L1108-1116.

49. Salathe, M. 2002. Effects of beta-agonists on airway epithelial cells. J Allergy Clin Immunol 110: S275-281.

50. Allen-Gipson, D.S., Wong, J., Spurzem, J.R., Sisson, J.H., and Wyatt, T.A. 2006. Adenosine A2A receptors promote adenosine-stimulated wound healing in bronchial epithelial cells. Am J Physiol Lung Cell Mol Physiol 290: L849-855.

51. Kuhn, C., 3rd, Boldt, J., King, T.E., Jr., Crouch, E., Vartio, T., and McDonald, J.A. 1989. An immunohistochemical study of architectural remodeling and connective tissue synthesis in pulmonary fibrosis. Am Rev Respir Dis 140: 1693-1703.

52. Herard, A.L., Pierrot, D., Hinnrasky, J., Kaplan, H., Sheppard, D., Puchelle, E., et al. 1996. Fibronectin and its 5 1 integrin receptor are involved in the wound repair process of the airway epithelium. Am J Physiol 271: L726-L733.

53. Rickard, K.A., Taylor, J., Spurzem, J.R., and Rennard, S.I. 1992. Extracellular matrix and bronchial epithelial cell migration. Chest 101: 17S-18S.

54. White, S.R., Dorscheid, D.R., Rabe, K.F., Wojcik, K.R., and Hamann, K.J. 1999. Role of very late adhesion integrins in mediating repair of human airway epithelial cell monolayers after mechanical injury. Am J Respir Cell Mol Biol 20: 787-796.

55. Legrand, C., Gilles, C., Zahm, J.M., Polette, M., Buisson, A.C., Kaplan, H., et al. 1999. Airway epithelial cell migration dynamics. MMP-9 role in cell-extracellular matrix remodeling. J Cell Biol 146: 517-529.

56. Zhu, M., Tian, D., Li, J., Ma, Y., Wang, Y., and Wu, R. 2007. Glycogen synthase kinase 3beta and beta-catenin are involved in the injury and repair of bronchial epithelial cells induced by scratching. Exp Mol Pathol 83: 30-38.

57. Lesur, O., Arsalane, K., Lane, D. 1996. Lung alveolar epithelial cell migration in vitro: modulators and regulation processes. Am J Physiol Lung Cell Mol Physiol 270: L311-L319.

58. Kheradmand, F., Folkesson, H.G., Shum, L., Derynk, R., Pytela, R., Matthay, M.A. 1994. Transforming growth factor-alpha enhances alveolar epithelial cell repair in a new in vitro model. Am J Physiol Lung Cell Mol Physiol 267: L728-L738.

59. Kim, H.J., Henke, C.A., Savik, S.K., Ingbar, D.H. 1997. Integrin mediation of alveolar epithelial cell migration on fibronectin and type I collagen. Am J Physiol Lung Cell Mol Physiol 273: L134-L141.

194

60. Clark, R.A., Mason, R.J., Folkvord, J.M., and McDonald, J.A. 1986. Fibronectin mediates adherence of rat alveolar type II epithelial cells via the fibroblastic cell-attachment domain. J Clin Invest 77: 1831-1840.

61. Crouch, E. 1990. Pathobiology of pulmonary fibrosis. Am J Physiol 259: L159-184.

62. Torikata, C., Villiger, B., Kuhn, C., 3rd, and McDonald, J.A. 1985. Ultrastructural distribution of fibronectin in normal and fibrotic human lung. Lab Invest 52: 399-408.

63. Madtes, D.K., Busby, H.K., Strandjord, T.P., and Clark, J.G. 1994. Expression of transforming growth factor-alpha and epidermal growth factor receptor is increased following bleomycin-induced lung injury in rats. Am J Respir Cell Mol Biol 11: 540-551.

64. Yu, H., Konigshoff, M., Jayachandran, A., Handley, D., Seeger, W., Kaminski, N., and Eickelberg, O. 2008. Transgelin is a direct target of TGF-beta/Smad3-dependent epithelial cell migration in lung fibrosis. FASEB J 22: 1778-1789.

65. Nagahori, T., Dohi, M., Matsumoto, K., Saitoh, K., Honda, Z.I., Nakamura, T., and Yamamoto, K. 1999. Interferon-gamma upregulates the c-Met/hepatocyte growth factor receptor expression in alveolar epithelial cells. Am J Respir Cell Mol Biol 21: 490-497.

66. Zou, Y., Lim, S., Lee, K., Deng, X., and Friedman, E. 2003. Serine/threonine kinase Mirk/Dyrk1B is an inhibitor of epithelial cell migration and is negatively regulated by the Met adaptor Ran-binding protein M. J Biol Chem 278: 49573-49581.

67. Galiacy, S., Planus, E., Lepetit, H., Fereol, S., Laurent, V., Ware, L., Isabey, D., Matthay, M., Harf, A., and d'Ortho, M.P. 2003. Keratinocyte growth factor promotes cell motility during alveolar epithelial repair in vitro. Exp Cell Res 283: 215-229.

68. Geiser, T., Jarreau, P.H., Atabai, K., and Matthay, M.A. 2000. Interleukin-1beta augments in vitro alveolar epithelial repair. Am J Physiol Lung Cell Mol Physiol 279: L1184-1190.

69. Lesur, O., Brisebois, M., Thibodeau, A., Chagnon, F., Lane, D., Fullop, T. 2004. Role if IFN-gamma and IL-2 in rat lung epithelial cell migration and apoptosis after oxidant injury. Am J Physiol Lung Cell Mol Physiol 286: L4-L14.

70. Planus, E., Galiacy, S., Matthay, M., Laurent, V., Gavrilovic, J., Murphy, G., Clerici, C., Isabey, D., Lafuma, C., and d'Ortho, M.P. 1999. Role of collagenase in mediating in vitro alveolar epithelial wound repair. J Cell Sci 112 ( Pt 2): 243-252.

71. Pardo, A., Gibson, K., Cisneros, J., Richards, T.J., Yang, Y., Becerril, C., Yousem, S., Herrera, I., Ruiz, V., Selman, M., et al. 2005. Up-regulation and profibrotic role of osteopontin in human idiopathic pulmonary fibrosis. PLoS Med 2: e251.

72. Lazar, M.H., Christensen, P.J., Du, M., Yu, B., Subbotina, N.M., Hanson, K.E., Hansen, J.M., White, E.S., Simon, R.H., and Sisson, T.H. 2004. Plasminogen activator inhibitor-1

195

impairs alveolar epithelial repair by binding to vitronectin. Am J Respir Cell Mol Biol 31: 672-678.

73. Barlow, P.G., Brown, D.M., Donaldson, K., MacCallum, J., and Stone, V. 2008. Reduced alveolar macrophage migration induced by acute ambient particle (PM10) exposure. Cell Biol Toxicol 24: 243-252.

74. Rickard, K.A., Shoji, S., Spurzem, J.R., Rennard, S.I. 1991. Attachment characteristics of bovine bronchial epithelial cells to extracellular matrix components. Am J Respir Cell Mol Biol 4: 440-448.

75. Erjefalt, J.S., Persson, C.G.A. 1997. Airway epithelial repair: breathtakingly quick and multipotentially pathogenic. Thorax 52: 1010-1012.

76. Yamada, M., Kubo, H., Kobayashi, S., et al. 2004. Bone marrow-derived progenitor cells are important for lung repair after lipopolysaccharide-induced lung injury. J Immunol172: 1266-1272.

77. Loebinger, M.R., Janes, S.M. 2007. Stem cells for lung disease. Chest 132: 279-285.

78. Kotton, D.N., Ma, B.Y., Cardoso, W.V. et al. 2001. Bone marrow-derived cells as progenitors of lung alveolar epithelium. Development 128: 5181-5188.

79. Loi, R., Beckett, T., Goncz, K.K., et al. 2006. Limited restoration of cystic fibrosis lung epithelium in vivo with adult bone marrow derived cells. Am J Respir Crit Care Med.173: 171-179.

80. Gomperts, B.N., Belperio, J.A., Rao, P.N., Randell, S.H., Fishbein, M.C., Burdick, M.D., et al. 2006. Circulating progenitor epithelial cells traffic via CXCR4/CXCL12 in response to airway injury. J Immunol 176: 1916-1927.

81. Wong, A.P., Dutly, A.E., Sacher, A., Lee, H., Hwang, D.M., Liu, M., Keshavjee, S., Hu, J., and Waddell, T.K. 2007. Targeted cell replacement with bone marrow cells for airway epithelial regeneration. Am J Physiol Lung Cell Mol Physiol 293: L740-752.

82. Kim, C.F., Jackson, E.L., Woolfenden, A.E., Lawrence, S., Babar, I., Vogel, S., Crowley, D., Bronson, R.T., and Jacks, T. 2005. Identification of bronchioalveolar stem cells in normal lung and lung cancer. Cell 121: 823-835.

83. Kim, C.F. 2007. Paving the road for lung stem cell biology: bronchioalveolar stem cells and other putative distal lung stem cells. Am J Physiol Lung Cell Mol Physiol 293: L1092-1098.

84. Roman, J.R., MacDonald, J.A. 1991. Cellular processes in lung repair. Chest 100: 245-248.

85. Horwitz, A.R., Parsons, J.T. 1999. Cell migration- movin' on. Science 286: 1102-1103.

196

86. Lauffenburger, D.A., Horwitz, A.F. 1996. Cell migration: a physically integrated molecular process. Cell 84: 359-369.

87. Legrand, C., Gilles, C., Zahm, J.M., Polette, M., Buisson, A.C., Kaplan, H., Birembaut, P., and Tournier, J.M. 1999. Airway epithelial cell migration dynamics. MMP-9 role in cell-extracellular matrix remodeling. J Cell Biol 146: 517-529.

88. Kaverina, I., Krylyshkina, O., Small, J.V. 1999. Microtubule targeting of substrate contacts promotes their relaxation and dissociation. J Cell Biol 146: 1033-1044.

89. Hall, A. 1998. Rho GTPases and the actin cytoskeleton. Science 279: 509-514.

90. Waterman-Storer, C.M., Salmon, E. 1999. Positive feedback interactions between microtubule and actin dynamics during cell motility. Curr Opin Cell Biol 11: 61-67.

91. Vicente-Manzanares, M., Koach, M.A., Whitmore, L., Lamers, M.L., and Horwitz, A.F. 2008. Segregation and activation of myosin IIB creates a rear in migrating cells. J Cell Biol 183:543-554.

92. Coniglio, S.J., Zavarella, S., and Symons, M.H. 2008. Pak1 and Pak2 mediate tumor cell invasion through distinct signaling mechanisms. Mol Cell Biol 28: 4162-4172.

93. Yin, J., and Yu, F.S. 2008. Rho kinases regulate corneal epithelial wound healing. Am J Physiol Cell Physiol 295: C378-387.

94. Pinner, S., and Sahai, E. 2008. PDK1 regulates cancer cell motility by antagonising inhibition of ROCK1 by RhoE. Nat Cell Biol 10: 127-137.

95. Lo, S.H., Weisberg, E., Chen, L.B. 1994. Tensin: a potential link between the cytoskeleton and signal transduction. Bioassay 16: 817-823.

96. Schaller, M., Parsons, J.T. . 1994. Focal adhesion kinase and associated proteins. CurrOpin Cell Biol 6: 705-710.

97. Turner, C.E. 1994. Paxillin: a cytoskeletal target for tyrosine kinases. Bioassay 16:47-52.

98. Wu, D.Y., Goldberg, D.J. 1993. Regulated tyrosine phosphorylation at the tips of growth cone filopodia. . J Cell Biol 123: 653-664.

99. Klinghoffer, R.A., Sachsenmaier, C., Cooper, J.A., Soriano, P. 1999. Src family kinases are required for integrin but not PDGFR signal transduction. EMBO J 18: 2459-2471.

100. Sieg, D.J., Hauck, C.R., Schlaepfer, D.D. 1999. Required role of focal adhesion kinase (FAK) for integrin-stimulated cell migration. J Cell Sci 112: 2677-2691.

101. Huttenlocher, A., Palecek, S.P., Lu, Q., Zhang, W., Mellgren, R.L., Lauffenburger, D.A., Ginsberg, M.H., Horwitz, A.F. 1997. Regulation of cell migration by the calcium-dependent protease calpain. J Biol Chem 272: 32719-32722.

197

102. Franke, T.F., Kaplan, D.R., and Cantley, L.C. 1997. PI3K: downstream AKTion blocks apoptosis. Cell 88: 435-437.

103. Ling, K., Schill, N.J., Wagoner, M.P., Sun, Y., and Anderson, R.A. 2006. Movin' on up: the role of PtdIns(4,5)P(2) in cell migration. Trends Cell Biol 16: 276-284.

104. Janetopoulos, C., and Devreotes, P. 2006. Phosphoinositide signaling plays a key role in cytokinesis. J Cell Biol 174: 485-490.

105. Insall, R.H., and Weiner, O.D. 2001. PIP3, PIP2, and cell movement--similar messages, different meanings? Dev Cell 1: 743-747.

106. Insall, R.H., and Machesky, L.M. 2004. Regulation of WASP: PIP2 Pipped by Toca-1? Cell 118: 140-141.

107. Takenawa, T., and Suetsugu, S. 2007. The WASP-WAVE protein network: connecting the membrane to the cytoskeleton. Nat Rev Mol Cell Biol 8: 37-48.

108. Deeks, M.J., and Hussey, P.J. 2005. Arp2/3 and SCAR: plants move to the fore. Nat Rev Mol Cell Biol 6: 954-964.

109. Svitkina, T.M., and Borisy, G.G. 1999. Progress in protrusion: the tell-tale scar. TrendsBiochem Sci 24: 432-436.

110. Ruoslahti, E. 1997. Integrins as signaling molecules and targets for tumor therapy. Kidney Int 51: 1413-1417.

111. Nayal, A., Webb, D.J., and Horwitz, A.F. 2004. Talin: an emerging focal point of adhesion dynamics. Curr Opin Cell Biol 16: 94-98.

112. Le Clainche, C., and Carlier, M.F. 2008. Regulation of actin assembly associated with protrusion and adhesion in cell migration. Physiol Rev 88: 489-513.

113. Ziegler, W.H., Liddington, R.C., and Critchley, D.R. 2006. The structure and regulation of vinculin. Trends Cell Biol 16: 453-460.

114. Wehrle-Haller, B., and Imhof, B. 2002. The inner lives of focal adhesions. Trends Cell Biol 12: 382-389.

115. Geiger, B., Bershadsky, A., Pankov, R., and Yamada, K.M. 2001. Transmembrane crosstalk between the extracellular matrix--cytoskeleton crosstalk. Nat Rev Mol Cell Biol2: 793-805.

116. Chrzanowska-Wodnicka, M., and Burridge, K. 1996. Rho-stimulated contractility drives the formation of stress fibers and focal adhesions. J Cell Biol 133: 1403-1415.

198

117. Nobes, C.D., and Hall, A. 1995. Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell 81: 53-62.

118. Rottner, K., Hall, A., and Small, J.V. 1999. Interplay between Rac and Rho in the control of substrate contact dynamics. Curr Biol 9: 640-648.

119. Zamir, E., Katz, M., Posen, Y., Erez, N., Yamada, K.M., Katz, B.Z., Lin, S., Lin, D.C., Bershadsky, A., Kam, Z., et al. 2000. Dynamics and segregation of cell-matrix adhesions in cultured fibroblasts. Nat Cell Biol 2: 191-196.

120. Adams, J.C. 2002. Regulation of protrusive and contractile cell-matrix contacts. J Cell Sci 115: 257-265.

121. Chen, W.T., and Singer, S.J. 1982. Immunoelectron microscopic studies of the sites of cell-substratum and cell-cell contacts in cultured fibroblasts. J Cell Biol 95: 205-222.

122. Xiao, H., Eves, R., Yeh, C., Kan, W., Xu, F., Mak, A.S., and Liu, M. 2009. Phorbol ester-induced podosomes in normal human bronchial epithelial cells. J Cell Physiol 218: 366-375.

123. Riveline, D., Zamir, E., Balaban, N.Q., Schwarz, U.S., Ishizaki, T., Narumiya, S., Kam, Z., Geiger, B., and Bershadsky, A.D. 2001. Focal contacts as mechanosensors: externally applied local mechanical force induces growth of focal contacts by an mDia1-dependent and ROCK-independent mechanism. J Cell Biol 153: 1175-1186.

124. Nix, D.A., and Beckerle, M.C. 1997. Nuclear-cytoplasmic shuttling of the focal contact protein, zyxin: a potential mechanism for communication between sites of cell adhesion and the nucleus. J Cell Biol 138: 1139-1147.

125. Hobert, O., Schilling, J.W., Beckerle, M.C., Ullrich, A., and Jallal, B. 1996. SH3 domain-dependent interaction of the proto-oncogene product Vav with the focal contact protein zyxin. Oncogene 12: 1577-1581.

126. Cavalcanti-Adam, E.A., Volberg, T., Micoulet, A., Kessler, H., Geiger, B., and Spatz, J.P. 2007. Cell spreading and focal adhesion dynamics are regulated by spacing of integrin ligands. Biophys J 92: 2964-2974.

127. Huveneers, S., Truong, H., Fassler, R., Sonnenberg, A., and Danen, E.H. 2008. Binding of soluble fibronectin to integrin alpha5 beta1 - link to focal adhesion redistribution and contractile shape. J Cell Sci 121: 2452-2462.

128. Teti, A., Migliaccio, S., Taranta, A., Bernardini, S., De Rossi, G., Luciani, M., Iacobini, M., De Felice, L., Boldrini, R., Bosman, C., et al. 1999. Mechanisms of osteoclast dysfunction in human osteopetrosis: abnormal osteoclastogenesis and lack of osteoclast-specific adhesion structures. J Bone Miner Res 14: 2107-2117.

199

129. Grimaldi, C., Pisanti, S., Laezza, C., Malfitano, A.M., Santoro, A., Vitale, M., Caruso, M.G., Notarnicola, M., Iacuzzo, I., Portella, G., et al. 2006. Anandamide inhibits adhesion and migration of breast cancer cells. Exp Cell Res 312: 363-373.

130. Zeller, Y., Lohr, J., Sammar, M., Butcher, E.C., and Altevogt, P. 1998. Asp-698 and Asp-811 of the integrin alpha4-subunit are critical for the formation of a functional heterodimer. J Biol Chem 273: 6786-6795.

131. Hyland, R.H., Douglass, W.A., Tan, S.M., and Law, S.K. 2001. Chimeras of the integrin beta subunit mid-region reveal regions required for heterodimer formation and for activation. Cell Commun Adhes 8: 61-69.

132. Jannuzi, A.L., Bunch, T.A., West, R.F., and Brower, D.L. 2004. Identification of integrin beta subunit mutations that alter heterodimer function in situ. Mol Biol Cell 15: 3829-3840.

133. Wippler, J., Kouns, W.C., Schlaeger, E.J., Kuhn, H., Hadvary, P., and Steiner, B. 1994. The integrin alpha IIb-beta 3, platelet glycoprotein IIb-IIIa, can form a functionally active heterodimer complex without the cysteine-rich repeats of the beta 3 subunit. J Biol Chem269: 8754-8761.

134. Gupta, V., Alonso, J.L., Sugimori, T., Essafi, M., Xiong, J.P., and Arnaout, M.A. 2008. Role of the beta-subunit arginine/lysine finger in integrin heterodimer formation and function. J Immunol 180: 1713-1718.

135. Lo, S.H. 2004. Tensin. Int J Biochem Cell Biol 36: 31-34.

136. Zhang, Z., Baron, R., and Horne, W.C. 2000. Integrin engagement, the actin cytoskeleton, and c-Src are required for the calcitonin-induced tyrosine phosphorylation of paxillin and HEF1, but not for calcitonin-induced Erk1/2 phosphorylation. J Biol Chem 275: 37219-37223.

137. Manie, S.N., Beck, A.R., Astier, A., Law, S.F., Canty, T., Hirai, H., Druker, B.J., Avraham, H., Haghayeghi, N., Sattler, M., et al. 1997. Involvement of p130(Cas) and p105(HEF1), a novel Cas-like docking protein, in a cytoskeleton-dependent signaling pathway initiated by ligation of integrin or antigen receptor on human B cells. J Biol Chem 272: 4230-4236.

138. Ohmori, T., Yatomi, Y., Asazuma, N., Satoh, K., and Ozaki, Y. 2000. Involvement of proline-rich tyrosine kinase 2 in platelet activation: tyrosine phosphorylation mostly dependent on alphaIIbbeta3 integrin and protein kinase C, translocation to the cytoskeleton and association with Shc through Grb2. Biochem J 347: 561-569.

139. Obergfell, A., Judd, B.A., del Pozo, M.A., Schwartz, M.A., Koretzky, G.A., and Shattil, S.J. 2001. The molecular adapter SLP-76 relays signals from platelet integrin alphaIIbbeta3 to the actin cytoskeleton. J Biol Chem 276: 5916-5923.

200

140. Mainiero, F., Pepe, A., Wary, K.K., Spinardi, L., Mohammadi, M., Schlessinger, J., and Giancotti, F.G. 1995. Signal transduction by the alpha 6 beta 4 integrin: distinct beta 4 subunit sites mediate recruitment of Shc/Grb2 and association with the cytoskeleton of hemidesmosomes. EMBO J 14: 4470-4481.

141. Buccione, R., Orth, J.D., and McNiven, M.A. 2004. Foot and mouth: podosomes, invadopodia and circular dorsal ruffles. Nat Rev Mol Cell Biol 5: 647-657.

142. Linder, S., and Aepfelbacher, M. 2003. Podosomes: adhesion hot-spots of invasive cells. Trends Cell Biol 13: 376-385.

143. Condeelis, J., and Pollard, J.W. 2006. Macrophages: obligate partners for tumor cell migration, invasion, and metastasis. Cell 124: 263-266.

144. Evans, J.G., Correia, I., Krasavina, O., Watson, N., and Matsudaira, P. 2003. Macrophage podosomes assemble at the leading lamella by growth and fragmentation. J Cell Biol 161: 697-705.

145. Evans, J.G., and Matsudaira, P. 2006. Structure and dynamics of macrophage podosomes. Eur J Cell Biol 85: 145-149.

146. Eves, R., Webb, B.A., Zhou, S., and Mak, A.S. 2006. Caldesmon is an integral component of podosomes in smooth muscle cells. J Cell Sci 119: 1691-1702.

147. Varon, C., Tatin, F., Moreau, V., Van Obberghen-Schilling, E., Fernandez-Sauze, S., Reuzeau, E., Kramer, I., and Genot, E. 2006. Transforming growth factor beta induces rosettes of podosomes in primary aortic endothelial cells. Mol Cell Biol 26: 3582-3594.

148. Seals, D.F., Azucena, E.F., Jr., Pass, I., Tesfay, L., Gordon, R., Woodrow, M., Resau, J.H., and Courtneidge, S.A. 2005. The adaptor protein Tks5/Fish is required for podosome formation and function, and for the protease-driven invasion of cancer cells. Cancer Cell 7: 155-165.

149. Buccione, R., Baldassarre, M., Trapani, V., Catalano, C., Pompeo, A., Brancaccio, A., Giavazzi, R., Luini, A., and Corda, D. 2005. Glycerophosphoinositols inhibit the ability of tumour cells to invade the extracellular matrix. Eur J Cancer 41:470-476.

150. Spinardi, L., and Marchisio, P.C. 2006. Podosomes as smart regulators of cellular adhesion. Eur J Cell Biol 85: 191-194.

151. Spinardi, L., Rietdorf, J., Nitsch, L., Bono, M., Tacchetti, C., Way, M., and Marchisio, P.C. 2004. A dynamic podosome-like structure of epithelial cells. Exp Cell Res 295: 360-374.

152. Linder, S. 2007. The matrix corroded: podosomes and invadopodia in extracellular matrix degradation. Trends Cell Biol 17: 107-117.

201

153. Collin, O., Na, S., Chowdhury, F., Hong, M., Shin, M.E., Wang, F., and Wang, N. 2008. Self-organized podosomes are dynamic mechanosensors. Curr Biol 18: 1288-1294.

154. Stylli, S.S., Kaye, A.H., and Lock, P. 2008. Invadopodia: at the cutting edge of tumour invasion. J Clin Neurosci 15: 725-737.

155. Mueller, S.C., and Chen, W.T. 1991. Cellular invasion into matrix beads: localization of beta 1 integrins and fibronectin to the invadopodia. J Cell Sci 99 ( Pt 2): 213-225.

156. Baldassarre, M., Pompeo, A., Beznoussenko, G., Castaldi, C., Cortellino, S., McNiven, M.A., Luini, A., and Buccione, R. 2003. Dynamin participates in focal extracellular matrix degradation by invasive cells. Mol Biol Cell 14: 1074-1084.

157. Chen, W.T., Olden, K., Bernard, B.A., and Chu, F.F. 1984. Expression of transformation-associated protease(s) that degrade fibronectin at cell contact sites. J Cell Biol 98: 1546-1555.

158. Chen, W.T., Chen, J.M., Parsons, S.J., and Parsons, J.T. 1985. Local degradation of fibronectin at sites of expression of the transforming gene product pp60src. Nature 316: 156-158.

159. Artym, V.V., Zhang, Y., Seillier-Moiseiwitsch, F., Yamada, K.M., and Mueller, S.C. 2006. Dynamic interactions of cortactin and membrane type 1 matrix metalloproteinase at invadopodia: defining the stages of invadopodia formation and function. Cancer Res 66: 3034-3043.

160. Clark, E.S., and Weaver, A.M. 2008. A new role for cortactin in invadopodia: regulation of protease secretion. Eur J Cell Biol 87: 581-590.

161. Tehrani, S., Tomasevic, N., Weed, S., Sakowicz, R., and Cooper, J.A. 2007. Src phosphorylation of cortactin enhances actin assembly. Proc Natl Acad Sci U S A 104: 11933-11938.

162. Clark, E.S., Whigham, A.S., Yarbrough, W.G., and Weaver, A.M. 2007. Cortactin is an essential regulator of matrix metalloproteinase secretion and extracellular matrix degradation in invadopodia. Cancer Res 67: 4227-4235.

163. Block, M.R., Badowski, C., Millon-Fremillon, A., Bouvard, D., Bouin, A.P., Faurobert, E., Gerber-Scokaert, D., Planus, E., and Albiges-Rizo, C. 2008. Podosome-type adhesions and focal adhesions, so alike yet so different. Eur J Cell Biol 87: 491-506.

164. Frame, M.C. 2004. Newest findings on the oldest oncogene; how activated src does it. JCell Sci 117: 989-998.

165. Manning, G., Whyte, D.B., Martinez, R., Hunter, T., and Sudarsanam, S. 2002. The protein kinase complement of the human genome. Science 298: 1912-1934.

202

166. Berdeaux, R.L., Diaz, B., Kim, L., and Martin, G.S. 2004. Active Rho is localized to podosomes induced by oncogenic Src and is required for their assembly and function. JCell Biol 166: 317-323.

167. Moreau, V., Tatin, F., Varon, C., and Genot, E. 2003. Actin can reorganize into podosomes in aortic endothelial cells, a process controlled by Cdc42 and RhoA. Mol Cell Biol 23: 6809-6822.

168. Mizutani, K., Miki, H., He, H., Maruta, H., and Takenawa, T. 2002. Essential role of neural Wiskott-Aldrich syndrome protein in podosome formation and degradation of extracellular matrix in src-transformed fibroblasts. Cancer Res 62: 669-674.

169. Destaing, O., Sanjay, A., Itzstein, C., Horne, W.C., Toomre, D., De Camilli, P., and Baron, R. 2008. The tyrosine kinase activity of c-Src regulates actin dynamics and organization of podosomes in osteoclasts. Mol Biol Cell 19: 394-404.

170. Heckel, T., Czupalla, C., Expirto Santo, A.I., Anitei, M., Arantzazu Sanchez-Fernandez, M., Mosch, K., Krause, E., and Hoflack, B. 2009. Src-dependent repression of ARF6 is required to maintain podosome-rich sealing zones in bone-digesting osteoclasts. ProcNatl Acad Sci U S A 106: 1451-1456.

171. Dorfleutner, A., Cho, Y., Vincent, D., Cunnick, J., Lin, H., Weed, S.A., Stehlik, C., and Flynn, D.C. 2008. Phosphorylation of AFAP-110 affects podosome lifespan in A7r5 cells. J Cell Sci 121: 2394-2405.

172. Blouw, B., Seals, D.F., Pass, I., Diaz, B., and Courtneidge, S.A. 2008. A role for the podosome/invadopodia scaffold protein Tks5 in tumor growth in vivo. Eur J Cell Biol 87: 555-567.

173. Buschman, M.D., Bromann, P.A., Cejudo-Martin, P., Wen, F., Pass, I., and Courtneidge, S.A. 2009. The Novel Adaptor Protein Tks4 (SH3PXD2B) Is Required for Functional Podosome Formation. Mol Biol Cell 20: 1302-11.

174. Oikawa, T., Itoh, T., and Takenawa, T. 2008. Sequential signals toward podosome formation in NIH-src cells. J Cell Biol 182: 157-169.

175. Yang, X., Qiao, D., Meyer, K., and Friedl, A. 2009. Signal Transducers and Activators of Transcription Mediate Fibroblast Growth Factor-Induced Vascular Endothelial Morphogenesis. Cancer Res 69: 1668-77.

176. Zhao, J., and Guan, J.L. 2009. Signal transduction by focal adhesion kinase in cancer. Cancer Metastasis Rev 28: 35-49.

177. Narumiya, S., Tanji, M., and Ishizaki, T. 2009. Rho signaling, ROCK and mDia1, in transformation, metastasis and invasion. Cancer Metastasis Rev 28: 65-76.

178. Wang, S.E., Xiang, B., Zent, R., Quaranta, V., Pozzi, A., and Arteaga, C.L. 2009. Transforming growth factor beta induces clustering of HER2 and integrins by activating

203

Src-focal adhesion kinase and receptor association to the cytoskeleton. Cancer Res 69: 475-482.

179. Brozzi, F., Arcuri, C., Giambanco, I., and Donato, R. 2009. S100B protein regulates astrocyte shape and migration via interaction with Src kinase: implications for astrocyte development, activation and tumor growth. J Biol Chem 284: 8797-8811.

180. Huot, M.E., Brown, C.M., Lamarche-Vane, N., and Richard, S. 2009. An adaptor role for cytoplasmic Sam68 in modulating Src activity during cell polarization. Mol Cell Biol 29: 1933-1943.

181. Vanhaesebroeck, B., Leevers, S.J., Ahmadi, K., Timms, J., Katso, R., Driscoll, P.C., Woscholski, R., Parker, P.J., and Waterfield, M.D. 2001. Synthesis and function of 3-phosphorylated inositol lipids. Annu Rev Biochem 70: 535-602.

182. Okkenhaug, K., and Vanhaesebroeck, B. 2003. PI3K in lymphocyte development, differentiation and activation. Nat Rev Immunol 3: 317-330.

183. Vanhaesebroeck, B., Leevers, S.J., Panayotou, G., and Waterfield, M.D. 1997. Phosphoinositide 3-kinases: a conserved family of signal transducers. Trends Biochem Sci 22: 267-272.

184. Falasca, M., and Maffucci, T. 2007. Role of class II phosphoinositide 3-kinase in cell signalling. Biochem Soc Trans 35: 211-214.

185. Hawkins, P.T., Anderson, K.E., Davidson, K., and Stephens, L.R. 2006. Signalling through Class I PI3Ks in mammalian cells. Biochem Soc Trans 34: 647-662.

186. Jiang, B.H., and Liu, L.Z. 2008. PI3K/PTEN signaling in tumorigenesis and angiogenesis. Biochim Biophys Acta 1784: 150-158.

187. Stocker, H., Andjelkovic, M., Oldham, S., Laffargue, M., Wymann, M.P., Hemmings, B.A., and Hafen, E. 2002. Living with lethal PIP3 levels: viability of flies lacking PTEN restored by a PH domain mutation in Akt/PKB. Science 295: 2088-2091.

188. Vanhaesebroeck, B., and Alessi, D.R. 2000. The PI3K-PDK1 connection: more than just a road to PKB. Biochem J 346 Pt 3: 561-576.

189. Luo, J., Manning, B.D., and Cantley, L.C. 2003. Targeting the PI3K-Akt pathway in human cancer: rationale and promise. Cancer Cell 4: 257-262.

190. Toker, A., and Cantley, L.C. 1997. Signalling through the lipid products of phosphoinositide-3-OH kinase. Nature 387: 673-676.

191. Franke, T.F., Kaplan, D.R., Cantley, L.C., and Toker, A. 1997. Direct regulation of the Akt proto-oncogene product by phosphatidylinositol-3,4-bisphosphate. Science 275: 665-668.

204

192. Shaw, M., Cohen, P., and Alessi, D.R. 1997. Further evidence that the inhibition of glycogen synthase kinase-3beta by IGF-1 is mediated by PDK1/PKB-induced phosphorylation of Ser-9 and not by dephosphorylation of Tyr-216. FEBS Lett 416: 307-311.

193. Alessi, D.R., James, S.R., Downes, C.P., Holmes, A.B., Gaffney, P.R., Reese, C.B., and Cohen, P. 1997. Characterization of a 3-phosphoinositide-dependent protein kinase which phosphorylates and activates protein kinase Balpha. Curr Biol 7: 261-269.

194. Alessi, D.R., Deak, M., Casamayor, A., Caudwell, F.B., Morrice, N., Norman, D.G., Gaffney, P., Reese, C.B., MacDougall, C.N., Harbison, D., et al. 1997. 3-Phosphoinositide-dependent protein kinase-1 (PDK1): structural and functional homology with the Drosophila DSTPK61 kinase. Curr Biol 7: 776-789.

195. Sarbassov, D.D., Guertin, D.A., Ali, S.M., and Sabatini, D.M. 2005. Phosphorylation and regulation of Akt/PKB by the rictor-mTOR complex. Science 307: 1098-1101.

196. Mora, A., Komander, D., van Aalten, D.M., and Alessi, D.R. 2004. PDK1, the master regulator of AGC kinase signal transduction. Semin Cell Dev Biol 15: 161-170.

197. Datta, S.R., Brunet, A., and Greenberg, M.E. 1999. Cellular survival: a play in three Akts. Genes Dev 13: 2905-2927.

198. Stiles, B., Gilman, V., Khanzenzon, N., Lesche, R., Li, A., Qiao, R., Liu, X., and Wu, H. 2002. Essential role of AKT-1/protein kinase B alpha in PTEN-controlled tumorigenesis. Mol Cell Biol 22: 3842-3851.

199. Chellaiah, M.A. 2006. Regulation of podosomes by integrin alphavbeta3 and Rho GTPase-facilitated phosphoinositide signaling. Eur J Cell Biol 85: 311-317.

200. Takenawa, T., and Itoh, T. 2001. Phosphoinositides, key molecules for regulation of actin cytoskeletal organization and membrane traffic from the plasma membrane. BiochimBiophys Acta 1533: 190-206.

201. Walker, V.G., Ammer, A., Cao, Z., Clump, A.C., Jiang, B.H., Kelley, L.C., Weed, S.A., Zot, H., and Flynn, D.C. 2007. PI3K activation is required for PMA-directed activation of cSrc by AFAP-110. Am J Physiol Cell Physiol 293: C119-132.

202. Junttila, M.R., Li, S.P., and Westermarck, J. 2008. Phosphatase-mediated crosstalk between MAPK signaling pathways in the regulation of cell survival. FASEB J 22: 954-965.

203. Raman, M., Chen, W., and Cobb, M.H. 2007. Differential regulation and properties of MAPKs. Oncogene 26: 3100-3112.

204. Dhillon, A.S., Hagan, S., Rath, O., and Kolch, W. 2007. MAP kinase signalling pathways in cancer. Oncogene 26: 3279-3290.

205

205. Schaeffer, H.J., and Weber, M.J. 1999. Mitogen-activated protein kinases: specific messages from ubiquitous messengers. Mol Cell Biol 19: 2435-2444.

206. Chen, Z., Gibson, T.B., Robinson, F., Silvestro, L., Pearson, G., Xu, B., Wright, A., Vanderbilt, C., and Cobb, M.H. 2001. MAP kinases. Chem Rev 101: 2449-2476.

207. Avruch, J., Khokhlatchev, A., Kyriakis, J.M., Luo, Z., Tzivion, G., Vavvas, D., and Zhang, X.F. 2001. Ras activation of the Raf kinase: tyrosine kinase recruitment of the MAP kinase cascade. Recent Prog Horm Res 56: 127-155.

208. Kyriakis, J.M., and Avruch, J. 2001. Mammalian mitogen-activated protein kinase signal transduction pathways activated by stress and inflammation. Physiol Rev 81: 807-869.

209. Davis, R.J. 2000. Signal transduction by the JNK group of MAP kinases. Cell 103: 239-252.

210. Weston, C.R., and Davis, R.J. 2007. The JNK signal transduction pathway. Curr Opin Cell Biol 19: 142-149.

211. Katz, M., Amit, I., and Yarden, Y. 2007. Regulation of MAPKs by growth factors and receptor tyrosine kinases. Biochim Biophys Acta 1773: 1161-1176.

212. Shaul, Y.D., and Seger, R. 2007. The MEK/ERK cascade: from signaling specificity to diverse functions. Biochim Biophys Acta 1773: 1213-1226.

213. Hai, C.M., and Gu, Z. 2006. Caldesmon phosphorylation in actin cytoskeletal remodeling. Eur J Cell Biol 85: 305-309.

214. Gu, Z., Kordowska, J., Williams, G.L., Wang, C.L., and Hai, C.M. 2007. Erk1/2 MAPK and caldesmon differentially regulate podosome dynamics in A7r5 vascular smooth muscle cells. Exp Cell Res 313: 849-866.

215. Luckashenak, N.A., Ryszkiewicz, R.L., Ramsey, K.D., and Clements, J.L. 2006. The Src homology 2 domain-containing leukocyte protein of 76-kDa adaptor links integrin ligation with p44/42 MAPK phosphorylation and podosome distribution in murine dendritic cells. J Immunol 177: 5177-5185.

216. Szczur, K., Xu, H., Atkinson, S., Zheng, Y., and Filippi, M.D. 2006. Rho GTPase CDC42 regulates directionality and random movement via distinct MAPK pathways in neutrophils. Blood 108: 4205-4213.

217. Schramp, M., Ying, O., Kim, T.Y., and Martin, G.S. 2008. ERK5 promotes Src-induced podosome formation by limiting Rho activation. J Cell Biol 181: 1195-1210.

218. Kuo, L., Chang, H.C., Leu, T.H., Maa, M.C., and Hung, W.C. 2006. Src oncogene activates MMP-2 expression via the ERK/Sp1 pathway. J Cell Physiol 207: 729-734.

206

219. Ruhul Amin, A.R., Oo, M.L., Senga, T., Suzuki, N., Feng, G.S., and Hamaguchi, M. 2003. SH2 domain containing protein tyrosine phosphatase 2 regulates concanavalin A-dependent secretion and activation of matrix metalloproteinase 2 via the extracellular signal-regulated kinase and p38 pathways. Cancer Res 63: 6334-6339.

220. Hauck, C.R., Sieg, D.J., Hsia, D.A., Loftus, J.C., Gaarde, W.A., Monia, B.P., and Schlaepfer, D.D. 2001. Inhibition of focal adhesion kinase expression or activity disrupts epidermal growth factor-stimulated signaling promoting the migration of invasive human carcinoma cells. Cancer Res 61: 7079-7090.

221. Kelly, T., Mueller, S.C., Yeh, Y., Chen, W.T. 1994. Invadopodia promote proteolysis of a wide variety of extracellular matrix proteins. J Cell Physiol. 158: 299-308.

222. Nakahara, H., Nomizu, M., Akiyama, S.K., Yamada, Y., Yeh, Y., Chen, W.T. 1996. A mechanism for regulation of melanoma invasion. Ligation of alpha6beta1 integrin by laminin G peptides. J Biol Chem 271: 27221-27224.

223. Nakahara, H., Howard, L, Thompson, E.W., Sato, H., Seiki, M., Yeh, Y.Y., Chen, W.T. 1997. Transmembrane/cytoplasmic domain-mediated membrane type 1-matrix metalloprotease docking to invadopodia is required for cell invasion. Proc Natl Acad Sci94: 7959-7964.

224. Chang, C., Werb, Z. 2001. The many faces of metalloproteinases: cell growth, invasion, angiogenesis and metastasis. Trends Cell Biol 11: S37-S43.

225. Shin, Y., Yoon, S.H., Choe, E.Y., Cho, S.H., Woo, C.H., Rho, J.Y., Kim, J.H. 2007. PMA-induced up-regulation of MMP-9 is regulated by a PKCalpha-NF-kappaB cascade in human lung epithelial cells. Exp Mol Med 39: 97-105.

226. Chetty, C., Bhoopathi, P., Joseph, P., Chittivelu, S., Rao, J.S., Lakka, S. 2006. Adenovirus-mediated small interfering RNA against matrix metalloproteinase-2 suppresses tumor growth and lung metastasis in mice. Mol Cancer Ther 5: 2289-2299.

227. Nagase, H., Woessner, J.F. Jr. 1999. Matrix Metalloproteinases. J Biol Chem 274: 21491-21494.

228. Kinoshita, T., Sato, H., Okada, A., Ohuchi, E., Imai, K., Okada, Y., Seiki, M. 1998. TIMP-2 promotes activation of progelatinase A by membrane-type 1 matrix metalloproteinase immobilized on agarose beads. J Biol Chem 273: 16098-16103.

229. Strongin, A.Y., Collier, I., Bannikov, G., Marmer, B.L., Grant, G.A., Goldberg, G.I. 1995. Mechanism of cell surface activation of 72-kDa type IV collagenase. Isolation of the activated form of the membrane metalloprotease. J Biol Chem 270: 5331-5338.

230. Juarez, J., Clayman, G., Nakajima, M., Tanabe, K.K., Saya, H., Nicolson, G.L., and Boyd, D. 1993. Role and regulation of expression of 92-kDa type-IV collagenase (MMP-9) in 2 invasive squamous-cell-carcinoma cell lines of the oral cavity. Int J Cancer 55: 10-18.

207

231. Simon, C., Goepfert, H., and Boyd, D. 1998. Inhibition of the p38 mitogen-activated protein kinase by SB 203580 blocks PMA-induced Mr 92,000 type IV collagenase secretion and in vitro invasion. Cancer Res 58: 1135-1139.

232. Genersch, E., Hayess, K., Neuenfeld, Y., and Haller, H. 2000. Sustained ERK phosphorylation is necessary but not sufficient for MMP-9 regulation in endothelial cells: involvement of Ras-dependent and -independent pathways. J Cell Sci 113 Pt 23: 4319-4330.

233. Park, M.J., Park, I.C., Hur, J.H., Rhee, C.H., Choe, T.B., Yi, D.H., Hong, S.I., and Lee, S.H. 2000. Protein kinase C activation by phorbol ester increases in vitro invasion through regulation of matrix metalloproteinases/tissue inhibitors of metalloproteinases system in D54 human glioblastoma cells. Neurosci Lett 290: 201-204.

234. Park, M.J., Park, I.C., Lee, H.C., Woo, S.H., Lee, J.Y., Hong, Y.J., Rhee, C.H., Lee, Y.S., Lee, S.H., Shim, B.S., et al. 2003. Protein kinase C-alpha activation by phorbol ester induces secretion of gelatinase B/MMP-9 through ERK 1/2 pathway in capillary endothelial cells. Int J Oncol 22: 137-143.

235. Hah, N., and Lee, S.T. 2003. An absolute role of the PKC-dependent NF-kappaB activation for induction of MMP-9 in hepatocellular carcinoma cells. Biochem Biophys Res Commun 305: 428-433.

236. Ko, F.W., Diba, C., Roth, M., McKay, K., Johnson, P.R., Salome, C., and King, G.G. 2005. A comparison of airway and serum matrix metalloproteinase-9 activity among normal subjects, asthmatic patients, and patients with asthmatic mucus hypersecretion. Chest 127: 1919-1927.

237. Esteve, P.O., Chicoine, E., Robledo, O., Aoudjit, F., Descoteaux, A., Potworowski, E.F., and St-Pierre, Y. 2002. Protein kinase C-zeta regulates transcription of the matrix metalloproteinase-9 gene induced by IL-1 and TNF-alpha in glioma cells via NF-kappa B. J Biol Chem 277: 35150-35155.

238. Esteve, P.O., Robledo, O., Potworowski, E.F., and St-Pierre, Y. 2002. Induced expression of MMP-9 in C6 glioma cells is inhibited by PDGF via a PI 3-kinase-dependent pathway. Biochem Biophys Res Commun 296: 864-869.

239. Liu, J.F., Crepin, M., Liu, J.M., Barritault, D., and Ledoux, D. 2002. FGF-2 and TPA induce matrix metalloproteinase-9 secretion in MCF-7 cells through PKC activation of the Ras/ERK pathway. Biochem Biophys Res Commun 293: 1174-1182.

240. Xie, B., Laouar, A., and Huberman, E. 1998. Fibronectin-mediated cell adhesion is required for induction of 92-kDa type IV collagenase/gelatinase (MMP-9) gene expression during macrophage differentiation. The signaling role of protein kinase C-beta. J Biol Chem 273: 11576-11582.

208

241. Goto, T., Maeda, H., Tanaka, T. 2002. A selective inhibitor of matrix metalloproteinases inhibits the migration of isolated osteoclasts by increasing the life span of podosomes. JBone Miner Metab 20: 98-105.

242. Artym, V.V., Zhang, Y., Seillier-Moiseiwitsch, F., Yamada, K.M., Mueller, S.C. 2006. Dynamic interactions of cortactin and membrane type 1 matrix metalloproteinase at invadopodia: defining the stages of invadopodia formation and function. Cancer Res 66: 3034-3043.

243. Itoh, Y., Seiki, M. 2004. MT1-MMP: an enzyme with multidimensional regulation. Trends Biochem Sci 29: 285–289.

244. Imai, K., Ohuchi, E., Aoki, T., Nomura, H, Fujii, Y., Sato, H., Seiki, M., Okada, Y. 1996. Membrane-type matrix metalloproteinase 1 is a gelatinolytic enzyme and is secreted in a complex with tissue inhibitor of metalloproteinases 2. Cancer Res 56: 2707-2710.

245. Pei, D., Weiss, S.J. . 1996. Transmembrane-deletion mutants of the membrane-type matrix metalloproteinase-1 process progelatinase A and express intrinsic matrix-degrading activity. J Biol Chem 271: 9135-9140.

246. Ohuchi, E., Imai, K., Fujii, Y., Sato, H., Seiki, M., Okada, Y. 1997. Membrane type 1 matrix metalloproteinase digests interstitial collagens and other extracellular matrix macromolecules. J Biol Chem 272: 2446-2451.

247. Bodey, B., Bodey B. Jr., Groger, A.M., Siegel, S.E., Kaiser, H.E. 2001. Invasion and metastasis: the expression and significance of matrix metalloproteinases in carcinomas of the lung. In Vivo 15: 175-180.

248. Monsky, W.L., Kelly, T., Lin, C.Y., Yeh, Y., Stetler-Stevenson, W.G., Mueller, S.C., Chen, W.T. 1993. Binding and localization of M(r) 72,000 matrix metalloproteinase at cell surface invadopodia. Cancer Res 53: 3159-3164.

249. Sato, H., Seiki, M. 1993. Regulatory mechanism of 92 kDa type IV collagenase gene expression which is associated with invasiveness of tumor cells. Oncogene 8: 395–405.

250. Poincloux, R., Vincent, C., Labrousse, A., Castandet, J., Rigo, M., Cougoule, C., et al. 2006. Re-arrangements of podosome structures are observed when Hck is activated in myeloid cells. Eur J Cell Biol 85: 327-332.

251. Buisson, A.C., Zahm, J.M., Polette, M., Pierrot, D., Bellon, G., Puchelle, E., Birembaut, P., Tournier, J.M. . 1996a. Gelatinase B is involved in the in vitro wound repair of human respiratory epithelium. J Cell Physiol 166: 413-426.

252. Ågren, M.S. 1994. Gelatinase activity during wound healing. Br J Dermatol 131: 634-640.

209

253. Stricklin, G.P., Li, L., Nanney, L.B. . 1994. Localization of mRNAs representing interstitial collagenase, 72-kDa gelatinase, and TIMP in healing porcine burn wounds. JInvest Dermatol 103: 352-358.

254. Bendeck, M.P., Zempo, N., Clowes, A.W., Galardy, R.E., Reidy, M.A. 1994. Smooth muscle cell migration and matrix metalloproteinase expression after arterial injury in the rat. Circ Res 75: 539-545.

255. Daniels, J.T., Geerling, G., Alexander, R.A., Murphy, G., Khaw, P.T., Saarialho-Kere, U. 2003. Temporal and spatial expression of matrix metalloproteinases during wound healing of human corneal tissue. . Exp Eye Res 77: 653-664.

256. Matsubara, M., Girard, M.T., Kublin, C.L., Cintron, C., Fini, M.E. . 1991. Differential roles for two gelatinolytic enzymes of the matrix metalloproteinase family in the remodeling cornea. Dev Biol 147: 425-439.

257. Oikarinen, A., Kylmäniemi, M., Autio-Harmainen, H., Autio, P., Salo, T. 1993. Demonstration of 72-kDa and 92-kDa forms of type IV collagenase in human skin: variable expression in various blistering diseases, induction during re-epithelialization,and decrease by topical glucocorticoids. J Invest Dermatol 101: 205-210.

258. Salo, T., Mäkelä, M., Kylmäniemi, M., Autio-Harmainen, H., Larjava, H. 1994. Expression of matrix metalloproteinase-2 and -9 during early human wound healing. LabInvest 70: 176-182.

259. Yu, Q., Stamenkovic, I. 2000. Cell surface-localized matrix metalloproteinase-9 proteolytically activates TGF-beta and promotes tumor invasion and angiogenesis. GenesDev 14: 163-176.

260. Pluznik, D.H., Fridman, R., Reich, R. 1992. Correlation in the expression of type IV collagenase and the invasive and chemotactic abilities of myelomonocytic cells differentiation into macrophages. Exp Hematol 20: 57-63.

261. Leppert, D., Waubant, E., Galardy, R., Bunnett, N.W., Hauser, S.L. 1995. T cell gelatinases mediate basement membrane transmigration in vitro. J Immunol 154: 4379-4389.

262. Delclaux, C., Delacourt, C., d'Ortho, M.P., Boyer, V., Lafuma, C., Harf, A. 1996. Role of gelatinase B and elastase in human polymorphonuclear neutrophil migration across basement membrane. Am J Respir Cell Mol Biol 14: 288-295.

263. Okada, S., Kita, H., George, J., Gleich, G.J., Leiferman, K. 1997. Migration of eosinophils through basement membrane components in vitro: role of matrix metalloproteinase-9. Am J Respir Cell Mol Biol 17: 519-528.

264. McCawley, L.J., O'Brien, P., Hudson, L.G. 1998. Epidermal growth factor (EGF)- and scatter factor/hepatocyte growth factor (SF/HGF)-mediated keratinocyte migration is

210

coincident with induction of matrix metalloproteinase (MMP-9). J Cell Physiol 176: 255-265.

265. Vu, T.H., Werb, Z. . 1998. Gelatinase B: structure, regulation and function. In Matrix Metalloproteinases (Biology of Extracellular Matrix Series). W.C. Parks, Mecham, R.P., editor. New York: Academic Press. 115-148.

266. Lee, J., Ishihara, A., Jacobson, K. 1993. How do cells move along surfaces? Trends Cell Biol 3: 366-370.

267. Small, J.V., Anderson, K., Rottner, K. 1996. Actin and the coordination of protrusion, attachment and retraction in cell crawling. Biosci Rep 16: 351-368.

268. Vicente-Manzanares, M., Cruz-Adalia, A., Martin-Cofreces, N.B., Cabrero, J.R., Dosil, M., Alvarado-Sanchez, B., Bustelo, X.R., and Sanchez-Madrid, F. 2005. Control of lymphocyte shape and the chemotactic response by the GTP exchange factor Vav. Blood105: 3026-3034.

269. Vicente-Manzanares, M., Webb, D.J., and Horwitz, A.R. 2005. Cell migration at a glance. J Cell Sci 118: 4917-4919.

270. Bouvard, D., Aszodi, A., Kostka, G., Block, M.R., Albiges-Rizo, C., and Fassler, R. 2007. Defective osteoblast function in ICAP-1-deficient mice. Development 134: 2615-2625.

271. Larsen, M., Artym, V.V., Green, J.A., and Yamada, K.M. 2006. The matrix reorganized: extracellular matrix remodeling and integrin signaling. Curr Opin Cell Biol 18: 463-471.

272. Linder, S. 2007. The matrix corroded: podosomes and invadopodia in extracellular matrix degradation. Trends Cell Biol 17: 107-117.

273. Gimona, M., Buccione, R., Courtneidge, S.A., Linder, S. 2008. Assembly and biological role of podosomes and invadopodia. Curr Opin Cell Biol 20: 235-241.

274. Yamaguchi, H., Pixley, F., and Condeelis, J. 2006. Invadopodia and podosomes in tumor invasion. Eur J Cell Biol 85: 213-218.

275. Yamaguchi, H., Lorenz, M., Kempiak, S., et al. 2005. Molecular mechanisms of invadopodium formation: the role of the N-WASP-Arp2/3 complex pathway and cofilin.J Cell Biol 168: 441–452.

276. Carman, C.V., Sage, P.T., Sciuto, T.E., de la Fuente, M.A., Geha, R.S., Ochs, H.D., Dvorak, H.F., Dvorak, A.M., and Springer, T.A. 2007. Transcellular diapedesis is initiated by invasive podosomes. Immunity 26: 784-797.

277. Carman, C.V., and Springer, T.A. 2008. Trans-cellular migration: cell-cell contacts get intimate. Curr Opin Cell Biol 20: 533-540.

211

278. Dejana, E. 2006. The transcellular railway: insights into leukocyte diapedesis. Nat Cell Biol 8: 105-107.

279. Yamaguchi, H., Wyckoff, J., and Condeelis, J. 2005. Cell migration in tumors. Curr Opin Cell Biol 17: 559-564.

280. Yamaguchi, H., Lorenz, M., Kempiak, S., Sarmiento, C., Coniglio, S., Symons, M., Segall, J., Eddy, R., Miki, H., Takenawa, T., et al. 2005. Molecular mechanisms of invadopodium formation: the role of the N-WASP-Arp2/3 complex pathway and cofilin. J Cell Biol 168: 441-452.

281. Lang, D., Beermann, M.L., Hauser, G., Cressman, C.M., and Shea, T.B. 1995. Phospholipids inhibit proteolysis of protein kinase C alpha by mM calcium-requiring calpain. Neurochem Res 20: 1361-1364.

282. Baekgaard, A., and Bindslev, N. 1998. Novel protein kinase C, nPKC, inhibition of murine bumetanide-sensitive Na+,K+,2Cl-cotransporter BSC1 in Xenopus oocyte. PfluArch Eur J Phy 436: 189-198.

283. Sehic, E., and Malik, K.U. 1988. Effect of Protein Kinase-C (Pkc) Inhibitors on Adrenergic Neurotransmitter Release in the Isolated Rat-Kidney. FASEB J 2: A1733-A1733.

284. George, O.K., Edwards, L.P., Hall, R.G., and Gonzalez, R.R. 2002. PKC modulation of vascular smooth muscle contraction in rat-tail artery is primarily through the AT(1) receptor. Mol Biol Cell 13: 274a-274a.

285. Makagiansar, I.T., Williams, S., Mustelin, T., and Stallcup, W.B. 2007. Differential phosphorylation of NG2 proteoglycan by ERK and PKC alpha helps balance cell proliferation and migration. J Cell Biol 178: 155-165.

286. Fong, D.K., Rao, A., Crump, F.T., and Craig, A.M. 2002. Rapid synaptic remodeling by protein kinase C: Reciprocal translocation of NMDA receptors and calcium/calmodulin-dependent kinase II. J Neurosci 22: 2153-2164.

287. Hori, K., Yasuda, H., Konno, D., Maruoka, H., Tsumoto, T., and Sobue, K. 2005. NMDA receptor-dependent synaptic translocation of insulin receptor substrate p53 via protein kinase C signaling. J Neurosci 25: 2670-2681.

288. Takeuchi, K., Nakamura, N., and Dzau, V.J. 1990. Angiotensin-Ii Can Regulate Gene-Expression by the Ap-1 Sequence Via a Pkc Dependent Pathway. Hypertension 16: 345-345.

289. Nakashima, S. 2002. Protein kinase C alpha (PKC alpha): Regulation and biological function. J Biochem 132: 669-675.

290. Liu, W.S., and Heckman, C.A. 1998. The sevenfold way of PKC regulation. Cell Signal10: 529-542.

212

291. Parker, P.J., and Murray-Rust, J. 2004. PKC at a glance. J Cell Sci 117: 131-132.

292. Griner, E.M., and Kazanietz, M.G. 2007. Protein kinase C and other diacylglycerol effectors in cancer. Nat Rev Cancer 7: 281-294.

293. Wang, Q.M.J. 2006. PKD at the crossroads of DAG and PKC signaling. TrendsPharmacol Sci 27: 317-323.

294. Oliva, J.L., Griner, E.M., and Kazanietz, M.G. 2005. PKC isozymes and diacylglycerol-regulated proteins as effectors of growth factor receptors. Growth Factors 23: 245-252.

295. Corbalan-Garcia, S., and Gomez-Fernandez, J.C. 2006. Protein kinase C regulatory domains: The art of decoding many different signals in membranes. BBA-Mol Cell Biol L1761: 633-654.

296. Ono, Y., Fujii, T., Igarashi, K., Kuno, T., Tanaka, C., Kikkawa, U., and Nishizuka, Y. 1989. Phorbol Ester Binding to Protein Kinase-C Requires a Cysteine-Rich Zinc-Finger-Like Sequence. Proc Natl Acad Sci USA 86: 4868-4871.

297. Hurley, J.H., Newton, A.C., Parker, P.J., Blumberg, P.M., and Nishizuka, Y. 1997. Taxonomy and function of C1 protein kinase C homology domains. Protein Sci 6: 477-480.

298. Gomez-Fernandez, J.C., Torrecillas, A., and Corbalan-Garcia, S. 2004. Diacylglycerols as activators of protein kinase C (Review). Mol Membr Biol 21: 339-349.

299. Newton, A.C., and Johnson, J.J. 1998. Protein kinase C: a paradigm for regulation of protein function by two membrane-targeting modules. BBA-Biomembranes 1376: 155-172.

300. Rizo, J., and Sudhof, T.C. 1998. C-2-domains, structure and function of a universal Ca2+-binding domain. J Biol Chem 273: 15879-15882.

301. Nalefski, E.A., and Falke, J.J. 1996. The C2 domain calcium-binding motif: Structural and functional diversity. Protein Sci 5: 2375-2390.

302. Colon-Gonzalez, F., and Kazanietz, M.G. 2006. C1 domains exposed: from diacylglycerol binding to protein-protein interactions. Biochim Biophys Acta 1761: 827-837.

303. Parekh, D.B., Ziegler, W., and Parker, P.J. 2000. Multiple pathways control protein kinase C phosphorylation. EMBO J 19: 496-503.

304. Keranen, L.M., Dutil, E.M., and Newton, A.C. 1995. Protein kinase C is regulated in vivo by three functionally distinct phosphorylations. Current Biology 5: 1394-1403.

305. Casabona, G. 1997. Intracellular signal modulation: A pivotal role for protein kinase C. Prog Neuro-Psychoph 21: 407-425.

213

306. Eshete, F., Chung, H., Gannon-Murakami, L., and Murakami, K. 1998. Ca2+ modulation of cis-unsaturated fatty acid-induced mutant protein kinase C activity: indication of inhibitory Ca2+-binding site in protein kinase C-alpha. Biochem J 333: 215-221.

307. Lambert, I.H., and Falktoft, B. 2001. Lysophosphatidylcholine-induced taurine release in HeLa cells involves protein kinase activity. Comp Biochem Phys A 130: 577-584.

308. Ways, D.K., Kukoly, C.A., Devente, J., Hooker, J.L., Bryant, W.O., Posekany, K.J., Fletcher, D.J., Cook, P.P., and Parker, P.J. 1995. Mcf-7 Breast-Cancer Cells Transfected with Protein-Kinase C-Alpha Exhibit Altered Expression of Other Protein-Kinase-C Isoforms and Display a More Aggressive Neoplastic Phenotype. J Clin Invest 95: 1906-1915.

309. Tan, M., Li, P., Sun, M., Yin, G., and Yu, D. 2006. Upregulation and activation of PKC alpha by ErbB2 through Src promotes breast cancer cell invasion that can be blocked by combined treatment with PKCa and Src inhibitors. Oncogene 25: 3286-3295.

310. Zhang, J., Anastasiadis, P.Z., Liu, Y., Thompson, E.A., and Fields, A.P. 2004. Protein kinase C (PKC) beta II induces cell invasion through a Ras/Mek-, PKC iota/Rac1-dependent signaling pathway. J Biol Chem 279: 22118-22123.

311. Ng, T., Shima, D., Squire, A., Bastiaens, P.I.H., Gschmeissner, S., Humphries, M.J., and Parker, P.J. 1999. PKC alpha regulates beta 1 integrin-dependent cell motility through association and control of integrin traffic. EMBO J 18: 3909-3923.

312. Parsons, M., Keppler, M.D., Kline, A., Messent, A., Humphries, M.J., Gilchrist, R., Hart, I.R., Quittau-Prevostel, C., Hughes, W.E., Parker, P.J., et al. 2002. Site-directed perturbation of protein kinase C- integrin interaction blocks carcinoma cell chemotaxis. Mol Cell Biol 22: 5897-5911.

313. Masur, K., Lang, K., Niggemann, B., Zanker, K.S., and Entschladen, F. 2001. High PKC alpha and low E-cadherin expression contribute to high migratory activity of colon carcinoma cells. Mol Biol Cell 12: 1973-1982.

314. Liu, B., Maher, R.J., Hannun, Y.A., Porter, A.T., and Honn, K.V. 1994. 12(S)-HETE enhancement of prostate tumor cell invasion: selective role of PKC alpha. J Natl Cancer Inst 86: 1145-1151.

315. Choe, Y., Jung, H., Khang, I., and Kim, K. 2003. Selective roles of protein kinase C isoforms on cell motility of GT1 immortalized hypothalamic neurones. JNeuroendocrinol 15: 508-515.

316. Liao, L., and Jaken, S. 1993. Effect of alpha-protein kinase C neutralizing antibodies and the pseudosubstrate peptide on phosphorylation, migration, and growth of REF52 cells. Cell Growth Differ 4: 309-316.

214

317. Fanning, A., Volkov, Y., Freeley, M., Kelleher, D., and Long, A. 2005. CD44 cross-linking induces protein kinase C-regulated migration of human T lymphocytes. Int Immunol 17: 449-458.

318. Gauthier, M.L., Torretto, C., Ly, J., Francescutti, V., and O'Day, D.H. 2003. Protein kinase Calpha negatively regulates cell spreading and motility in MDA-MB-231 human breast cancer cells downstream of epidermal growth factor receptor. Biochem Biophys Res Commun 307: 839-846.

319. Kikkawa, U., Matsuzaki, H., and Yamamoto, T. 2002. Protein kinase C delta (PKC delta): Activation mechanisms and functions. J Biochem 132: 831-839.

320. Gschwendt, M. 1999. Protein kinase C delta. Eur J Biochem 259: 555-564.

321. Gschwendt, M., Stempka, L., Schnolzer, M., Radke, S., Rincke, G., and Marks, F. 1999. Requirements of protein kinase C delta for catalytic function. FASEB J 13: A1530-A1530.

322. Brenner, W., Greber, I., Gudejko-Thiel, J., Beitz, S., Schneider, E., Walenta, S., Peters, K., Unger, R., and Thuroff, J.W. 2008. Migration of renal carcinoma cells is dependent on protein kinase Cdelta via beta1 integrin and focal adhesion kinase. Int J Oncol 32: 1125-1131.

323. Villar, J., Arenas, M.I., MacCarthy, C.M., Blanquez, M.J., Tirado, O.M., and Notario, V. 2007. PCPH/ENTPD5 expression enhances the invasiveness of human prostate cancer cells by a protein kinase C delta-dependent mechanism. Cancer Res 67: 10859-10868.

324. Kho, D.H., Bae, J.A., Lee, J.H., Cho, H.J., Cho, S.H., Lee, J.H., Seo, Y.W., Ahn, K.Y., Chung, I.J., and Kim, K.K. 2008. KITENIN recruits Dishevelled/PKC{delta} to form a functional complex and controls the migration and invasiveness of colorectal cancer cells. Gut 58: 509-19.

325. Lin, C.W., Hou, W.C., Shen, S.C., Juan, S.H., Ko, C.H., Wang, L.M., and Chen, Y.C. 2008. Quercetin inhibition of tumor invasion via suppressing PKC delta/ERK/AP-1-dependent matrix metalloproteinase-9 activation in breast carcinoma cells. Carcinogenesis 29: 1807-1815.

326. Jane, E.P., Premkumar, D.R., and Pollack, I.F. 2006. Coadministration of sorafenib with rottlerin potently inhibits cell proliferation and migration in human malignant glioma cells. J Pharmacol Exp Ther 319: 1070-1080.

327. Kruger, J.S., and Reddy, K.B. 2003. Distinct mechanisms mediate the initial and sustained phases of cell migration in epidermal growth factor receptor-overexpressing cells. Mol Cancer Res 1: 801-809.

328. Leng, L., Du, B., Consigli, S., and McCaffrey, T.A. 1996. Translocation of protein kinase C-delta by PDGF in cultured vascular smooth muscle cells: inhibition by TGF-beta 1. Artery 22: 140-154.

215

329. Li, C., Wernig, F., Leitges, M., Hu, Y., and Xu, Q. 2003. Mechanical stress-activated PKCdelta regulates smooth muscle cell migration. FASEB J 17: 2106-2108.

330. Disatnik, M.H., Boutet, S.C., Lee, C.H., Mochly-Rosen, D., and Rando, T.A. 2002. Sequential activation of individual PKC isozymes in integrin-mediated muscle cell spreading: a role for MARCKS in an integrin signaling pathway. J Cell Sci 115: 2151-2163.

331. Iwabu, A., Smith, K., Allen, F.D., Lauffenburger, D.A., and Wells, A. 2004. Epidermal growth factor induces fibroblast contractility and motility via a protein kinase C delta-dependent pathway. J Biol Chem 279: 14551-14560.

332. Fan, J., Guan, S., Cheng, C.F., Cho, M., Fields, J.W., Chen, M., Denning, M.F., Woodley, D.T., and Li, W. 2006. PKCdelta clustering at the leading edge and mediating growth factor-enhanced, but not ecm-initiated, dermal fibroblast migration. J Invest Dermatol 126: 1233-1243.

333. Hsieh, H.L., Wu, C.Y., and Yang, C.M. 2008. Bradykinin induces matrix metalloproteinase-9 expression and cell migration through a PKC-delta-dependent ERK/Elk-1 pathway in astrocytes. Glia 56: 619-632.

334. Renault-Mihara, F., Beuvon, F., Iturrioz, X., Canton, B., De Bouard, S., Leonard, N., Mouhamad, S., Sharif, A., Ramos, J.W., Junier, M.P., et al. 2006. Phosphoprotein enriched in astrocytes-15 kDa expression inhibits astrocyte migration by a protein kinase C delta-dependent mechanism. Mol Biol Cell 17: 5141-5152.

335. Langlois, A., Chouinard, F., Flamand, N., Ferland, C., Rola-Pleszczynski, M., and Laviolette, M. 2009. Crucial implication of protein kinase C (PKC)-{delta}, PKC-{zeta}, ERK-1/2, and p38 MAPK in migration of human asthmatic eosinophils. J Leukoc Biol85: 656-63.

336. Gorshkova, I., He, D., Berdyshev, E., Usatuyk, P., Burns, M., Kalari, S., Zhao, Y., Pendyala, S., Garcia, J.G., Pyne, N.J., et al. 2008. Protein kinase C-epsilon regulates sphingosine 1-phosphate-mediated migration of human lung endothelial cells through activation of phospholipase D2, protein kinase C-zeta, and Rac1. J Biol Chem 283: 11794-11806.

337. Harrington, E.O., Doyle, K.E., Brunelle, J.L., and Ware, J.A. 2000. Endothelial proliferation, migration, and differentiation are blunted by conditionally expressed protein kinase C pseudosubstrate peptides. Biochem Biophys Res Commun 271: 499-508.

338. Carpenter, A.C., and Alexander, J.S. 2008. Endothelial PKC delta activation attenuates neutrophil transendothelial migration. Inflamm Res 57: 216-229.

339. Chaudhuri, P., Colles, S.M., Fox, P.L., and Graham, L.M. 2005. Protein kinase Cdelta-dependent phosphorylation of syndecan-4 regulates cell migration. Circ Res 97: 674-681.

216

340. Hirai, T., and Chida, K. 2003. Protein kinase Czeta (PKCzeta): activation mechanisms and cellular functions. J Biochem (Tokyo) 133: 1-7.

341. Ways, D.K., Cook, P.P., Webster, C., and Parker, P.J. 1992. Effect of Phorbol Esters on Protein Kinase-C-Zeta. J Biol Chem 267: 4799-4805.

342. Qiu, R.G., Abo, A., and Steven Martin, G. 2000. A human homolog of the C. elegans polarity determinant Par-6 links Rac and Cdc42 to PKCzeta signaling and cell transformation. Curr Biol 10: 697-707.

343. Wooten, M.W., Vandenplas, M.L., Seibenhener, M.L., Geetha, T., and Diaz-Meco, M.T. 2001. Nerve growth factor stimulates multisite tyrosine phosphorylation and activation of the atypical protein kinase C's via a src kinase pathway. Mol Cell Biol 21: 8414-8427.

344. Regala, R.P., Weems, C., Jamieson, L., Copland, J.A., Thompson, E.A., and Fields, A.P. 2005. Atypical protein kinase Ciota plays a critical role in human lung cancer cell growth and tumorigenicity. J Biol Chem 280: 31109-31115.

345. Regala, R.P., Weems, C., Jamieson, L., Khoor, A., Edell, E.S., Lohse, C.M., and Fields, A.P. 2005. Atypical protein kinase C iota is an oncogene in human non-small cell lung cancer. Cancer Res 65: 8905-8911.

346. Forde, S., Tye, B.J., Newey, S.E., Roubelakis, M., Smythe, J., McGuckin, C.P., Pettengell, R., and Watt, S.M. 2007. Endolyn (CD164) modulates the CXCL12-mediated migration of umbilical cord blood CD133+ cells. Blood 109: 1825-1833.

347. Petit, I., Goichberg, P., Spiegel, A., Peled, A., Brodie, C., Seger, R., Nagler, A., Alon, R., and Lapidot, T. 2005. Atypical PKC-zeta regulates SDF-1-mediated migration and development of human CD34+ progenitor cells. J Clin Invest 115: 168-176.

348. Guo, H., Ma, Y., Zhang, B., Sun, B., Niu, R., Ying, G., and Zhang, N. 2009. Pivotal Advance: PKC{zeta} is required for migration of macrophages. J Leukoc Biol 85: 911-8.

349. Kuribayashi, K., Nakamura, K., Tanaka, M., Sato, T., Kato, J., Sasaki, K., Takimoto, R., Kogawa, K., Terui, T., Takayama, T., et al. 2007. Essential role of protein kinase C zeta in transducing a motility signal induced by superoxide and a chemotactic peptide, fMLP. J Cell Biol 176: 1049-1060.

350. Sun, R., Gao, P., Chen, L., Ma, D., Wang, J., Oppenheim, J.J., and Zhang, N. 2005. Protein kinase C zeta is required for epidermal growth factor-induced chemotaxis of human breast cancer cells. Cancer Res 65: 1433-1441.

351. Laudanna, C., Sorio, C., Tecchio, C., Butcher, E.C., Bonora, A., Bassi, C., and Scarpa, A. 2003. Motility analysis of pancreatic adenocarcinoma cells reveals a role for the atypical zeta isoform of protein kinase C in cancer cell movement. Lab Invest 83: 1155-1163.

352. Xiao, A., Yin, C., Yang, C., Di Cristofano, A., Pandolfi, P.P., and Van Dyke, T. 2005. Somatic induction of Pten loss in a preclinical astrocytoma model reveals major roles in

217

disease progression and avenues for target discovery and validation. Cancer Res 65: 5172-5180.

353. Kanmogne, G.D., Schall, K., Leibhart, J., Knipe, B., Gendelman, H.E., and Persidsky, Y. 2007. HIV-1 gp120 compromises blood-brain barrier integrity and enhances monocyte migration across blood-brain barrier: implication for viral neuropathogenesis. J Cereb Blood Flow Metab 27: 123-134.

354. Loitto, V.M., Huang, C., Sigal, Y.J., and Jacobson, K. 2007. Filopodia are induced by aquaporin-9 expression. Exp Cell Res 313: 1295-1306.

355. Triplett, J.W., O'Riley, R., Tekulve, K., Norvell, S.M., and Pavalko, F.M. 2007. Mechanical loading by fluid shear stress enhances IGF-1 receptor signaling in osteoblasts in a PKCzeta-dependent manner. Mol Cell Biomech 4: 13-25.

356. Furlong, F., Crean, J., Thornton, L., O'Leary, R., Murphy, M., and Martin, F. 2007. Dysregulated intracellular signaling impairs CTGF-stimulated responses in human mesangial cells exposed to high extracellular glucose. Am J Physiol Renal Physiol 292: F1691-1700.

357. Crean, J.K., Furlong, F., Finlay, D., Mitchell, D., Murphy, M., Conway, B., Brady, H.R., Godson, C., and Martin, F. 2004. Connective tissue growth factor [CTGF]/CCN2 stimulates mesangial cell migration through integrated dissolution of focal adhesion complexes and activation of cell polarization. FASEB J 18: 1541-1543.

358. Urtreger, A.J., Grossoni, V.C., Falbo, K.B., Kazanietz, M.G., and Bal de Kier Joffe, E.D. 2005. Atypical protein kinase C-zeta modulates clonogenicity, motility, and secretion of proteolytic enzymes in murine mammary cells. Mol Carcinog 42: 29-39.

359. Huang, H.C., Huang, C.Y., Lin-Shiau, S.Y., and Lin, J.K. 2008. Ursolic acid inhibits IL-1beta or TNF-alpha-induced C6 glioma invasion through suppressing the association ZIP/p62 with PKC-zeta and downregulating the MMP-9 expression. Mol Carcinog 48: 517-531.

360. Powell, C.T., Fair, W.R., and Heston, W.D. 1994. Differential expression of protein kinase C isozyme messenger RNAs in dunning R-3327 rat prostatic tumors. Cell Growth Differ 5: 143-149.

361. Powell, C.T., Gschwend, J.E., Fair, W.R., Brittis, N.J., Stec, D., and Huryk, R. 1996. Overexpression of protein kinase C-zeta (PKC-zeta) inhibits invasive and metastatic abilities of Dunning R-3327 MAT-LyLu rat prostate cancer cells. Cancer Res 56: 4137-4141.

362. Vertommen, D., Rider, M., Ni, Y., Waelkens, E., Merlevede, W., Vandenheede, J.R., and Van Lint, J. 2000. Regulation of protein kinase D by multisite phosphorylation. Identification of phosphorylation sites by mass spectrometry and characterization by site-directed mutagenesis. J Biol Chem 275: 19567-19576.

218

363. Bowden, E.T., Barth, M., Thomas, D., Glazer, R.I., Mueller, S.C. 1999. An invasion-related complex of cortactin, paxillin, and PKCµ associates with invadopodia at sites of extracellular matrix degradation. Oncogene 18: 4440-4449.

364. Sacco, O., Silvestri, M., Sabatini, F., Sale, R., Defilippi, A.C., and Rossi, G.A. 2004. Epithelial cells and fibroblasts: structural repair and remodelling in the airways. PaediatrRespir Rev 5 Suppl A: S35-40.

365. Thompson, A.B., Robbins, R.A., Romberger, D.J., Sisson, J.H., Spurzem, J.R., Teschler, H., and Rennard, S.I. 1995. Immunological functions of the pulmonary epithelium. Eur Respir J 8: 127-149.

366. Liu, M. 2001. Alveolar epithelium in host defense: cytokine production. In Sepsis and Organ Dysfunction: From chaos to rationale. G.B. A. E. Baue, A. Gullo, J.-L. Vincent, editor. Milano: Springer. 37-50.

367. Montefort, S., and Holgate, S.T. 1991. Adhesion molecules and their role in inflammation. Respir Med 85:91-99.

368. Dufour, S., Duband, J.L., and Thiery, J.P. 1986. Role of a major cell-substratum adhesion system in cell behavior and morphogenesis. Biol Cell 58: 1-13.

369. Marx, J. 2006. Cell biology. Podosomes and invadopodia help mobile cells step lively. Science 312: 1868-1869.

370. Calle, Y., Carragher, N.O., Thrasher, A.J., and Jones, G.E. 2006. Inhibition of calpain stabilises podosomes and impairs dendritic cell motility. J Cell Sci 119: 2375-2385.

371. West, M.A., Wallin, R.P., Matthews, S.P., Svensson, H.G., Zaru, R., Ljunggren, H.G., Prescott, A.R., and Watts, C. 2004. Enhanced dendritic cell antigen capture via toll-like receptor-induced actin remodeling. Science 305: 1153-1157.

372. Krzewski, K., Chen, X., Orange, J.S., and Strominger, J.L. 2006. Formation of a WIP-, WASp-, actin-, and myosin IIA-containing multiprotein complex in activated NK cells and its alteration by KIR inhibitory signaling. J Cell Biol 173: 121-132.

373. Bruzzaniti, A., Neff, L., Sanjay, A., Horne, W.C., De Camilli, P., and Baron, R. 2005. Dynamin forms a Src kinase-sensitive complex with Cbl and regulates podosomes and osteoclast activity. Mol Biol Cell 16: 3301-3313.

374. Tehrani, S., Faccio, R., Chandrasekar, I., Ross, F.P., and Cooper, J.A. 2006. Cortactin has an essential and specific role in osteoclast actin assembly. Mol Biol Cell 17: 2882-2895.

375. Amadio, M., Battaini, F., and Pascale, A. 2006. The different facets of protein kinases C: old and new players in neuronal signal transduction pathways. Pharmacol Res 54: 317-325.

219

376. Tatin, F., Varon, C., Genot, E., and Moreau, V. 2006. A signalling cascade involving PKC, Src and Cdc42 regulates podosome assembly in cultured endothelial cells in response to phorbol ester. J Cell Sci 119: 769-781.

377. Gatesman, A., Walker, V.G., Baisden, J.M., Weed, S.A., and Flynn, D.C. 2004. Protein kinase Calpha activates c-Src and induces podosome formation via AFAP-110. Mol Cell Biol 24: 7578-7597.

378. Dwyer-Nield, L.D., Miller, A.C., Neighbors, B.W., Dinsdale, D., and Malkinson, A.M. 1996. Cytoskeletal architecture in mouse lung epithelial cells is regulated by protein-kinase C-alpha and calpain II. Am J Physiol 270: L526-534.

379. Han, B., Mura, M., Andrade, C.F., Okutani, D., Lodyga, M., Dos Santos, C.C., Keshavjee, S., Matthay, M., and Liu, M. 2005. TNF{alpha}-Induced Long Pentraxin PTX3 Expression in Human Lung Epithelial Cells via JNK. J Immunol 175: 8303-8311.

380. Xu, J., Bai, X.H., Lodyga, M., Han, B., Xiao, H., Keshavjee, S., Hu, J., Zhang, H., Yang, B.B., and Liu, M. 2007. XB130, A novel adaptor protein for signal transduction. J Biol Chem 282: 16401-16412. .

381. Lodyga, M., De Falco, V., Bai, X.H., Kapus, A., Melillo, R.M., Santoro, M., and Liu, M. 2009. XB130, a tissue-specific adaptor protein that couples the RET/PTC oncogenic kinase to PI 3-kinase pathway. Oncogene 28: 937-949.

382. Xu, J., Bai, X.H., Lodyga, M., Han, B., Xiao, H., Keshavjee, S., Hu, J., Zhang, H., Yang, B.B., and Liu, M. 2007. XB130, a novel adaptor protein for signal transduction. J Biol Chem 282: 16401-16412.

383. Webb, B.A., Eves, R., Crawley, S.W., Zhou, S., Cote, G.P., and Mak, A.S. 2005. PAK1 induces podosome formation in A7r5 vascular smooth muscle cells in a PAK-interacting exchange factor-dependent manner. Am J Physiol Cell Physiol 289: C898-907.

384. Han, B., Bai, X.H., Lodyga, M., Xu, J., Yang, B.B., Keshavjee, S., Post, M., and Liu, M. 2004. Conversion of mechanical force into biochemical signaling. J Biol Chem 279: 54793-54801.

385. Webb, B.A., Jia, L., Eves, R., and Mak, A.S. 2007. Dissecting the functional domain requirements of cortactin in invadopodia formation. Eur J Cell Biol 86: 189-206.

386. Kaneda, H., Waddell, T.K., de Perrot, M., Bai, X.H., Gutierrez, C., Arenovich, T., Chaparro, C., Liu, M., and Keshavjee, S. 2006. Pre-implantation multiple cytokine mRNA expression analysis of donor lung grafts predicts survival after lung transplantation in humans. Am J Transplant 6: 544-551.

387. Fischer, S., Hopkinson, D., Liu, M., MacLean, A.A., Edwards, V., Cutz, E., and Keshavjee, S. 2001. Raffinose improves 24-hour lung preservation in low potassium dextran glucose solution: a histologic and ultrastructural analysis. Ann Thorac Surg 71: 1140-1145.

220

388. Tang, P.S., Mura, M., Seth, R., and Liu, M. 2008. Acute lung injury and cell death: how many ways can cells die? Am J Physiol Lung Cell Mol Physiol 294: L632-641.

389. McDowell, E.M., Becci, P.J., Schurch, W., and Trump, B.F. 1979. The respiratory epithelium. VII. Epidermoid metaplasia of hamster tracheal epithelium during regeneration following mechanical injury. J Natl Cancer Inst 62: 995-1008.

390. Nikula, K.J., Wilson, D.W., Giri, S.N., Plopper, C.G., and Dungworth, D.L. 1988. The response of the rat tracheal epithelium to ozone exposure. Injury, adaptation, and repair. Am J Pathol 131: 373-384.

391. Higgs, H.N., and Pollard, T.D. 1999. Regulation of actin polymerization by Arp2/3 complex and WASp/Scar proteins. J Biol Chem 274: 32531-32534.

392. Castellano, F., Le Clainche, C., Patin, D., Carlier, M.F., and Chavrier, P. 2001. A WASp-VASP complex regulates actin polymerization at the plasma membrane. EMBO J 20: 5603-5614.

393. Chen, W.T. 1989. Proteolytic activity of specialized surface protrusions formed at rosette contact sites of transformed cells. J Exp Zool 251: 167-185.

394. Han, B., Lodyga, M., and Liu, M. 2005. Ventilator-induced lung injury: role of protein-protein interaction in mechanosensation. Proc Am Thorac Soc 2: 181-187.

395. Okutani, D., Lodyga, M., Han, B., and Liu, M. 2006. Src protein tyrosine kinase family and acute inflammatory responses. Am J Physiol Lung Cell Mol Physiol 291: L129-141.

396. Atkinson, J.J., Toennies, H.M., Holmbeck, K., and Senior, R.M. 2007. Membrane type 1 matrix metalloproteinase is necessary for distal airway epithelial repair and keratinocyte growth factor receptor expression after acute injury. Am J Physiol Lung Cell Mol Physiol293: L600-610.

397. Chen, W.T., and Wang, J.Y. 1999. Specialized surface protrusions of invasive cells, invadopodia and lamellipodia, have differential MT1-MMP, MMP-2, and TIMP-2 localization. Ann N Y Acad Sci 878: 361-371.

398. Schliwa, M., Nakamura, T., Porter, K.R., and Euteneuer, U. 1984. A tumor promoter induces rapid and coordinated reorganization of actin and vinculin in cultured cells. JCell Biol 99: 1045-1059.

399. Isowa, N., Xavier, A.M., Dziak, E., Opas, M., McRitchie, D.I., Slutsky, A.S., Keshavjee, S.H., and Liu, M. 1999. LPS-induced depolymerization of cytoskeleton and its role in TNF-alpha production by rat pneumocytes. Am J Physiol 277: L606-615.

400. dos Santos, C.C., Han, B., Andrade, C.F., Bai, X., Uhlig, S., Hubmayr, R., Tsang, M., Lodyga, M., Keshavjee, S., Slutsky, A.S., et al. 2004. DNA microarray analysis of gene expression in alveolar epithelial cells in response to TNFalpha, LPS, and cyclic stretch. Physiol Genomics 19: 331-342.

221

401. Tang, P.S., Tsang, M.E., Lodyga, M., Bai, X.H., Miller, A., Han, B., and Liu, M. 2006. Lipopolysaccharide accelerates caspase-independent but cathepsin B-dependent death of human lung epithelial cells. J Cell Physiol 209: 457-467.

402. Isowa, N., and Liu, M. 2001. Role of LPS-induced microfilament depolymerization in MIP-2 production from rat pneumocytes. Am J Physiol Lung Cell Mol Physiol 280: L762-770.

403. Tarone, G., Cirillo, D., Giancotti, F.G., Comoglio, P.M., and Marchisio, P.C. 1985. Rous sarcoma virus-transformed fibroblasts adhere primarily at discrete protrusions of the ventral membrane called podosomes. Exp Cell Res 159: 141-157.

404. Zhu, T., Goh, E.L., Graichen, R., Ling, L., and Lobie, P.E. 2001. Signal transduction via the growth hormone receptor. Cell Signal 13: 599-616.

405. Hai, C.M., Hahne, P., Harrington, E.O., and Gimona, M. 2002. Conventional protein kinase C mediates phorbol-dibutyrate-induced cytoskeletal remodeling in a7r5 smooth muscle cells. Exp Cell Res 280: 64-74.

406. Steinberg, S.F. 2004. Distinctive activation mechanisms and functions for protein kinase Cdelta. Biochem J 384: 449-459.

407. Schechtman, D., and Mochly-Rosen, D. 2001. Adaptor proteins in protein kinase C-mediated signal transduction. Oncogene 20: 6339-6347.

408. Hamplova, B., Novakova, O., Tvrzicka, E., Kolar, F., and Novak, F. 2005. Protein kinase C activity and isoform expression during early postnatal development of rat myocardium. Cell Biochem Biophys 43: 105-117.

409. Gangopadhyay, S.S., Takizawa, N., Gallant, C., Barber, A.L., Je, H.D., Smith, T.C., Luna, E.J., and Morgan, K.G. 2004. Smooth muscle archvillin: a novel regulator of signaling and contractility in vascular smooth muscle. J Cell Sci 117: 5043-5057.

410. Teti, A., Colucci, S., Grano, M., Argentino, L., and Zambonin Zallone, A. 1992. Protein kinase C affects microfilaments, bone resorption, and [Ca2+]o sensing in cultured osteoclasts. Am J Physiol 263: C130-139.

411. Zhou, S., Webb, B.A., Eves, R., and Mak, A.S. 2006. Effects of tyrosine phosphorylation of cortactin on podosome formation in A7r5 vascular smooth muscle cells. Am J Physiol Cell Physiol 290: C463-471.

412. Wilkinson, S.E., Parker, P.J., and Nixon, J.S. 1993. Isoenzyme specificity of bisindolylmaleimides, selective inhibitors of protein kinase C. Biochem J 294 ( Pt 2): 335-337.

413. Martiny-Baron, G., Kazanietz, M.G., Mischak, H., Blumberg, P.M., Kochs, G., Hug, H., Marme, D., and Schachtele, C. 1993. Selective inhibition of protein kinase C isozymes by the indolocarbazole Go 6976. J Biol Chem 268: 9194-9197.

222

414. Gschwendt, M., Muller, H.J., Kielbassa, K., Zang, R., Kittstein, W., Rincke, G., and Marks, F. 1994. Rottlerin, a novel protein kinase inhibitor. Biochem Biophys Res Commun 199: 93-98.

415. Bain, J., Plater, L., Elliott, M., Shpiro, N., Hastie, C.J., McLauchlan, H., Klevernic, I., Arthur, J.S., Alessi, D.R., and Cohen, P. 2007. The selectivity of protein kinase inhibitors: a further update. Biochem J 408: 297-315.

416. Liu, X.J., Yang, C., Gupta, N., Zuo, J., Chang, Y.S., and Fang, F.D. 2007. Protein kinase C-zeta regulation of GLUT4 translocation through actin remodeling in CHO cells. J Mol Med 85: 851-861.

417. Liu, M., Xu, J., Liu, J., Kraw, M.E., Tanswell, A.K., and Post, M. 1995. Mechanical strain-enhanced fetal lung cell proliferation is mediated by phospholipase C and D and protein kinase C. Am J Physiol 268: L729-738.

418. Keenan, C., and Kelleher, D. 1998. Protein kinase C and the cytoskeleton. Cell Signal 10: 225-232.

419. West, M.A., Prescott, A.R., Chan, K.M., Zhou, Z., Rose-John, S., Scheller, J., and Watts, C. 2008. TLR ligand-induced podosome disassembly in dendritic cells is ADAM17 dependent. J Cell Biol 182: 993-1005.

420. Han, Y.P., Yan, C., Zhou, L., Qin, L., and Tsukamoto, H. 2007. A matrix metalloproteinase-9 activation cascade by hepatic stellate cells in trans-differentiation in the three-dimensional extracellular matrix. J Biol Chem 282: 12928-12939.

421. Toth, M., Chvyrkova, I., Bernardo, M.M., Hernandez-Barrantes, S., and Fridman, R. 2003. Pro-MMP-9 activation by the MT1-MMP/MMP-2 axis and MMP-3: role of TIMP-2 and plasma membranes. Biochem Biophys Res Commun 308: 386-395.

422. Billottet, C., Rottiers, P., Tatin, F., Varon, C., Reuzeau, E., Maitre, J.L., Saltel, F., Moreau, V., and Genot, E. 2008. Regulatory signals for endothelial podosome formation. Eur J Cell Biol 87: 543-554.

423. Xiao, H., Bai, X., Lu, W., Kapus, A., Mak, A.S., and Liu, M. 2009. PKC regulates recruitment, release and activation of MMP-9 at podosomes. Mol Biol Cell: Under revision.

424. Burgstaller, G., and Gimona, M. 2005. Podosome-mediated matrix resorption and cell motility in vascular smooth muscle cells. Am J Physiol Heart Circ Physiol 288: H3001-3005.

425. Guegan, F., Tatin, F., Leste-Lasserre, T., Drutel, G., Genot, E., and Moreau, V. 2008. p190B RhoGAP regulates endothelial-cell-associated proteolysis through MT1-MMP and MMP2. J Cell Sci 121: 2054-2061.

223

426. Mukhopadhyay, U.K., Eves, R., Jia, L., Mooney, P., and Mak, A.S. 2009. p53 suppresses Src-induced podosome and rosette formation and cellular invasiveness through the upregulation of caldesmon. Mol Cell Biol 29: 3088-3098.

427. Takenawa, T., and Itoh, T. 2006. Membrane targeting and remodeling through phosphoinositide-binding domains. IUBMB Life 58: 296-303.

428. Yu, L., Qi, M., Sheff, M.A., and Elion, E.A. 2008. Counteractive control of polarized morphogenesis during mating by mitogen-activated protein kinase Fus3 and G1 cyclin-dependent kinase. Mol Biol Cell 19: 1739-1752.

429. Dow, L.E., Elsum, I.A., King, C.L., Kinross, K.M., Richardson, H.E., and Humbert, P.O. 2008. Loss of human Scribble cooperates with H-Ras to promote cell invasion through deregulation of MAPK signalling. Oncogene 27: 5988-6001.

430. Furmaniak-Kazmierczak, E., Crawley, S.W., Carter, R.L., Maurice, D.H., and Cote, G.P. 2007. Formation of extracellular matrix-digesting invadopodia by primary aortic smooth muscle cells. Circ Res 100: 1328-1336.

431. Buschman, M.D., Bromann, P.A., Cejudo-Martin, P., Wen, F., Pass, I., and Courtneidge, S.A. 2009. The Novel Adaptor Protein Tks4 (SH3PXD2B) Is Required for Functional Podosome Formation. Mol Biol Cell 20: 1302-1311.

432. Xu, W., Harrison, S.C., and Eck, M.J. 1997. Three-dimensional structure of the tyrosine kinase c-Src. Nature 385: 595-602.

433. Mandal, S., Johnson, K.R., and Wheelock, M.J. 2008. TGF-beta induces formation of F-actin cores and matrix degradation in human breast cancer cells via distinct signaling pathways. Exp Cell Res 314: 3478-3493.

434. Hong, I.K., Jin, Y.J., Byun, H.J., Jeoung, D.I., Kim, Y.M., and Lee, H. 2006. Homophilic interactions of Tetraspanin CD151 up-regulate motility and matrix metalloproteinase-9 expression of human melanoma cells through adhesion-dependent c-Jun activation signaling pathways. J Biol Chem 281: 24279-24292.

435. Byun, H.J., Hong, I.K., Kim, E., Jin, Y.J., Jeoung, D.I., Hahn, J.H., Kim, Y.M., Park, S.H., and Lee, H. 2006. A splice variant of CD99 increases motility and MMP-9 expression of human breast cancer cells through the AKT-, ERK-, and JNK-dependent AP-1 activation signaling pathways. J Biol Chem 281: 34833-34847.

436. Dempsey, E.C., Cool, C.D., and Littler, C.M. 2007. Lung disease and PKCs. PharmacolRes 55: 545-559.

437. Xu, L., and Deng, X. 2006. Protein kinase Ciota promotes nicotine-induced migration and invasion of cancer cells via phosphorylation of micro- and m-calpains. J Biol Chem281: 4457-4466.

224

438. Xu, L., and Deng, X. 2004. Tobacco-specific nitrosamine 4-(methylnitrosamino)-1-(3-pyridyl)-1-butanone induces phosphorylation of mu- and m-calpain in association with increased secretion, cell migration, and invasion. J Biol Chem 279: 53683-53690.

439. Slager, R.E., Allen-Gipson, D.S., Sammut, A., Heires, A., DeVasure, J., Von Essen, S., Romberger, D.J., and Wyatt, T.A. 2007. Hog barn dust slows airway epithelial cell migration in vitro through a PKCalpha-dependent mechanism. Am J Physiol Lung Cell Mol Physiol 293: L1469-1474.

440. Rodriguez, E.M., Dunham, E.E., and Martin, G.S. 2009. Atypical protein kinase C activity is required for extracellular matrix degradation and invasion by Src-transformed cells. J Cell Physiol 221: 171-182.

441. Iyer, V., Pumiglia, K., and DiPersio, C.M. 2005. Alpha3beta1 integrin regulates MMP-9 mRNA stability in immortalized keratinocytes: a novel mechanism of integrin-mediated MMP gene expression. J Cell Sci 118: 1185-1195.

442. DiPersio, C.M., Shao, M., Di Costanzo, L., Kreidberg, J.A., and Hynes, R.O. 2000. Mouse keratinocytes immortalized with large T antigen acquire alpha3beta1 integrin-dependent secretion of MMP-9/gelatinase B. J Cell Sci 113 ( Pt 16): 2909-2921.

443. Harris, J.E., Fernandez-Vilaseca, M., Elkington, P.T., Horncastle, D.E., Graeber, M.B., and Friedland, J.S. 2007. IFNgamma synergizes with IL-1beta to up-regulate MMP-9 secretion in a cellular model of central nervous system tuberculosis. FASEB J 21:356-365.

444. Stuelten, C.H., DaCosta Byfield, S., Arany, P.R., Karpova, T.S., Stetler-Stevenson, W.G., and Roberts, A.B. 2005. Breast cancer cells induce stromal fibroblasts to express MMP-9 via secretion of TNF-alpha and TGF-beta. J Cell Sci 118: 2143-2153.

445. Turner, N.A., O'Regan, D.J., Ball, S.G., and Porter, K.E. 2005. Simvastatin inhibits MMP-9 secretion from human saphenous vein smooth muscle cells by inhibiting the RhoA/ROCK pathway and reducing MMP-9 mRNA levels. FASEB J 19: 804-806.

446. Sbai, O., Ferhat, L., Bernard, A., Gueye, Y., Ould-Yahoui, A., Thiolloy, S., Charrat, E., Charton, G., Tremblay, E., Risso, J.J., et al. 2008. Vesicular trafficking and secretion of matrix metalloproteinases-2, -9 and tissue inhibitor of metalloproteinases-1 in neuronal cells. Mol Cell Neurosci 39: 549-568.

447. Hsia, D.A., Mitra, S.K., Hauck, C.R., Streblow, D.N., Nelson, J.A., Ilic, D., Huang, S., Li, E., Nemerow, G.R., Leng, J., et al. 2003. Differential regulation of cell motility and invasion by FAK. J Cell Biol 160: 753-767.

448. Wang, I.C., Chen, Y.J., Hughes, D.E., Ackerson, T., Major, M.L., Kalinichenko, V.V., Costa, R.H., Raychaudhuri, P., Tyner, A.L., and Lau, L.F. 2008. FoxM1 Regulates Transcription of JNK1 to Promote the G1/S Transition and Tumor Cell Invasiveness. JBiol Chem 283: 20770-20778.

225

449. Radjabi, A.R., Sawada, K., Jagadeeswaran, S., Eichbichler, A., Kenny, H.A., Montag, A., Bruno, K., and Lengyel, E. 2008. Thrombin induces tumor invasion through the induction and association of matrix metalloproteinase-9 and beta1-integrin on the cell surface. JBiol Chem 283: 2822-2834.

450. Wang, X.Q., Sun, P., and Paller, A.S. 2003. Ganglioside GM3 inhibits matrix metalloproteinase-9 activation and disrupts its association with integrin. J Biol Chem 278: 25591-25599.

451. Kim, D.S., Jeon, O.H., Lee, H.D., Yoo, K.H., and Kim, D.S. 2008. Integrin alphavbeta3-mediated transcriptional regulation of TIMP-1 in a human ovarian cancer cell line. Biochem Biophys Res Commun 377: 479-483.

452. Mistry, N.N., Pollaro, J., Song, J., De Lin, M., and Johnson, G.A. 2008. Pulmonary perfusion imaging in the rodent lung using dynamic contrast-enhanced MRI. Magn Reson Med 59: 289-297.

453. Bullen, A. 2008. Microscopic imaging techniques for drug discovery. Nat Rev Drug Discov 7: 54-67.

Molecular Mechanism of Podosome Formation and Proteolytic ... · critical for podosome formation,...

Documents

Transcript of Molecular Mechanism of Podosome Formation and Proteolytic ... · critical for podosome formation,...