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
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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.
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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.
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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.
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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
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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
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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
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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
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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
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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
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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
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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
Extracellular matrix
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).
Extracellular matrix
Extracellular matrix
Extracellular matrix
Extracellular matrix
Extracellular matrix
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.
Extracellular matrix
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
Extracellular matrix
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.
Extracellular matrix
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.
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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.
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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).
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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.
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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).
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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).
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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
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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).
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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
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122
0
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PKC PKC PKC PKCµ
Pear
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s C
orre
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No TreatPDBu
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Pear
son'
s C
orre
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oeffi
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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
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% Cells Degrading Matrix
* * * *
*
AF-
actin
Mat
rixO
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10 µm
10 µm
10 µm
D
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DM
SO
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OB
IM I
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PTA
/AM
Gö6
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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
% Cells Degrading Matrix
Fig 7-4
** * *
F-ac
tinO
verla
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PKC siRNAControl siRNA PKC siRNAA
D % Cells with Podosomes
Con
trol
Con
trol s
iRN
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Con
trol s
iRN
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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
% Cells Degrading Matrix
No Treat PDBu
020406080
100
*
PI PS
F-ac
tinM
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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
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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
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yControl siRNA MMP-2 siRNA MMP-9 siRNA MMP-14 siRNA
F-ac
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mIg G MMP-9 AB
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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
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Fig 7-8
PKC -GFP Tks5-mCherry Overlay DIC
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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
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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
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Bu
(p)AktS473 (p)GSK3 S9
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a2
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a5p85 (p)PDK1 (p)Akt (p)GSK3
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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
Pearson's Correlation Coefficient
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
% Cells Degrade Matrix
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
% Cells Degrade Matrix
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
Pearson's Correlation Coefficient
* *
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
Pearson's Correlation Coefficient
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
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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
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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
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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
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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
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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
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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
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