Embed Size (px)
Citation preview
THE ROLE OF TYPE VIII COLLAGEN IN
VASCULAR OCCLUSIVE DISEASE
by
Ilkim Eser Adiguzel, Hons. B.Sc.
A thesis submitted in conformity with the requirements
for the degree of Doctor of Philosophy
Graduate Department of Laboratory Medicine and Pathobiology
and the Cardiovascular Sciences Collaborative Program
University of Toronto
© Copyright by Ilkim Eser Adiguzel 2009
ii
The role of type VIII collagen in vascular occlusive disease
Ilkim Eser Adiguzel Doctor of Philosophy, 2009
Department of Laboratory Medicine and Pathobiology Cardiovascular Sciences Collaborative Program
University of Toronto
Abstract
During atherosclerosis and restenosis, there is an extensive amount of collagen
synthesis and degradation. Changes in the types of collagen present can have profound
effects on vascular smooth muscle cell (SMC) proliferation and migration. Type VIII
collagen, which is normally present at low levels within the mature vascular system, is
greatly increased during atherogenesis. The central theme of this thesis is to determine
the role of type VIII collagen in the pathogenesis of atherosclerosis and restenosis.
In the first study, we demonstrated the importance of type VIII collagen in SMC
migration and proliferation. SMCs from type VIII collagen-deficient mice display
increased adhesion and decreased spreading, migration, and proliferation compared to
SMCs from wild-type mice. Treatment of SMCs from type VIII collagen-deficient mice
with exogenous type VIII collagen can rescue the defects.
In the second study, we determined that type VIII collagen exerts its effects
through regulation of MMP-2 expression. Type VIII collagen-deficient SMCs have
decreased levels of MMP-2 and are impaired in chemotaxis toward PDGF-BB and in
their ability to contract thick collagen gels. We found that decreasing endogenous MMP-
2 levels in normal SMCs or adding exogenous collagen to type VIII collagen-deficient
iii
SMCs is sufficient to recapitulate the type VIII collagen-deficient or wild-type SMC
phenotype, respectively.
In the third study, we investigated the contribution of type VIII collagen to intimal
hyperplasia after mechanical injury in the mouse. We found that type VIII collagen-
deficient mice display a 35% reduction in intimal hyperplasia and attenuated vessel
remodeling after femoral artery wire injury, establishing a role for type VIII collagen in
restenosis.
The results of the work presented in this thesis demonstrate that production of
type VIII collagen confers an SMC phenotype with a greater propencity for proliferation
and migration. These effects are in part mediated through regulation of MMP-2
expression and activation. We conclude that the increases in type VIII collagen
production that occur during atherosclerosis and restenosis contribute to the capacity of
SMCs to alter the existing extracellular matrix in a manner which permits enhanced
migration.
iv
Acknowledgements
The production of this thesis would not have been possible without the guidance, encouragement, and support of many people. I would like to take this opportunity to acknowledge these individuals for their contributions during my Ph.D. candidacy. First and foremost, I would like to thank my supervisor, Dr. Michelle Bendeck, for her guidance and support throughout my studies. Much of my success can be attributed to Dr. Bendeck’s contribution to my Ph.D. training. Not only did she encourage me to think independently, have confidence in my abilities, and provide critical feedback on my work, she also offered her friendship, by engaging in stimulating conversations and various outdoor activities with myself and our labmates. I would also like to sincerely thank my labmates and colleagues for their support, critical feedback, and friendship over the years. In particular, I would like to thank Bernard, Chris, Cristina, Dan, Diane, Dorota, Karen, Katherine, Mike, Pam, Peter, Rosalind, Shathiyah, Tony, Winsion, and especially Guangpei for his neverending wisdom and zany ideas. I would also like to acknowledge the members of my Program Advisory Committee and/or Oral Defense Committees: Dr. David Courtman, Dr. Tara Haas, Dr. Alek Hinek, the late Dr. Lowell Langille, Dr. Bradley Strauss, the late Dr. Wolfgang Vogel, and Dr. Michael Ward. I would like to thank my Program Advisory Committee members for guiding my progress during my studies. I would like to especially acknowledge Dr. Langille, for encouraging me to see the bigger picture, and Dr. Vogel, for often providing an alternative point of view to dilemmas encountered in my work. Most importantly, I am deeply indebted to my family for their unwavering support. I would like to thank Stefan for his constant encouragement, vibrant energy, infectious laughter, and vivacious spirit that helped me to really enjoy my work and life in general. I am most grateful to my parents, Emin and Firdevs, and brother, Aras, for pushing me to be the best I can and for their neverending support, encouragement, guidance, and, most importantly, patience. All of my achievements and success result from their love and dedication. George Bernard Shaw once said, “Life is not about finding yourself. Life is about creating yourself.” I would finally like to thank everyone above, (and countless unnamed friends, family, and professors), for helping create who I am today. Eser Adiguzel
v
Table of Contents
Abstract .............................................................................................................................. ii
Acknowledgements .......................................................................................................... iv
List of Tables and Figures ............................................................................................... ix
List of Abbreviations and Acronyms ............................................................................ xii
Chapter 1: Review of Literature .................................................................................... 1
Introduction ....................................................................................................................... 2
1.1 The cardiovascular system .................................................................................... 3 1.1.1 Arterial anatomy and the extracellular matrix ..................................................... 3
1.2 Atherosclerosis and restenosis ............................................................................... 6 1.2.1 General overview ................................................................................................. 6 1.2.2 Animal models of atherosclerosis and restenosis ................................................ 9
1.2.2.1 Atherosclerosis ........................................................................................ 9 1.2.2.2 Restenosis .............................................................................................. 10
1.2.3 Matrix proteolysis in atherosclerosis and restenosis ......................................... 11 1.2.3.1 Matrix metalloproteinases ..................................................................... 12 1.2.3.2 Matrix metalloproteinases and atherogenesis ........................................ 13
1.3 Mechanisms of cell migration .............................................................................. 16
1.4 Extracellular matrix receptors involved in atherogenesis ................................ 19 1.4.1 Integrins ............................................................................................................. 19
1.4.1.1 Integrins and atherogenesis .................................................................... 20 1.4.2 Discoidin domain receptors................................................................................ 24
1.4.2.1 Discoidin domain receptors and atherogenesis ..................................... 25
1.5 Changes in the extracellular matrix during atherosclerosis and restenosis ... 27 1.5.1 Collagens and atherogenesis .............................................................................. 28 1.5.2 Glycoproteins, proteoglycans, and atherogenesis .............................................. 32 1.5.3 Elastin and atherogenesis ................................................................................... 35
1.6 Type VIII collagen ................................................................................................ 36 1.6.1 Structure and localization ................................................................................... 36 1.6.2 Functions of type VIII collagen ......................................................................... 38 1.6.3 Type VIII collagen in vascular disease .............................................................. 41
vi
1.7 Hypothesis and objectives .................................................................................... 46
1.8 Tables ..................................................................................................................... 48
Chapter 2: Contribution of type VIII collagen to smooth muscle cell migration and proliferation ............................................................................................................. 51
2.1 Introduction .......................................................................................................... 52
2.2 Materials and methods ......................................................................................... 53 2.2.1 Chemicals and reagents ...................................................................................... 53 2.2.2 Animals ............................................................................................................... 53 2.2.3 Cell culture ......................................................................................................... 54 2.2.4 Cell morphology ................................................................................................ 55 2.2.5 Immunocytochemistry ....................................................................................... 56 2.2.6 Adhesion assays ................................................................................................. 58 2.2.7 Spreading and migration assays ......................................................................... 58 2.2.8 Gelatin zymography ........................................................................................... 59 2.2.9 Type VIII collagen rescue experiments ............................................................. 59 2.2.10 Proliferation assays .......................................................................................... 60 2.2.11 Immunoblotting ................................................................................................ 61 2.2.12 Cell viability assays ......................................................................................... 61 2.2.13 Statistics ........................................................................................................... 62
2.3 Results ................................................................................................................... 62 2.3.1 COL8-/- smooth muscle cells are phenotypically distinct from COL8+/+ smooth muscle cells ..................................................................................................... 62 2.3.2 The production of type VIII collagen was upregulated after injury ................... 66 2.3.3 The production of type VIII collagen decreased the attachment of smooth muscle cells to type I collagen, and facilitated spreading and migration ..................... 67 2.3.4 Type VIII collagen production increases MMP activity .................................... 68 2.3.5 Type VIII collagen facilitates smooth muscle cell proliferation ........................ 69
2.4 Discussion .............................................................................................................. 69
2.5 Figures ................................................................................................................... 76
Chapter 3: Type VIII collagen-dependent regulation of smooth muscle cell MMP-2 production and migration ................................................................................ 89
3.1 Introduction .......................................................................................................... 90
3.2 Materials and methods ......................................................................................... 91 3.2.1 Chemicals and reagents ...................................................................................... 91 3.2.2 Aortic smooth muscle cells ................................................................................ 91 3.2.3 Transwell migration assays ................................................................................ 92 3.2.4 Gel contraction assays ........................................................................................ 93
vii
3.2.5 Quantitative real-time polymerase chain reaction (qRT-PCR) .......................... 94 3.2.6 SiRNA experiments ........................................................................................... 95 3.2.7 Time-lapse migration assays ............................................................................. 96 3.2.8 Immunoblotting .................................................................................................. 97 3.2.9 Gelatin zymography ........................................................................................... 98 3.2.10 Statistics ........................................................................................................... 98
3.3 Results ................................................................................................................... 98 3.3.1 There are decreased levels of MMP-2 mRNA, protein and activity in the COL8-/- smooth muscle cells ........................................................................................ 98 3.3.2 MMP-2 mRNA, protein, and activity was decreased after treatment with siRNA in COL8+/+ smooth muscle cells ...................................................................... 99 3.3.3 COL8-/- smooth muscle cells display impaired chemotaxis ............................. 100 3.3.4 MMP-2 knockdown impairs the chemotactic migration of smooth muscle cells ............................................................................................................................ 101 3.3.5 COL8-/- smooth muscle cell migration deficiencies are due to decreased MMP-2 ....................................................................................................................... 101 3.3.6 COL8-/- smooth muscle cells are significantly impaired in contracting thick collagen gels ............................................................................................................... 102
3.4 Discussion ............................................................................................................ 103
3.5 Figures ................................................................................................................. 110
Chapter 4: The contribution of type VIII collagen in response to wire injury of mouse arteries................................................................................................................ 120
4.1 Introduction ........................................................................................................ 121
4.2 Materials and methods ....................................................................................... 121 4.2.1 Chemicals and reagents .................................................................................... 121 4.2.2 Animals ............................................................................................................ 122 4.2.3 Carotid artery wire injury ................................................................................. 122 4.2.4 Femoral artery wire injury ............................................................................... 123 4.2.5 Carotid and femoral artery processing ............................................................. 123 4.2.6 Determination of the extent of denudation and re-endothelization .................. 124 4.2.7 Immunostaining for Ki67 ................................................................................. 125 4.2.8 Gelatin zymography ......................................................................................... 126 4.2.9 Immunoblotting ................................................................................................ 127 4.2.10 Intimal hyperplasia ......................................................................................... 127 4.2.11 Statistics ......................................................................................................... 128
4.3 Results ................................................................................................................. 128 4.3.1 Type VIII collagen was increased in injured carotid arteries of COL8+/+ mice ............................................................................................................................ 128 4.3.2 The extent of injury was the same in both COL8-/- and COL8+/+ mice ........... 129
viii
4.3.3 There were no significant differences in smooth muscle cell proliferation in injured carotid arteries between the COL8-/- and COL8+/+ mice ............................... 129 4.3.4 There were no significant differences between MMP and TIMP activity in injured carotid arteries from COL8-/- and COL8+/+ mice ........................................... 130 4.3.5 There were no significant differences in intimal hyperplasia in injured carotid arteries from COL8+/+ and COL8-/- mice ....................................................... 131 4.3.6 COL8-/- mice had increased medial proliferation after femoral artery wire injury .......................................................................................................................... 132 4.3.7 COL8-/- mice demonstrated reduced outward remodeling after femoral artery wire injury ........................................................................................................ 133
4.4 Discussion ............................................................................................................ 133
4.5 Figures and tables ............................................................................................... 142
Chapter 5: General discussion and future directions ............................................... 160
5.1 The effects of type VIII collagen on migration and proliferation .................. 161
5.2 Regulation of MMP-2 and migration by type VIII collagen .......................... 164
5.3 Contribution of type VIII collagen in the arterial response to mechanical injury .......................................................................................................................... 166
5.4 Conclusion ........................................................................................................... 170
5.5 Figures ................................................................................................................. 172
References ...................................................................................................................... 174
ix
List of Tables and Figures
Chapter 1: Review of Literature .................................................................................... 1 Table 1.8.1 Collagens in the vessel wall ..................................................................... 49 Table 1.8.2 MMPs in the vasculature ......................................................................... 50
Chapter 2: Contribution of type VIII collagen to smooth muscle cell migration and proliferation ..................................................................................................................... 51
Figure 2.5.1 COL8-/- smooth muscle cells do not produce type VIII collagen and are morphologically different from COL8+/+ smooth muscle cells .............................. 77 Figure 2.5.2 COL8-/- smooth muscle cells have increased prominent actin stress fibers compared to COL8+/+ smooth muscle cells, which are decreased in the presence of type VIII collagen ..................................................................................... 78 Figure 2.5.3 COL8-/- smooth muscle cells have a large stable microtubule network compared to COL8+/+ smooth muscle cells which is decreased in the presence of type VIII collagen ........................................................................................................ 79 Figure 2.5.4 COL8-/- smooth muscle cells contain more basal focal adhesions compared to COL8+/+ smooth muscle cells which are decreased in the presence of type VIII collagen ........................................................................................................ 80 Figure 2.5.5 COL8-/- smooth muscle cells revert to a size and shape similar to COL8+/+ smooth muscle cells in the presence of type VIII collagen ........................... 81 Figure 2.5.6 Type VIII collagen is upregulated in COL8+/+ smooth muscle cells after wounding ...................................................................................................................... 82 Figure 2.5.7 Type VIII collagen is deposited into the extracellular matrix ................ 83 Figure 2.5.8 COL8-/- smooth muscle cells display increased adhesion compared to COL8+/+ smooth muscle cells ....................................................................................... 84 Figure 2.5.9 COL8-/- smooth muscle cells are impaired in their ability to spread after plating .................................................................................................................. 85 Figure 2.5.10 Migration of COL8-/- smooth muscle cells is impaired compared to COL8+/+ smooth muscle cells ....................................................................................... 86 Figure 2.5.11 COL8-/- smooth muscle cells have less MMP-2 activity than COL8+/+ smooth muscle cells ....................................................................................... 87 Figure 2.5.12 COL8-/- smooth muscle cells proliferate less than COL8+/+ smooth muscle cells .................................................................................................................. 88
Chapter 3: Type VIII collagen-dependent regulation of smooth muscle cell MMP-2 production and migration .............................................................................................. 89
Figure 3.5.1 COL8-/- smooth muscle cells contain less MMP-2 mRNA than COL8+/+ smooth muscle cells ..................................................................................... 111 Figure 3.5.2 Levels of MMP-2 are effectively knocked-down after administration of MMP-2 siRNA ....................................................................................................... 112 Figure 3.5.3 COL8-/- smooth muscle cells display less chemotaxis towards PDGF-BB than COL8+/+ smooth muscle cells ........................................................... 113 Figure 3.5.4 MMP-2 siRNA inhibits chemotaxis in COL8+/+ smooth muscle cells . 114
x
Figure 3.5.5 MMP-2 siRNA reduces COL8+/+ migration levels to those of COL8-/- smooth muscle cells ...................................................................................... 115 Figure 3.5.6 MMP-2-/- smooth muscle cells display reduced rates of migration ...... 116 Figure 3.5.7 COL8-/- smooth muscle cells display attenuated collagen gel contraction compared to COL8+/+ smooth muscle cells ............................................. 117 Figure 3.5.8 In the 3-dimensional collagen gel assay, COL8-/- smooth muscle cells produce less MMP-2 and MMP-9 than COL8+/+ smooth muscle cells ...................... 118 Figure 3.5.9 MMP-2 siRNA reduces COL8+/+ gel contraction to that of COL8-/- smooth muscle cells ................................................................................................... 119
Chapter 4: The contribution of type VIII collagen in response to wire injury of mouse arteries................................................................................................................ 120
Figure 4.5.1 Type VIII collagen is increased in COL8+/+ mouse carotid arteries after wire injury .......................................................................................................... 143 Figure 4.5.2 Extent of injury is the same in COL8-/- and COL8+/+ mice ................. 144 Figure 4.5.3 Images of cross-sections from uninjured COL8-/- and COL8+/+ carotid arteries ............................................................................................................ 145 Figure 4.5.4 Images of cross-sections from COL8-/- and COL8+/+ carotid arteries four and seven days after injury ................................................................................. 146 Figure 4.5.5 There were no differences in proliferation or total cell number between COL8-/- and COL8+/+ mice four days after carotid artery wire injury ......... 147 Figure 4.5.6 There were no differences in proliferation or total cell number between COL8-/- and COL8+/+ mice seven days after carotid artery wire injury ...... 148 Figure 4.5.7 There were no differences in MMP or TIMP activity between COL8-/- and COL8+/+ mice after carotid artery injury ................................................ 149 Figure 4.5.8 There were no differences in gelatinase activity between COL8-/- and COL8+/+ mice after carotid artery injury ............................................................. 150 Figure 4.5.9 There were no significant differences in vessel wall hypertrophy after carotid artery injury between COL8-/- and COL8+/+ mice ................................. 151 Figure 4.5.10 There were no differences in lumen size or outward remodeling between COL8-/- and COL8+/+ mice after carotid artery injury ................................. 152 Figure 4.5.11 Images of cross-sections from COL8-/- and COL8+/+ carotid arteries twenty-one days after injury....................................................................................... 152 Figure 4.5.11 Images of cross-sections from COL8-/- and COL8+/+ carotid arteries twenty-one days after injury....................................................................................... 153 Figure 4.5.12 Images of cross-sections from uninjured and injured COL8-/- and COL8+/+ femoral arteries .......................................................................................... 154 Figure 4.5.13 COL8-/- mice demonstrated increased proliferation in the media seven days after femoral artery injury compared to COL8+/+ mice ........................... 155 Figure 4.5.14 There were no significant differences in vessel wall hypertrophy after femoral artery injury between COL8-/- and COL8+/+ mice ................................ 156 Figure 4.5.15 COL8-/- mice demonstrated attenuated outward remodeling after femoral injury ............................................................................................................. 157 Figure 4.5.16 Images of cross sections from COL8-/- and COL8+/+ femoral arteries twenty-one days after injury .......................................................................... 158 Table 4.5.1 Comparison of different mouse injury models of intimal hyperplasia .. 159
xi
Chapter 5: General discussion and future directions ............................................... 160 Figure 5.5.1 The role of type VIII collagen in smooth muscle cells in vascular occlusive disease ........................................................................................................ 173
xii
List of Abbreviations and Acronyms
AEC 3-amino-9-ethylcarbazole ApoE apolipoprotein E ANOVA analysis of variance ARP acidic ribosomal protein ATP adenosine triphosphate bFGF basic fibroblast growth factor BSA bovine serum albumin cDNA complimentary deoxyribonucleic acid cdk cyclin-dependent kinase CMHD Centre for Modeling Human Disease COL8+/+ wild-type, Col8a1+/+/Col8a2+/+
COL8-/- knockout, Col8a1-/-/Col8a2-/- CPM counts per minute Ct critical threshold Cy3 cyanine DDR discoidin domain receptor DMEM Dulbecco’s Modified Eagle’s Medium DNA deoxyribonucleic acid DTT dithiothreitol EDTA ethylene diamine tetraacetic acid EGTA ethylene glycol tetraacetic acid ERK extracellular related kinase FAK focal adhesion kinase FBS fetal bovine serum FITC fluorescein isothiocyanate FRNK focal adhesion kinase-related non-kinase GAP guanosine-5’-triphosphatase-activating protein GAPDH glyceraldehyde 3-phosphate dehydrogenase GASGER glycine-alanine-serine-glycine-glutamate-arginine GDP guanosine-5’-diphosphate GEF guanine-nucleotide exchange factor GFOGER glycine-phenylalanine-hydroxyproline-glycine-glutamate-arginine GLOGER glycine-leucine-hydroxyproline-glycine-glutamate-arginine GM-CSF granulocyte colony stimulating factor GTP guanosine-5’-triphosphate HBSS Hanks’ Balanced Salt Solution HEPES N-2-hydroxyethylpiperazone-n-2-ethanesulfonic acid IEL internal elastic lamina Klf4 Krüppel-like transcription factor-4 LDL low-density-lipoprotein LDLR low-density-lipoprotein receptor MAPK mitogen-activated protein kinase
xiii
M-CSF macrophage colony stimulating factor MEF mouse embryonic fibroblasts MLCP myosin light chain phosphatase MMP matrix metalloproteinase mRNA messenger ribonucleic acid MT-MMP membrane-type matrix metalloproteinase NC non-collagenous PAGE polyacrylamide gel electrophoresis PBS phosphate-buffered saline PCR polymerase chain reaction PDGF platelet-derived growth factor PI3K phosphatidylinositol 3-kinase PKC protein kinase C PMSF phenylmethanesulphonylfluoride qRT-PCR quantitative real-time polymerase chain reaction RGD arginine-glycine-aspartate RNA ribonucleic acid SDS sodium dodecyl sulphate SH2 src-homology-2 SHP-2 src-homology-2 containing protein tyrosine phosphatase siRNA small interfering RNA SMC smooth muscle cell TBS-T Tris-buffered saline containing Tween 20 TCA trichloroacetic acid TGF-β transforming growth factor β TIMP tissue inhibitor of matrix metalloproteinase TRITC tetramethyl rhodamine VEGF vascular endothelial growth factor
1
Chapter 1
Review of Literature
2
Introduction
Cardiovascular diseases, including occlusive vascular diseases such as
atherosclerosis and restenosis, are the leading cause of death in developed nations. These
diseases cause narrowing of vital blood vessels and leading to inadequate blood supply to
the heart and brain, resulting in myocardial and cerebral infarction. Remodeling of the
vascular extracellular matrix in these diseases contributes to the restriction of blood flow.
This process includes an increase in the synthesis of extracellular matrix proteins with a
disproportionate increase in matrix proteins not normally found within the vasculature,
such as type VIII collagen. Many cardiovascular research laboratories today analyze the
substances produced during atherosclerosis and restenosis to understand the mechanisms
behind these complex diseases. However, this observational approach has not yet led to
an understanding of the mechanisms by which these changes contribute to the
pathogenesis of disease. In the following review of literature, the vascular architecture is
described to give an overview of the cellular and extracellular matrix components present
and the cellular receptors that link the two. Atherosclerosis and restenosis are then
described, with attention given to the cellular receptors involved and the changes in
extracellular matrix that accompany these diseases. Particular attention is given to the
role of cellular motility within the vasculature, as this is a key process regulating disease
progression. Most notably, the emergence of type VIII collagen as an important
component of atherosclerosis and restenosis is discussed, as this is an understudied
molecule and its potential role in vascular disease is the central theme of this thesis.
3
1.1 The cardiovascular system
1.1.1 Arterial anatomy and the extracellular matrix
The circulatory system consists of the heart and an arterial and venous
vasculature. Arteries transport oxygenated and nutrient-rich blood away from the heart at
high pressure and branch into arterioles, which branch into capillaries within tissues to
supply them with oxygen and nutrients. De-oxygenated and nutrient-poor blood is then
drained from the capillaries into the venules after which it enters the veins, which
transport the blood back to the heart at low pressure. The exception to this is within the
pulmonary circulation where deoxygenated blood is carried into the lungs from the heart
via the pulmonary arteries. After oxygenation, the blood is returned via the pulmonary
veins to the heart and ready to enter the systemic circulation described above. For the
purpose of this thesis, we will be concentrating mainly on the arteries.
Arterial blood vessels consist of three distinct layers of cells embedded in a
surrounding extracellular matrix. The innermost intimal layer consists of a single layer
of endothelial cells resting on a basement membrane of type IV collagen along with
laminin and perlecan. Endothelial cells serve to provide an anti-thrombotic and non-
adhesive surface to blood and also regulate the passage of molecules from the
bloodstream into tissues and vice versa. The medial layer consists of concentric and
alternating layers of smooth muscle cells (SMCs) and elastic lamina with a matrix rich in
collagens, particularly fibrillar types I and III collagens, along with glycoproteins and the
proteoglycans hyaluronan, decorin, versican and perlecan. The contraction or relaxation
4
of smooth muscle cells in the media regulates the diameter of the artery. The outermost
adventitial layer contains fibroblasts and longitudinally-oriented type I and III collagens
and elastic fibers. The components of the vascular extracellular matrix, while providing
structural support for smooth muscle cells, also transmit mechanical and biochemical
stimuli to the cells, resulting in cell signaling which can in turn modulate cellular
responses and cause changes in most components of the extracellular matrix.
The most abundant matrix components of large arteries are comprised of
collagens, providing tensile strength, and elastin, providing elastic recoil. Collagens are
characterized by a triple helix consisting of either three identical or different α chains,
each coiled into a left-handed helix, which wrap around each other to form a right-handed
super triple helix. In order to form a helical structure, the α chains of collagen contain
multiple long repeats of a Gly-X-Y sequence, where X is usually proline and Y is usually
hydroxyproline. The multiple types of collagens are divided into two major groups based
on their ability to form macromolecular fibrils in culture, the fibrillar and nonfibrillar
collagens, which are then divided into further subgroups based on structural
characteristics (for reviews of different collagen structures, please see(van der and
Garrone, 1991; Prockop and Kivirikko, 1995). Many different collagen types are present
within the vessel wall (Table 1.8.1), with the fibrillar types I and III collagen representing
60% and 30%, respectively, of vascular collagens (vascular collagens reviewed
in(Mayne, 1986).
Elastic fibers within the vessel wall are composed of elastin and associated
microfibrils, and serve to provide elasticity to the vessel. Elastin is one of the most
abundant proteins in large arteries subject to variations in stress and blood pressure and
5
this elasticity is responsible for their resilience. Elastin is synthesized by smooth muscle
cells as a soluble monomer which is organized into insoluble polymers to form the
concentric elastic lamellae (for a review of the structure and formation of elastic fibers,
see(Rosenbloom et al., 1993). Elastin is also responsible for stabilization of developed
arteries as elastin-deficient mice die shortly after birth, due to uncontrolled proliferation
of smooth muscle cells leading to arterial thickening and occlusion (Li et al., 1998a).
Abnormal elastic fiber formation due to the loss of one elastin allele in humans and mice
demonstrated that both had thinner, yet increased number of elastic lamellae. In parallel
with the increased vessel wall thickness seen in elastin-deficient mice, both humans and
mice with loss of one elastin allele have increased proliferation of smooth muscle cells
leading to increased vessel wall thickness (Li et al., 1998b).
Proteoglycans are core proteins having one or more covalently bound
glycosaminoglycan chains, such as chondroitin sulfate and heparan sulfate. Within the
vasculature, these include the large aggregating aggrecan and versican, and the small
non-aggregating biglycan and decorin (Iozzo, 1998). Proteoglycans interact with other
extracellular matrix components through their protein and carbohydrate domains and also
function to regulate cellular responses due to their ability to sequester cytokines. The
glycoproteins of the vessel wall include fibronectin, vitronectin, laminin, and tenascin,
which have multidomain structures, enabling simultaneous interactions between cells and
other components of the extracellular matrix (Raines, 2000).
6
1.2 Atherosclerosis and restenosis
1.2.1 General overview
Atherosclerosis and restenosis are vascular diseases involving remodeling and a
change in cellular composition of the vasculature. Both result in a narrowing of the
arterial lumen due to expansion of either the atherosclerotic plaque or the restenotic
neointima, and constrictive remodeling. Both can also result in arterial occlusion and,
ultimately, death. Since type VIII collagen has been implicated in the progression of
atherosclerosis and restenosis (discussed in Section 1.6.3), the major goal of this thesis
was to determine its role in the molecular mechanisms underlying atherogenesis.
Atherosclerosis is a vascular disease affecting mainly large muscular and some
elastic arteries and involving chronic inflammation. It starts with an accumulation of
oxidized low-density-lipoproteins (LDL) below the endothelium, causing endothelial cell
dysfunction and increased adherence of monocytes, macrophages and T-lymphocytes,
followed by their subsequent penetration into the subendothelial layer, and further
accumulation of intracellular lipids (reviewed in(Hansson and Libby, 2006). Release of
various growth factors, such as platelet-derived growth factor (PDGF) and basic
fibroblast growth factor (bFGF) results in the medial smooth muscle cells switching from
a differentiated, contractile phenotype to a dedifferentiated, synthetic and proliferative
phenotype and migrating from the media into the intima (Campbell and Campbell, 1994a;
Campbell and Campbell, 1994b). Intermediate-sized lesions are formed by the
proliferation of layers of vascular smooth muscle cells and continued accumulation of
macrophages, with an extensive synthesis of extracellular matrix by the smooth muscle
7
cells. Advanced lesions, or plaques, generally consist of a lipid or necrotic core with a
fibrous cap, while the core and edges, or shoulders, of the lesion contain inflammatory
cells. Plaques consisting of many layers of smooth muscle cells and a thick fibrous cap
are stable plaques, yet can lead to occlusion if they intrude into the lumen. As plaque
growth encroaches on the lumen, the vessel may undergo dilation, or outward
remodeling, to compensate for about 30% of the lumen occlusion in an effort to maintain
lumen size. Although this can prevent luminal narrowing, it is maladaptive because it
can mask the presence of an unstable plaque (arterial remodeling reviewed in(Pasterkamp
et al., 2000). Unstable, or vulnerable plaques usually contain a large lipid core, a very
thin fibrous cap, and many inflammatory cells in the shoulder regions, which can lead to
plaque rupture due to macrophage matrix metalloproteinase (MMP) degradation of the
fibrous cap and subsequent thrombosis.
Substantiating the importance of hemodynamic forces within the vasculature,
atherosclerosis has a predilection to form at areas of disturbed blood flow, such as
bifurcations and curvatures within the vasculature (Asakura and Karino, 1990). Most
notably, plaque formation appears at areas where shear stress and flow are reduced or
where there is turbulent rather than laminar flow. This is due to a switch of the
endothelial cells lining the lumen from an atheroprotective phenotype to an atherogenic
phenotype in which they recruit and activate monocytes, increase platelet activation,
increase vasoconstriction, and increase endothelial cell apoptosis and turnover.
One clinical treatment for advanced atherosclerosis is angioplasty, in which a
deflated balloon is advanced into the lumen and inflated to increase the size of the lumen.
Although initially successful, this method often results in the development of restenosis.
8
Restenosis is an arterial remodeling response to the injury caused by angioplasty, and is
characterized by further intimal thickening and constrictive inward remodeling of the
vessel. Following angioplasty, shear stress decreases, inducing remodeling and leading
to narrowing of the lumen (Pasterkamp et al., 2000; Ward et al., 2000). The lumen
narrowing is due at first mainly to vasoconstriction by the vascular smooth muscle cells
and then later to the collagen-rich intimal accumulation (Clowes et al., 1983a; Lafont et
al., 1999). One remedy used to avoid vasoconstriction, and thus prevent restenosis, is the
use of stents in balloon angioplasty, which serve to keep the vessel open at a defined
diameter. However, vessels may still undergo luminal narrowing due to lesion
development and in-stent restenosis. Examination of follow-up reports of stented patients
for up to eleven years demonstrated a triphasic response to stenting consisting of
development of early in-stent restenosis up to 6 months, a regressional phase in lesion
growth from 6 months to three years, and a late, re-narrowing phase past four years after
the procedure (Kimura et al., 2002). Further research led to the development of drug-
eluting stents, which slowly release anti-proliferative and anti-migratory substances, such
as sirolimus and paclitaxel, which have demonstrated success in laboratory tests and
clinical trials at preventing or decreasing restenosis (Vaina et al., 2005). Long-term
studies showed that, while there were no differences in outcome or stent-thrombosis
between drug-eluting and bare metal stents 4 years after surgery, late stent-thrombosis in
drug-eluting stents is increased in the first year after placement. This was attributed to
delayed or incomplete endothelization or late polymer reactions (Stone et al., 2007),
indicating the need for further study in this field.
9
1.2.2 Animal models of atherosclerosis and restenosis
Most of our knowledge of the molecular mechanisms underlying atherosclerosis
and restenosis has come from studying animal models of atherogenesis. In the next two
sections, animal models commonly used to study atherogenesis are described,
highlighting what has been learned about the molecular mechanisms underlying the
smooth muscle cell responses.
1.2.2.1 Atherosclerosis
Historically, animal models used to study atherosclerosis have been rabbits, pigs,
dogs, and non-human primates fed a high-cholesterol diet to stimulate atherogenesis.
While similarities to the human disease are high, these models have their limitations due
to the difficulty of genetic manipulation and/or high cost. In the early 1990s, two mouse
models were developed that have a genetic predisposition to atherosclerosis. Due to the
relative low-cost, ease of genetic manipulation, and the rapid time course of disease
progression, the use of these mouse models have become commonplace for studying the
factors involved in atherosclerosis.
Both models are based on mechanisms of cholesterol uptake and clearance from
the circulation. ApolipoproteinE (ApoE), is a lipid-binding glycoprotein that mediates
the cellular uptake of cholesterol-rich particles in the liver. In 1992, two research groups
produced apoE-deficient mice, which develop spontaneous hypercholesterolemia and
atherosclerosis (Plump et al., 1992; Zhang et al., 1992). These mice develop lesions
similar to humans in that they are located in the same pre-disposed regions of the
vasculature and lesions progress from initial fatty streaks to advanced plaques with
necrotic cores (Reddick et al., 1994). The LDL receptor (LDLR) is a cell-surface
10
receptor involved in clearance of intermediate- and low-density lipoproteins from the
circulation. Generation of LDLR-deficient mice has provided another mouse model of
atherogenesis with one advantage over the apoE-deficient mouse model: the LDLR-
deficient mouse model will undergo a predictable and reproducible time-course of
atherogenesis only with administration of a high-fat diet, enabling control over the onset
of atherosclerosis. These mice also undergo atherogenesis in a manner similar to the
apoE-deficient mice (Ishibashi et al., 1994).
1.2.2.2 Restenosis
An animal model of restenosis has been established by denudation of the intimal
endothelial cell layer via balloon catheter injury in the rat or rabbit, where a catheter is
advanced through the external carotid artery into the common carotid and then inflated
and pulled along the length of the artery for a few passes (Clowes et al., 1983a). A
similar model is used in the mouse by advancing a copper wire into the common carotid
artery through the external carotid artery (Lindner et al., 1993). The compression from
the denudation procedure also causes injury to the inner layers of the media, resulting in
the switch from a contractile to a proliferative and synthetic phenotype of the smooth
muscle cells (Clowes et al., 1986). Injury to the endothelial cells or smooth muscle cells
in the arterial wall causes release of basic fibroblast growth factor (bFGF), which
stimulates the surrounding smooth muscle cells to proliferate (Lindner and Reidy, 1991).
Medial smooth muscle cell proliferation begins immediately after injury to the vessel
wall, and the smooth muscle cells begin to migrate to the intima due to platelet-derived
growth factor (PDGF) release by platelets at sites of arterial injury (Jackson et al., 1993).
Migration is facilitated by the matrix metalloproteinases (MMPs), released by smooth
11
muscle cells, which degrade the surrounding extracellular matrix components (Bendeck
et al., 1994). Examination of denuded areas of rat carotid arteries subjected to balloon
catheter injury demonstrated thickening of the intima and vessel contraction, leading to
net loss of lumen area (Clowes et al., 1983a). The intimal smooth muscle cells continue
to proliferate, with greatest activity being in the area closest to the lumen, and synthesize
new extracellular matrix consisting of 90% collagens, forming the intimal lesion (Clowes
et al., 1983b; Clowes et al., 1986; Bendeck et al., 1996b; Plenz et al., 1999a).
In Chapter 4, we studied the functions of type VIII collagen in the mouse
following arterial wire injury. Since this thesis is centered on alterations in the
extracellular matrix which occur in atherosclerosis and restenosis, the following sections
summarize how the matrix composition of the vessel wall is modulated during
atherogenesis.
1.2.3 Matrix proteolysis in atherosclerosis and restenosis
During atherogenesis, the extracellular matrix composition is affected by
degradation and turnover. Research over the last decade has identified important roles
for several families of matrix-degrading proteinases, including the MMPs, elastases,
plasmin/plasminogen activators and cathepsins. Production of proteases by smooth
muscle cells and inflammatory cells allows infiltration of these cells into the
atherosclerotic plaque and ultimately may lead to the degradation of matrix and
destabilization of the plaque structure. The expression of proteinases can be stimulated
by certain matrix molecules, thus providing the potential for feedback regulation of
matrix turnover. Because our studies concentrate on the role of type VIII collagen in
12
regulating MMPs, the roles of MMPs in atherosclerosis are discussed in the following
sections.
1.2.3.1 Matrix metalloproteinases
MMPs are a family of zinc and calcium-dependent endopeptidases responsible for
the degradation of extracellular matrix proteins. MMPs exist as secreted proteins which
can be divided into the following classes according to substrate specificity: interstitial
collagenases, which degrade fibrillar collagens I and III, collagenase/gelatinases, which
degrade basement membrane collagens and gelatins, and stromelysins, which degrade
proteoglycans, laminin, fibronectin, gelatin, and basement membrane collagens. There is
also a group of membrane-bound MMPs, called MT-MMPs, capable of activating other
MMPs. The MMPs present within the vasculature include the interstitial collagenases
MMP-1, -8, and -13, the gelatinases MMP-2 and -9, the stromelysins MMP-3, -7, and -
10, the matrix metalloelastase MMP-12, and the MT-MMPs MMP-14 and -16, which
degrade elastin, type I collagen, and activate pro-MMP-2 (Sasamura et al., 2005; Nagase
et al., 2006) (Table 1.8.2). Since, altogether, the MMPs can degrade all extracellular
matrix proteins, they are tightly regulated at three levels: transcription, proenzyme
activation, and enzymatic inhibition.
Various cytokines and growth factors can either stimulate or inhibit MMP
synthesis to aid in the regulation of MMPs at a transcriptional level. Once synthesized,
MMPs are latent proenzymes that are cleaved by various enzymes, a potent one being
plasmin, to become active enzymes. The third level of regulation is controlled via tissue
inhibitors of MMPs (TIMPs), naturally-occurring inhibitors of MMPs, which act at the
level of proenzyme activation. Interestingly, TIMP-2 will form a complex with
13
proMMP-2, which, when bound to a free MT1-MMP molecule, exposes the propeptide of
proMMP-2 to another MT1-MMP molecule, causing formation of a quaternary proMMP-
2 activation complex and auto-activation of proMMP-2 (Nagase et al., 2006).
1.2.3.2 Matrix metalloproteinases and atherogenesis
MMPs are elevated during the course of atherogenesis and particularly in the
shoulders and cores of atherosclerotic lesions, contributing to lesion growth and plaque
instability in these areas (reviewed in(Dollery and Libby, 2006). After arterial injury,
MMP-1, MMP-2, MMP-3, MMP-9, and MT1-MMP facilitate smooth muscle cell
migration by clearing a path through the extracellular matrix to aid in migration from the
media into the intima (Bendeck et al., 1994; Zempo et al., 1996; Bendeck et al., 1996a;
Lijnen et al., 1999). Increased levels of MMPs after injury are responsible for medial
smooth muscle cell migration, since using an MMP inhibitor resulted in significant
suppression of medial smooth muscle cell migration, but not proliferation (Bendeck et al.,
1996a), and overexpression of TIMP-1 (Forough et al., 1996) or gene transfer of TIMP-2
(Cheng et al., 1998) inhibited smooth muscle cell migration after rat carotid arterial
injury.
Past studies have correlated activation of MMP-2 and MMP-9 with collagen
synthesis and degradation, as use of GM6001, a general MMP inhibitor, inhibited the
increased turnover in collagen occurring after double balloon iliac artery injury in the
rabbit (Strauss et al., 1996), indicating an integral role of MMPs in collagen turnover and
lesion formation. After arterial injury in the rat, an increase in MMP-9 mRNA six hours
after injury and a decrease in MMP-2 mRNA during the first week after injury were
demonstrated (Bendeck et al., 1994) with increases in MMP-9 and MMP-2 activity
14
observed on zymograms at one and five days after injury, respectively (Bendeck et al.,
1994; Zempo et al., 1994). Paradoxically, these studies demonstrated that the decrease in
MMP-2 mRNA was not correlated with the increase in MMP-2 activity after injury,
underscoring the importance of examination of MMPs at multiple levels of regulation.
Performing rat carotid balloon catheter injury in the presence of GM6001 resulted
in a significant decrease in the number of smooth muscle cells that migrated to the intima
four days after injury while having no effect on medial smooth muscle cell replication
(Bendeck et al., 1994). In contrast, treatment of rat carotid balloon catheter injured
arteries with doxycycline, another broad-spectrum MMP inhibitor, resulted in a decrease
in intimal smooth muscle cell proliferation and elastin accumulation in addition to
inhibition of MMP-2, MMP-9, and smooth muscle cell migration (Bendeck et al., 2002).
However, doxycycline is a derivative of the tetracycline antibiotics, which are capable of
inhibiting proliferation (Guerin et al., 1992). Nonetheless, these experiments once again
suggest that increased MMP activity as a result of injury is involved in smooth muscle
cell migration. In fact, a recent study with in situ zymography examining intimal
formation in apoE-deficient mice fed a high-fat diet and subjected to carotid wire injury
demonstrated an increase in MMP-2 and MMP-9 activity from one to three weeks after
injury during the growth of the intima (Zhang et al., 2008). Using RP782, a radio-
labelled tracer that specifically targets activated MMPs, microSPECT (Single Photon
Emission Computed Tomography) to image the radiolabel, and angiography to localize
the carotid arteries, the researchers showed localized activation of MMPs in injured
carotid arteries at 2 and 3 weeks after injury. Autoradiography of excised carotid arteries
also demonstrated increased MMP activity levels at all time points tested after injury,
15
from 6 hours to 4 weeks. Examination of all data revealed a strong correlation between
increases in intimal hyperplasia and total MMP activity. While these are elegant
experiments, there are some limitations to these studies: first of all, the use of broad-
spectrum MMP inhibitors does not define which MMPs are responsible for observed
results; gelatin zymograms can only determine the presence of MMP-2 and MMP-9;
correlation of increases in specific MMPs with timecourses of specific events in the
response to injury does not necessarily indicate involvement of those MMPs in the
response; and, the use of RP782 can only indicate an increase in general MMP activity.
Much of the information on the role of specific MMPs in atherosclerosis has been
elucidated through the use of knock-out mice, especially through studying double
knockouts of apoE and various MMPs. Deletion of MMP-2 resulted in smaller, smooth
muscle cell-poor lesions in the apoE-deficient mouse demonstrating a role for this MMP
in promoting smooth muscle cell migration into the fibrous cap (Kuzuya et al., 2006).
Deletion of MMP-3 and MMP-9 resulted in larger lesions in the brachiocephalic artery
due to more matrix accumulation, but the lesions contained few smooth muscle cells and
increased macrophage accumulation, which indicates a role for these MMPs in plaque
stability (Johnson et al., 2005). However, these results may be dependent on where the
lesions are localized, as lesions within the descending and thoracic aorta contained fewer
macrophages and were less prone to rupture in the absence of MMP-3 (Silence et al.,
2001) and MMP-9 (Luttun et al., 2004). Deletion of MMP-12 resulted in more stable,
smaller plaques with increased smooth muscle cell content and decreased macrophage
accumulation (Johnson et al., 2005) while overexpression of MMP-12 in cholesterol-fed
rabbits led to advanced macrophage-rich atherosclerotic lesions (Liang et al., 2006).
16
These results demonstrate the varying effect that matrix degradation can have during
atherogenesis, based on both substrate and localization within the vasculature.
In addition to their roles in degrading existing matrix, MMPs are capable of
releasing growth factors from the matrix and activating growth factors and cytokines as
well. For example, both MMP-2 and MMP-9 are capable of directly activating latent
TGF-β sequestered within the matrix. Transgenic mice with over-expression of MMP-2
and MMP-9 gave rise to invasive breast cancer due to a large increase in activated TGF-β
(Jenkins, 2008). Likewise, MT1-MMP, MMP-1, -3, -13 (van Hinsbergh and Koolwijk,
2008) and MMP-9 (Bergers et al., 2000) were all capable of releasing active VEGF from
the matrix. This carries important implications for atherosclerosis and restenosis as
VEGF is capable of indirectly upregulating smooth muscle cell proliferation and
migration. Endothelial cells stimulated with VEGF increase their expression of bFGF
mRNA, and smooth muscle cells display increased proliferation in response to and
chemotaxis towards conditioned media from endothelial cells stimulated with VEGF
compared to control endothelial conditioned media (Li et al., 2009). Furthermore,
increased activity of MMP-2 and MMP-9 early in lesion develop may then lead to
increased levels of TGF-β in late, developed lesions, which can stimulate collagen and
proteoglycan synthesis by smooth muscle cells (Dadlani et al., 2008), resulting in lesions
with increased matrix accumulation.
1.3 Mechanisms of cell migration
Smooth muscle cell migration makes a very important contribution to lesion
growth in atherosclerosis and restenosis. Since the effect of type VIII collagen on
17
smooth muscle cell migration is studied in Chapters 2 and 3, a brief review of migration
is described below.
Migration is a multi-step process, consisting of protrusion of the cell membrane,
formation of new adhesions, myosin-based cell body contraction, and tail retraction, each
of which is influenced by the extracellular matrix components present, the cytoskeleton,
and Rho GTPases. Rho GTPases are a family of at least 20 members and play key roles
in coordinating cell migration by regulation of the cytoskeleton, with cdc42, rac1, and
RhoA regulating filopodia extension (Degani et al., 2002), lamellipodia protrusion (van
Hennik et al., 2003), and contractile actin stress fiber and stable microtubule formation
(Fukata et al., 2003), respectively.
Polymerization of the actin cytoskeleton and microtubule dynamics provide the
protrusive force at the leading edge of migrating cells to extend lamellipodia and
filopodia. Integrins in focal adhesions form attachments to the extracellular matrix
(discussed later in Section 1.4.1), which are localized in the lamellipodia of migrating
cells. Cell adhesion to the extracellular matrix activates rac and cdc42, with the
continuous formation of focal adhesions at the leading edge of cells maintaining rac
activity in a positive feedback loop (Allen et al., 1998; Bailly et al., 2000). In contrast,
strong cell adhesion correlates with high levels of Rho activity, which inhibits cell
migration, while rac can induce focal adhesion turnover by antagonizing Rho activity
(Sander et al., 1999), indicating that a proper balance of adhesive strength is required
(Cox et al., 2001).
Cell body contraction depends on the interaction of actin and myosin filaments
(Mitchison and Cramer, 1996). ATP-dependent myosin motors drive cell body
18
contraction; myosin binds to actin filaments anchored at focal adhesions and causes
contraction by pulling actin filaments past one another (Lauffenburger and Horwitz,
1996). Rho acts via Rho-kinases that phosphorylate myosin light chain and also inhibit
myosin light chain phosphatase to influence the contraction process (Amano et al., 2000).
Theoretical analysis suggests that the greatest migration speed occurs at
intermediate adhesiveness of cells (DiMilla et al., 1991), since they must both attach and
detach from substrate to migrate effectively. In fact, detachment of the cell tail is usually
the rate-limiting step of cell migration and depends upon the degradation of the focal
adhesions in the tail (Cox and Huttenlocher, 1998; Palecek et al., 1998). Microtubules
are also important for detachment and retraction of the tail. Microtubules are guided
along actin stress fibers towards focal adhesions (Krylyshkina et al., 2003) and induce
their disassembly (Ezratty et al., 2005). Removal of the adhesion to the matrix, followed
by Rho-mediated contraction of actin stress fibers previously anchored at that location,
serves to move the cell forward (Ishizaki et al., 2001). Two separate studies examining
the effects of the microtubule-stabilizing compound paclitaxel on human smooth muscle
cells in vitro demonstrated a significant decrease in both cell proliferation and migration
(Wiskirchen et al., 2004), even when co-cultured with endothelial cells and stimulated
with growth factors (Axel et al., 1997). In vivo administration of paclitaxel after injury
of rabbit arteries inhibited the formation of neointima (Axel et al., 1997). Most likely,
these preliminary results led to the current clinical use of paclitaxel on drug-eluting stents
(Vaina et al., 2005). Also, in addition to the intracellular disassembly of focal adhesions
by microtubules, MMP-1 is produced at the tail end of migrating human smooth muscle
cells, where it serves to degrade fibrillar type I collagen to release cells from the matrix
19
(Li et al., 2000). These results indicate that cell migration is a complex process
integrating multiple mechanisms for cell locomotion.
1.4 Extracellular matrix receptors involved in
atherogenesis
1.4.1 Integrins
Integrin receptors are the primary receptors for the extracellular matrix and serve
as transmembrane links between the extracellular matrix and the actin cytoskeleton of
cells. Integrins are heterodimers composed of noncovalently-associated α and β
subunits. There are 18 α and 8 β subunits that can form 24 different heterodimers which
bind to a variety of ligands (Hynes, 2002; Humphries et al., 2006). Integrins are capable
of bidirectional signaling; they can transmit information intracellularly upon ligand
binding for outside-in signaling, while they can also undergo inside-out signaling where
intracellular stimuli can cause activation of the integrins themselves. Resting and
inactive integrins have low affinity for their ligands, while activated integrins undergo
conformational changes to expose the ligand-binding site (Xiong et al., 2000). While
integrin activation increases the affinity of integrins for a matrix ligand, for cells to bind
strongly, the avidity of the interactions must be increased by clustering activated and
ligated integrins to form strong adhesions to extracellular matrix and linking through
multiprotein intracellular complexes to the actin cytoskeleton; these are termed focal
adhesions. Four main types of focal adhesion proteins are present in the integrin linkage
to the actin cytoskeleton: 1) integrin-bound proteins that directly bind actin, such as talin;
20
2) integrin-bound proteins that indirectly associate with the cytoskeleton, such as paxillin,
focal adhesion kinase (FAK), and Src; 3) non-integrin-bound actin-binding proteins, such
as vinculin; and 4) adaptor and signaling molecules regulating interactions of the focal
adhesion proteins (for a detailed review of focal adhesion proteins and integrin signaling,
see(Legate et al., 2009).
1.4.1.1 Integrins and atherogenesis
Because this thesis is focused on collagen-dependent responses in smooth muscle
cells and the factors involved in atherogenesis, in this section we will concentrate on
collagen-binding integrins and integrins implicated in atherogenesis. The predominant β
integrins in vascular smooth muscle cells in vivo and in vitro are β1 integrins (Skinner et
al., 1994). The predominant α integrins normally expressed in vivo are α1, α3, and α5
(Skinner et al., 1994; Hillis et al., 1998). Four integrin combinations function primarily
as collagen receptors: α1β1, α2β1, α10β1, and α11β1, all through the I domain in the α
subunit (Tulla et al., 2001). α1β1 and α2β1 integrins interact mainly with native fibrillar
collagens through GFOGER, GLOGER, and GASGER sequences (Siljander et al., 2004)
independent of Arg-Gly-Asp (RGD) sequences. However, once the collagen triple helix
is denatured, RGD sequences are exposed, and other integrins recognizing the RGD
sequence, in particular αvβ3, are then utilized, as was found in vitro with synthetic
smooth muscle cells exposed to heat-denatured type I collagen (Yamamoto et al., 1995).
One study showed the major collagen integrin expressed on normal human
smooth muscle cells in vivo is the α1β1 integrin with very little α2β1 and, reciprocally, in
21
vitro the α2β1 integrin predominates with very little α1β1 (Skinner et al., 1994). Other
studies also demonstrated that expression of the α1β1 integrin correlated with the
expression of the differentiated, contractile smooth muscle cell phenotype present in the
medial wall of healthy arteries in mice (Yao et al., 1997) and humans (Belkin et al.,
1990). Futhermore, primary culture smooth muscle cells (Dilley et al., 1987; Campbell
and Campbell, 1993), along with smooth muscle cells in atherosclerotic lesions (Belkin et
al., 1990), contain much smaller amounts of α1 integrins than smooth muscle cells in the
media, which indicates the α1β1 integrin is the integrin normally found in vivo while the
α2β1 integrin is present only when smooth muscle cells are in an activated state.
However, a contrasting study in rats revealed the absence of both α1β1 and α2β1 integrins
in normal vessels and the expression of α1β1 integrin in injured vessels only (Gotwals et
al., 1996). In an attempt to reconcile these findings, it should be noted that different cell
types within the vasculature can have varying expression profiles of integrins (Gotwals et
al., 1996), and these observations were in different species under different conditions, all
of which could have an effect on integrin expression profiles, necessitating further
research to elucidate a definitive integrin expression profile.
Stimulation of integrins can also lead to varying cellular responses. For example,
both α1β1 and α2β1 integrins mediate collagen gel contraction by fibroblasts and vascular
smooth muscle cells (Langholz et al., 1995; Gotwals et al., 1996). However, the ligation
of these two receptors does not always produce the same response. For example, in
fibroblasts, α2β1 integrins increase type I collagen and MMP production (Langholz et al.,
1995) through p38 kinase signaling (Ravanti et al., 1999), while α1β1 integrins can
stimulate cell proliferation through the adaptor proteins Shc, and Ras and Erk (Pozzi et
22
al., 1998). On the contrary, α1β1 integrins can also maintain the contractile and quiescent
smooth muscle cell phenotype by binding to laminin, a component of the healthy
basement membrane in vivo (Hayward et al., 1995; Walker-Caprioglio et al., 1995;
Thyberg et al., 1997). These studies demonstrate while individual integrins can bind
multiple ligands and the same ligands can bind multiple integrins, different downstream
signaling pathways are stimulated and it is the combination of the ligands and integrins
present that determines signaling.
Integrins αvβ3 and αvβ5 are upregulated early after vascular injury (Corjay et al.,
1999; Bendeck et al., 2000), in parallel with an increase in their ligands fibrinogen, fibrin,
vitronectin, and osteopontin in diseased blood vessels (Wight et al., 1985; Valenzuela et
al., 1992; O'Brien et al., 1994; Stary et al., 1995; Dufourcq et al., 1998). In fact, αvβ3
integrins have been implicated in smooth muscle cell migration both in vitro (Brown et
al., 1994; Choi et al., 1994; Yue et al., 1994; Liaw et al., 1995; Jones et al., 1996; Bilato
et al., 1997; Panda et al., 1997; Clemmons et al., 1999; Baron et al., 2000; Ikari et al.,
2000) and in vivo (Choi et al., 1994; Matsuno et al., 1994; Srivatsa et al., 1997; Slepian et
al., 1998; Bendeck and Nakada, 2001). The αvβ3 integrin ligand osteopontin promoted
adhesion and migration of smooth muscle cells in a Boyden chamber in vitro (Liaw et al.,
1994) and stimulated smooth muscle cell expression of MMP-1 in vitro and was
upregulated after rat carotid balloon catheter injury, coincident with MMP-1 upregulation
and smooth muscle cell migration (Bendeck et al., 2000). In fact, inhibition of αvβ3
integrin with the antagonistic antibody m7E3 reduced smooth muscle cell migration,
MMP activity, and intimal area after rat carotid balloon catheter injury, without affecting
cell proliferation (Bendeck and Nakada, 2001). These studies highlight the significance
23
of the extracellular matrix composition and integrin signaling in influencing cell
behavior.
In addition to their individual effects on responses, signaling pathways of
integrins can interact with growth factor receptors, leading to enhanced, synergistic
signaling (reviewed in(Eliceiri, 2001). In fact, a recent study has demonstrated that type I
collagen is able to synergistically enhance proliferation of smooth muscle cells in
response to PDGF-BB through src-dependent cross-talk with the α1β1 integrin.
Stimulation of smooth muscle cells with both type I collagen and PDGF-BB resulted in
cell proliferation at much higher levels than the additive effects of both, which was
abolished in the presence of a src inhibitor (Hollenbeck et al., 2004). Further evidence
of receptor cross-talk was demonstrated by examination of FAK activity. Cell migration
stimulated by PDGF is associated with increased PI3K activity (Kundra et al., 1994), and
PI3K in turn associates with FAK and increases its phosphorylation (Abedi and Zachary,
1995; Saito et al., 1996). This activation of FAK is required for both integrin and growth
factor-mediated cell motility (Carragher et al., 1999; Sieg et al., 1999; Sieg et al., 2000),
again demonstrating the cross-talk that occurs between the growth factor and integrin
receptors. FAK is inhibited by overexpression of FAK-related non-kinases (FRNK)
(Gilmore and Romer, 1996; Richardson and Parsons, 1996; Zheng et al., 1999) causing
decreased PDGF-BB-mediated recruitment of FAK to PDGF receptor complexes, and
decreased phosphorylation of FAK, suggesting that FAK can serve as a connection
between growth factor receptors, integrins, and downstream signaling leading to
migration (Hauck et al., 2000).
24
1.4.2 Discoidin domain receptors
Another class of collagen receptors present within the vasculature are the
discoidin domain receptor (DDR) tyrosine kinases, the first receptor tyrosine kinases
found that bind directly to the extracellular matrix (Shrivastava et al., 1997; Vogel et al.,
1997). Recent studies have demonstrated DDRs are also involved in collagen turnover
(discussed in Section 1.4.2.1). DDRs are aptly named as they contain an extracellular
domain homologous to discoidin-1, a lectin present in Dictyostelium discoideum which
meditates intercellular adhesion. There are two separate genes for DDR1 and DDR2, and
6 known splice variants of DDR1. DDR1 binds types I-V and type VIII collagens while
DDR2 binds the fibrillar type I, III, and the network-forming type X collagen
(Shrivastava et al., 1997; Vogel et al., 1997; Hou et al., 2001; Leitinger and Kwan, 2006).
Targeted deletion of the DDR1 gene in mice results in dwarfism and defects in placental
implantation and mammary gland development (Vogel et al., 2001) while deletion of the
DDR2 gene results in dwarfism, skeletal defects, and delayed wound healing (Labrador et
al., 2001).
Dimerization of the DDRs is necessary for collagen binding (Leitinger, 2003).
The minimal required sequence for DDR2 binding to fibrillar collagen was determined to
be GVMGFO (Konitsiotis et al., 2008). Upon ligand stimulation, the DDRs undergo
autophosphorylation and remain phosphorylated for up to 18 hours after stimulation.
Furthermore, neither gelatin nor collagenase-digested collagens are able to stimulate
DDR tyrosine phosphorylation, indicating a requirement for the native triple-helical
structure of collagen for receptor activation (Vogel et al., 1997).
25
Autophosphorylation of the cytoplasmic domain of DDR1 reveals various
consensus binding motifs for signaling and adaptor proteins containing SH2 and
phosphotyrosine-binding domains (Vogel, 1999). It has been shown, in human kidney
293 cells transfected with a DDR1 plasmid, that the adaptor protein Shc (Vogel et al.,
1997) and fibroblast growth factor receptor substrate-2 (Foehr et al., 2000) bind to the
juxtamembrane region of DDR1, but the ras-MAPK pathway is not activated in this
cascade. While not yet fully investigated, both DDR1 and DDR2 sequences contain
consensus sequences for the SH2 domains of Nck, GTP-activating protein, and the p85
subunit of PI3K (Vogel, 1999), which may indicate levels of possible interaction with
integrins or growth factor receptors. However, transfection of DDR1 plasmids into 293
cells demonstrated it is both active in the presence of integrin blocking antibodies and not
stimulated with growth factor administration, indicating integrin and growth factor
receptor-independent signaling (Vogel et al., 2000). One caveat of these experiments is
that this signaling was examined in cells transfected with DDR1 plasmids, so further
work is necessary to elucidate the signaling pathways present in cells which express
endogenous DDR1.
1.4.2.1 Discoidin domain receptors and atherogenesis
The functions of DDR1 in atherogenesis have received much more study than
DDR2. Using the rat carotid balloon catheter injury model, very little DDR1 expression
was present in the uninjured artery but both DDR1 mRNA and protein expression were
elevated 2 days after injury in the arterial media with continued expression in the intima
at 2 weeks (Hou et al., 2001). In vitro studies with DDR1-deficient and wild-type smooth
muscle cells demonstrated reduced adhesion to, proliferation on, and chemotaxis towards
26
type I and VIII collagens in the DDR1-deficient cells. MMP-2 and MMP-9 activity were
also dramatically downregulated in the DDR1-deficient smooth muscle cells, suggesting
an important role for DDR1 in regulating MMP activity in response to collagen
stimulation. Furthermore, intimal area after carotid wire injury in these animals showed a
70% reduction in the DDR1-deficient mice with reduced collagen accumulation (Hou et
al., 2001). Studies examining hypertension-induced renal disease (Flamant et al., 2006)
and bleomycin-induced lung fibrosis (Avivi-Green et al., 2006) in the DDR1-deficient
mice have also demonstrated decreased collagen accumulation. In contrast to DDR1 and
DDR2 deletion, overexpression of DDR1 and DDR2 in vascular smooth muscle cells led
to down-regulation of collagen production and increases in MMP expression and
activation (Ferri et al., 2004), indicating a complex role for the DDRs in regulating
collagen turnover.
Research in our laboratory with LDLR- and DDR1-doubly-deficient mice fed a
high fat diet demonstrated decreased lesion area, yet increased production of procollagen
mRNA and accumulation of collagen within the lesions (Franco et al., 2008), further
demonstrating the role of DDR1 as a collagen sensor. These doubly-deficient mice
displayed increased levels of total and fibrillar collagen and elastin at 12 weeks of fat-
feeding, but comparable levels at 24 weeks to LDLR-deficient mice. Lesions at 12
weeks in the doubly-deficient mice also contained fewer macrophages and decreased
gelatinase activity, contributing to the decreased lesion size. Although these studies seem
to contradict one another, where in one instance DDR1 seems to promote collagen
accumulation and in another decrease it, the findings actually quite similar on closer
examination. For instance, overexpression of DDRs resulted in decreased collagen
27
mRNA and increased MMP activity (Ferri et al., 2004) while the studies of LDLR- and
DDR1-doubly-deficient mice demonstrated decreased MMP activity and increased
collagen mRNA (Franco et al., 2008). Deletion of DDR1 resulted in a smaller intima in
mouse carotid wire injury models (Hou et al., 2001) and also decreased plaque burden
and lesion area in LDLR- and DDR1-doubly-deficient fat-fed mice (Franco et al., 2008).
Furthermore, the discrepancy between matrix accumulation in these two studies may be
due to the differences in atherogenesis between the two. The response in the wire injury
model is due mainly to smooth muscle cells as they were the only cell type present (Hou
et al., 2001); however, the fat-fed LDLR-deficient mouse model is characterized by an
increased accumulation of inflammatory cells (Ishibashi et al., 1994), which would serve
as a source of MMPs as well. The decreased MMP activity in the LDLR- and DDR1-
doubly-deficient mice was likely due to the decrease in macrophage accumulation,
resulting in increased matrix accumulation in the LDLR- and DDR1- doubly-deficient
mice, whereas the increase in MMP activity in the LDLR-deficient mouse resulted in
decreased matrix accumulation.
1.5 Changes in the extracellular matrix during
atherosclerosis and restenosis
The normal vascular extracellular matrix is altered in atherosclerosis and
restenosis. In particular, the dramatic increase in type VIII collagen following arterial
injury and in atherosclerotic lesions compared to healthy arteries (discussed in Section
1.6.3) provided the basis for studying the function of type VIII collagen. During
28
atherogenesis, smooth muscle cells switch from a quiescent, contractile phenotype to a
proliferative, and extracellular matrix synthetic phenotype (Campbell and Campbell,
1994b). The changes in synthesized extracellular matrix, in turn, have profound effects
on cell behavior and are discussed in the following sections.
1.5.1 Collagens and atherogenesis
Collagens make up a major portion of the extracellular matrix during
atherogenesis and can greatly influence disease progression (reviewed in(Adiguzel et al.,
2009). In fact, the extracellular matrix of fibrous atherosclerotic plaques consists of
about 60% collagens (Stary et al., 1995), with type I collagen accounting for 70% of the
collagens present (Katsuda et al., 1992). In experimental animal models, during the first
week after balloon catheter carotid artery injury, mRNAs for type I, type III (Majesky et
al., 1991), and type VIII collagen (Bendeck et al., 1996b) are significantly increased,
coincident with smooth muscle cell migration and intimal hyperplasia. Examination of
collagen turnover rates in double balloon-injured rabbit iliac arteries demonstrated a
significant increase in both collagen synthesis and collagen degradation up to four weeks
after the second injury compared to uninjured arteries, with peak rates occurring at one
week (Strauss et al., 1996). In fact, collagen synthesis rates are 50% higher in rat carotid
arteries seven days after balloon catheter injury compared to controls, while total
collagen content is the same at seven days and only increased between three weeks and
two months after injury, demonstrating the early influence of collagen turnover to inhibit
matrix accumulation (Nili et al., 2002). Continued collagen accumulation contributes to
29
lesion growth and vessel contraction, while collagen degradation can lead to plaque
instability and rupture.
In addition to their roles providing structural support in the arterial wall, in vitro
studies suggest that collagens act as signaling molecules stimulating changes in the
phenotype and behavior of smooth muscle cells, endothelial cells and macrophages. It is
not surprising that collagen production is increased after arterial injury, since studies
using collagen synthesis inhibitors revealed that de novo production of collagens was
necessary for porcine smooth muscle cell spreading and migration (Rocnik et al., 1998).
This study demonstrated that de novo collagen production affected spreading and
migration only; attachment to collagen matrices was not affected in the presence of the
collagen synthesis inhibitors. This was due to the inability of smooth muscle cells, in the
absence of collagen synthesis, to form fibrillar actin stress fibers and to cluster β1
integrins to form focal adhesions, while total numbers of actin monomers or β1 integrins
were not affected. The authors demonstrated a decrease in type I collagen production
after use of the inhibitors and that smooth muscle cell spreading was inhibited on
preformed matrices of type I, III, IV, and V collagens with collagen synthesis inhibitor
treatment, indicating the deposition of new collagen was required to modify the pre-
existing matrix. They did not identify which new collagens were needed for cell
spreading and migration to occur. This raises the possibility that type VIII collagen may
be able to modify a pre-existing matrix to allow cell spreading and migration, a question
that we examined in Chapter 2.
The same group later demonstrated that degradation of existing collagen was
necessary for smooth muscle cell migration to proceed (Li et al., 2000). Administration
30
of MMP-1 inhibitors decreased migration in response to bFGF or PDGF. Culture of
smooth muscle cells on collagenase-resistant collagen also inhibited migration, and,
furthermore, immunocytochemistry demonstrated that smooth muscle cells produce
MMP-1 beneath the tail and leading edge of the cell to facilitate migration. These studies
elegantly demonstrate that smooth muscle cells must be able to remodel their matrix for
migration.
Type IV collagen, normally found in the basement membrane surrounding smooth
muscle cells, likely serves to maintain cell quiescence. Synthetic and proliferating rabbit
smooth muscle cells have decreased expression of type IV collagen in culture and slowly
increased expression of type IV collagen as they became more contractile and quiescent
(Okada et al., 1990). In fact, PDGF, which stimulates smooth muscle cell migration after
experimental injury in vivo (Jackson et al., 1993), decreased type IV collagen synthesis in
smooth muscle cells in vitro (Okada et al., 1992), suggesting a decreased production of
basement membrane to allow for more motile smooth muscle cells.
Research also suggests that smooth muscle cells respond differently to different
states of collagen. Polymerized collagen is normally found within the vascular
extracellular matrix in vivo. A polymerized collagen matrix can be created by
neutralizing solubilized monomers of type I collagen, which then spontaneously form
fibrils over time in vitro. Plating smooth muscle cells upon 2-dimensional polymerized
type I collagen causes cells to maintain a rounded morphology and decrease focal
adhesions. Polymerized type I collagen also maintains cell quiescence by inhibiting
DNA synthesis via upregulation of p27Kip1 and p21Cip1 (Koyama et al., 1996), inhibitors
of cyclin E and cdk-2 kinase which are required for cell cycle transition. Suspending
31
smooth muscle cells within polymerized type I collagen gels retards proliferation by
upregulation of p21Cip1 (Li et al., 2003). Polymerized collagen also maintains cell
quiescence in vivo, as p27Kip1 levels decrease immediately following arterial injury
during smooth muscle cell proliferation, but are increased in correlation with collagen
deposition one week after injury (Tanner et al., 1998).
In contrast to the effect of polymerized collagen, when smooth muscle cells were
plated on type I collagen in its monomeric form (Koyama et al., 1996), cyclin E and cdk-
2 kinase activity were increased and when cells were stimulated with solubilized
monomers of type I collagen (Liu et al., 2004), there was increased phosphatidylinositol
3-kinase (PI3K) activity. Activation of these pathways results in increased proliferation,
suggesting that increased smooth muscle cell proliferation in vivo can be partially
attributed to smooth muscle cell stimulation by newly-synthesized collagens.
Examination of the different matrix constituents produced by the smooth muscle cells
when cultured on polymerized collagen compared to culture on monomeric collagen
demonstrated suppressed expression of extracellular matrix molecules, such as
fibronectin and thrombospondin-1, in smooth muscle cells on polymerized collagen.
Even stimulation of smooth muscle cells cultured on polymerized type I collagen with
PDGF-BB was unable to increase fibronectin production, demonstrating that the state of
the extracellular matrix can influence cell behavior. Furthermore, examination of
uninjured and balloon catheter injured arteries demonstrated that the same matrix
molecules that were suppressed in smooth muscle cells cultured on polymerized collagen
in vitro were suppressed in normal arteries in vivo, while matrix molecules that were
32
upregulated after injury in vivo were the same as those upregulated after smooth muscle
cell culture on monomeric collagen in vitro (Ichii et al., 2001).
Adding degraded collagen fragments, created by treating polymerized type I
collagen gels with bacterial collagenase, to smooth muscle cells plated on monomeric
collagen caused cell rounding and a decrease in focal adhesions by calpain-mediated
cleavage of focal adhesion proteins, facilitating release from the existing extracellular
matrix and cell migration. Additionally, prolonged culture of smooth muscle cells on
polymerized collagen induced the production of MMPs -1 and -2, leading to subsequent
collagen matrix degradation and cleavage of focal adhesion proteins (Carragher et al.,
1999). These findings, however, bring into question previous claims that polymerized
type I collagen maintains cell quiescence, as quiescent cells would not be thought to be
creating a positive feedforward loop whereby the presence of polymerized collagen
induces MMP production, causing matrix degradation. Possible explanations for this
paradox are that these are findings from smooth muscle cells in vitro, which may not
behave exactly as smooth muscle cells in vivo or that this is a positive feedback
regulatory mechanism to limit remodeling.
1.5.2 Glycoproteins, proteoglycans, and atherogenesis
Expression of several extracellular matrix glycoproteins is increased in
developing atherosclerotic and restenotic lesions, including osteopontin, tenascin, and
fibronectin (general review in(Raines, 2000). Fibronectin promotes the phenotypic
switch of smooth muscle cells from the contractile, quiescent phenotype to the synthetic,
proliferative phenotype (Hedin et al., 1988). It accumulates at the luminal edge after
33
arterial injury, and its fibril assembly is necessary in atherosclerotic lesions for smooth
muscle cell growth (Pickering et al., 2000). Osteopontin is absent within the normal
human vessel wall and accumulates within the neointima in clinical atherosclerosis and
restenosis due to its production by smooth muscle cells, endothelial cells, and
macrophages (O'Brien et al., 1994). Furthermore, glycoproteins such as osteopontin
(Liaw et al., 1995) and thrombospondin (Sage and Bornstein, 1991) can provide a
chemotactic stimulus and act synergistically with growth factors, respectively, to
modulate smooth muscle cell migration.
Heparin and heparan sulfate proteoglycans are believed to maintain the quiescent
and contractile phenotype of the vascular smooth muscle cells as they prevent smooth
muscle cell proliferation in vitro and neointimal formation in vivo in a rabbit model of
arterial injury when applied periadventitially in a pluronic gel at the time of surgery
(Bingley et al., 1998). Within six hours after rabbit carotid balloon catheter injury, the
pericellular arrangement of heparin is lost and does not reappear until seven days after
injury (Bingley et al., 2001). Heparan sulfate proteoglycans produced by mast cells
directly inhibit proliferation of smooth muscle cells by blocking DNA synthesis (Wang
and Kovanen, 1999), indicating that activated mast cells in the atherosclerotic intima
serve to regulate smooth muscle cell growth. Perlecan, a heparan sulfate proteoglycan,
maintains smooth muscle cell quiescence by stimulating the tumor suppressor PTEN
(phosphatase and tensin homolog), by inhibiting growth factor and integrin-stimulated
signaling, leading to cell cycle arrest, decreased cell migration, and increased apoptosis
(Garl et al., 2004). Transgenic mice with heparan sulfate-deficient perlecan demonstrate
increased neointimal formation after carotid artery injury and increased proliferation of
34
smooth muscle cells in vitro due to the reduced ability to bind and sequester bFGF (Tran
et al., 2004). This demonstrates that perlecan can suppress proliferation by affecting cell
signaling and sequestering heparin-binding growth factors. However, breeding these
mice with apoE-deficient mice resulted in increased smooth muscle accumulation, yet the
formation of smaller atherosclerotic lesions, possibly due to decreased retention of
lipoproteins within the vessel wall (Tran-Lundmark et al., 2008), indicating that further
study is needed to understand the role of perlecan during atherogenesis.
In contrast to the heparan sulfate proteoglycans, hyaluronan deposition is
increased by proliferating smooth muscle cells in balloon-injured rat carotid arteries
(Riessen et al., 1996) and is necessary for smooth muscle cell proliferation and migration
in vitro (Evanko et al., 1999). With time-lapse microscopy, it was demonstrated that
human smooth muscle cells in culture produce a matrix rich in versican and hyaluronan
immediately prior to tail retraction, membrane ruffling and cell division, and this matrix
is absent in stationary cells. Pretreatment of cells with hyaluronan oligosaccharides that
compete with the hyaluronan receptor and prevent formation of hyaluronan matrices,
inhibited both smooth muscle cell proliferation and migration, even in the presence of
PDGF (Evanko et al., 1999).
Chondroitin sulfate proteoglycans, such as versican and biglycan, stimulate
smooth muscle cell migration by increasing production of fibronectin and promoting
detachment from elastin to facilitate migration through the elastic lamellae (Hinek et al.,
1992). Chondroitin sulfate proteoglycans also affect atherogenesis through regulation of
elastogenesis (the role of elastin in atherogenesis is discussed in the following section).
Overexpression of normal biglycan in smooth muscle cells in vitro and following balloon
35
catheter carotid artery injury in vivo lead to decreased elastin synthesis and fiber
formation and increased type I collagen synthesis and deposition. Conversely,
overexpression of a mutant form of biglycan lacking chondroitin sulfate resulted in a
marked upregulation of elastin synthesis and fiber formation along with decreased
collagen synthesis in vitro and in vivo, demonstrating the role of biglycan in regulating
elastogenesis and balance in the composition of the extracellular matrix (Hwang et al.,
2008). Similarly, both knockdown of versican expression with antisense vectors (Huang
et al., 2006) and overexpression of V3, a versican variant lacking chrondroitin sulfate
(Merrilees et al., 2002) in smooth muscle cells also resulted in increased elastogenesis
and elastin fiber assembly in vitro and in vivo. The intimas in both studies were highly
structured and contained smooth muscle cells arranged in lamellar layers, similar to the
media. Furthermore, both knockdown of versican production and overexpression of V3
resulted in increased adhesion and decreased migration and proliferation rates of smooth
muscle cells in vitro (Lemire et al., 2002; Huang et al., 2006) suggesting increases in
versican production contribute to atherogenic growth.
1.5.3 Elastin and atherogenesis
An increase in elastolytic activity, measured by elastin zymography, in balloon
catheter injured rat carotid arteries is observed two weeks after injury (Zempo et al.,
1994). However, demonstrating the importance of matrix turnover in determining total
matrix content, there was a 100% increase in elastin synthesis rates one week after rat
balloon catheter carotid injury with no change in total elastin content at the same
timepoint (Nili et al., 2002). At later timepoints, elastin content was significantly
36
increased in injured arteries at three weeks and two months after injury compared to
control, indicating synthesis of elastin at these later timepoints was greater than
degradation. The increased accumulation of elastin at these later times may serve to
inhibit smooth muscle cell proliferation, since intact elastin inhibits smooth muscle cell
proliferation in vitro (Urban et al., 2002). In contrast, elastin degradation products result
in transactivation of the PDGF receptor pathway and increases in cdks and cyclins for
increased smooth muscle cell proliferation in vitro (Mochizuki et al., 2002), indicating
elastolytic activity and PDGF stimulation during atherogenesis facilitate smooth muscle
proliferation and migration through the lamellae into the intima. In fact, within six hours
after rabbit carotid balloon catheter injury, smooth muscle cells are dissociated from the
elastic lamina. Seven days after injury, smooth muscle cells are still dissociated from the
elastic fibers, have a synthetic phenotype, and appear within the internal elastic lamina to
form the early intima. By two weeks after injury, the majority of smooth muscle cells
within the media are reassociated with elastic lamina and adopt a quiescent and
contractile phenotype once more (Bingley et al., 2001).
1.6 Type VIII collagen
1.6.1 Structure and localization
Our interest in type VIII collagen stems from earlier research in our laboratory
and several others that demonstrated that this matrix protein is upregulated in
atherosclerosis and following vascular injury, and is potentially important in stimulating
smooth muscle cell migration. Type VIII collagen is a short-chain molecule composed of
37
two chains, α1(VIII) and α2(VIII), which are encoded by the Col8a1 and Col8a2 genes,
respectively. Col8a1 is located on the long arm of human chromosome 3 (Muragaki et
al., 1991b), while Col8a2 is located on the short arm of human chromosome 1 (Muragaki
et al., 1991a). Type VIII collagen is a non-fibrillar molecule with a short triple-helical
domain constituting two-thirds of the molecule and it is flanked by non-collagenous (NC)
globular domains at both the amino- and carboxy-termini, totaling 160 nm in length
(Yamaguchi et al., 1989). The molecule is composed of three chains of α1(VIII) and/or
α2(VIII), with both heterotrimers and homotrimers found in vivo (Benya and Padilla,
1986; Kapoor et al., 1986; Greenhill et al., 2000) and produced in vitro in a translation
system (Illidge et al., 1998; Illidge et al., 2001). Recent work with atomic force
microscopy and rotary shadowing electron microscopy has demonstrated homotrimers of
recombinant α2(VIII) form triple helical chains 135 nm in length and arrange in a
hexagonal lattice (Stephan et al., 2004). Very little is known about the distribution or
relative abundance of homo- and hetero-trimers in tissues in vivo.
Type VIII collagen is most structurally similar to type X collagen (Yamaguchi et
al., 1989; Yamaguchi et al., 1991), produced by hypertrophic chondrocytes in cartilage.
The α1(VIII) and α1(X) chains contain 56% homology in their triple helical domains and
61% homology in their 3’ NC1 domains of the amino acid sequence (Yamaguchi et al.,
1989). Both collagens form unusually strong trimers through their NC1 domains, which
contain three hydrophobic strips, which is thought to initiate their supramolecular
assembly into a lattice network (Kvansakul et al., 2003). Furthermore, both the α1(VIII)
and the α1(X) chain contain a similar encoding sequence (Yamaguchi et al., 1989). The
α1(VIII) gene contains 4 exons while the α1(X) gene contains three. For the α1(VIII)
38
gene, the first and second exon encode 5’ untranslated sequences and the third encodes
most of the 5’ NC2 domain. The remainder of the NC2 domain, the triple helical
domain, the NC1 domain, and the 3’ untranslated region are all encoded by the fourth
exon (Yamaguchi et al., 1991). Both the α1(VIII) and α1(X) gene contain eight similarly
located imperfections in the triple helical domain, which consist of a Gly-X-Gly sequence
instead of the characteristic Gly-X-Y triplet of the triple helix (Yamaguchi et al., 1989).
For the α2(VIII) gene as well, the entire triple helical domain and the NC1 domain are
encoded by a single exon and the α2(VIII) gene encodes triple helical and NC1 domains
of similar sizes to the α1(VIII) gene and contains eight imperfections of the triple helix
in the same location as in the α1(VIII) gene (Muragaki et al., 1991a).
1.6.2 Functions of type VIII collagen
Type VIII collagen, originally identified in culture medium of bovine aortic
endothelial cells (Sage et al., 1980), is a principle component of Descemet’s membrane in
the cornea, where it forms a 3-dimensional hexagonal latticework (Sawada et al., 1990).
Further studies showed that type VIII collagen is also present in the arterioles of the
kidney, subintima of larger arteries (Kittelberger et al., 1990), and in normal brain
parenchyma (Hirano et al., 2004), is synthesized by smooth muscle cells, endothelial
cells, and macrophages, and upregulated in areas undergoing pathologic vascular
changes, such as in atherosclerosis (Yasuda et al., 2000; Yasuda et al., 2001) and
following vascular injury (Bendeck et al., 1996b; Sibinga et al., 1997; Plenz et al.,
1999a).
39
Type VIII collagen is detected in the murine embryonic heart, brain, lung,
thymus, and placental capillaries (Sage and Iruela-Arispe, 1990), during embryonic
development of both the murine and chick heart (Iruela-Arispe and Sage, 1991), in the
fetal calf perichondrium, brain, and optic nerve sheath (Kapoor et al., 1988). Type VIII
collagen is also expressed by both neonatal and adult smooth muscle cells in culture and
within human atherosclerotic lesions (Macbeath et al., 1996). Type VIII collagen has
been detected within the vessels of brain tumors, but not in the tumor cells themselves
nor in the normal adult brain (Paulus et al., 1991).
Type VIII collagen is implicated in various other pathologies. While its
expression is very low and localized only to blood vessels in the normal kidney, in
diabetic nephropathy, type VIII collagen expression is upregulated in the glomerular and
tubular regions of the kidney, possibly due to a glucose-sensitive regulatory element
located in the promoter of the Col8a1 gene (Gerth et al., 2007). Mutations within the
Col8a2 gene were found to cause early-onset Fuch’s corneal dystrophy (Biswas et al.,
2001; Gottsch et al., 2005a), a disease characterized by a thickened cornea and massive
accumulation and abnormal assembly of type VIII collagen in Descemet’s membrane
(Gottsch et al., 2005b). Col8a2 mutations were also linked to another form of corneal
endothelial dystrophy, posterior polymorphous dystrophy (Biswas et al., 2001).
Furthermore, both cultured astrocytes and astrocytes participating in glial scar formation
in the brain in vivo were found to express high levels of type VIII collagen. Astrocytes
were also able to adhere to type VIII collagen, which enhanced the rate of cell migration
far more than collagen types I, IV, and V, and fibronectin (Hirano et al., 2004).
40
Recent work indicates that type VIII collagen is an extremely adhesive substrate
for endothelial cells (Turner et al., 2006), and may be antithrombogenic because,
compared to type I and III collagen, it only weakly supports platelet adhesion (Saelman et
al., 1994). The α2(VIII) chain was found to greatly increase adhesion of endothelial
cells when compared to fibronectin and caused increased spreading of the endothelial
cells, mediated through the α2β1 integrin (Turner et al., 2006). The increased spreading
of endothelial cells on type VIII collagen is consistent with previous findings that type
VIII collagen is synthesized by endothelial cells participating in capillary tube formation
in vitro (Sage and Iruela-Arispe, 1990). This suggests that type VIII collagen may play
important roles mediating endothelial morphogenesis. The same integrin receptor
mediates binding of smooth muscle cells (Hou et al., 2000) and platelets (Saelman et al.,
1994) to type VIII collagen. Although the α2(VIII) chain contains three RGD motifs,
binding to the α2β1 integrin of endothelial cells was through a GLOGER motif,
suggesting that these RGD motifs are masked (Turner et al., 2006).
While these previous studies have elucidated both temporal and physical
localization of type VIII collagen, they have not fully addressed whether type VIII
collagen is necessary for development or angiogenesis. Type VIII collagen-deficient
mice are viable and fertile with no observable gross anatomical abnormalities (Hopfer et
al., 2005). However, these mice have an increased distance between the corneal
endothelium and lens, a thinned corneal stroma, and a markedly thinned and altered
Descemet’s membrane, demonstrating abnormalities in anterior segment of the eyes. In
vitro studies of corneal endothelial cells isolated from these mice demonstrate decreased
proliferation and increased cellular size, perhaps explaining the thinned corneal
41
membranes, due to the presence of fewer cells. While type VIII collagen-deficient mice
displayed abnormalities in the anterior segment of the eye, these were different from the
clinical corneal dystrophies caused by mutations in the Col8a2 gene (Biswas et al., 2001;
Gottsch et al., 2005a), which result in abnormal accumulation of collagen and thickened
corneas (Gottsch et al., 2005b). Whether the corneal abnormalities in the type VIII
collagen-deficient mice affected visual ability was not examined (Hopfer et al., 2005).
Nonetheless, these studies collectively suggest that type VIII collagen is necessary for
development of the cornea.
Examination of various organs in the singly-deficient Col8a1-/- or Col8a2-/- mice
demonstrated a slight increase in Col8a2 mRNA in the heart with large decreases in
Col8a2 mRNA in all other organs in Col8a1-/- mice. In contrast, while levels of Col8a1
mRNA were largely decreased in most organs, they were almost 1.5 and 2.5 times greater
in the heart and aorta, respectively, in Col8a2-/- mice, indicating that enhanced expression
of the α1(VIII) and α2(VIII) chains may compensate for each other’s absence within the
vasculature (Hopfer et al., 2005). To date, aside from the Hopfer et al., 2005 study and
the work presented in this thesis, no other experiments have been conducted on the
Col8a1/Col8a2 mice.
1.6.3 Type VIII collagen in vascular disease
Expression of type VIII collagen was transiently but dramatically increased by
vascular smooth muscle cells following arterial injury, while the protein was virtually
undetectable in uninjured arteries (Bendeck et al., 1996b). In the rat balloon injury model
both type VIII collagen mRNA and protein were expressed by migrating and proliferating
42
smooth muscle cells throughout the intima one week after injury, but then limited to
those areas closest to the lumen two weeks later (Sibinga et al., 1997), indicating it is
predominately a product of synthetic smooth muscle cells. Its expression coincided with
dramatic increases in MMP-2 and MMP-9 expression in rat neointimal smooth muscle
cells, the critical MMPs for medial smooth muscle cell migration following arterial injury
(Bendeck et al., 1994), suggesting, but not proving, an important role for type VIII
collagen in proteinase-dependent smooth muscle cell migration. The increase in type
VIII collagen mRNA and protein expression was found to be quite robust when
compared to the modest increase in type I collagen relative to the native amount in the
vessel wall (Sibinga et al., 1997).
Subsequent to these early observations, it was demonstrated that type VIII
collagen expression was increased in several arterial injury models, in the atherosclerotic
lesions of apolipoprotein E-deficient (apoE -/-) mice, and in human atherosclerotic
lesions. Expression of type VIII collagen was elevated in cholesterol-fed rabbits (Plenz
et al., 1999b), and in the atherosclerotic lesions of cholesterol-fed rabbits subject to
balloon injury (Plenz et al., 1999a), where expression was localized to intimal smooth
muscle cells, and to macrophage-rich areas of the plaque. In the rabbit model, elevated
levels of type VIII collagen were completely reversed after the termination of cholesterol
feeding (Plenz et al., 1999b). ApoE-/- mice expressed type VIII collagen in the fibrous
cap of the atherosclerotic plaque with extremely strong expression of type VIII collagen
by isolated intimal smooth muscle cells and little expression elsewhere in the plaque
(Yasuda et al., 2000). In contrast, in the cholesterol-fed and injured rabbits, type VIII
collagen expression was distributed in intimal, medial, and adventitial layers and in high
43
abundance in the fibrous cap, plaque, and plaque shoulders (Plenz et al., 1999a), again
suggesting it is produced by synthetic smooth muscle cells. Type VIII collagen has been
similarly localized in human atherosclerotic plaques (Macbeath et al., 1996; Weitkamp et
al., 1999; Plenz et al., 1999d).
Immunostaining of human atherosclerotic arteries demonstrated the co-
distribution of type VIII collagen and granulocyte colony stimulating factor (GM-CSF),
macrophage colony stimulating factor (M-CSF) and TGF-β (Plenz et al., 1999c; Plenz et
al., 1999d). In early atherosclerotic lesions, type VIII collagen was codistributed with
GM-CSF and M-CSF, but not with TGF-β while the inverse was true for late stage
lesions. Also, stimulation of human and porcine smooth muscle cells with GM-CSF, M-
CSF, and TGF-β transiently upregulated expression of type VIII collagen mRNA (Plenz
et al., 1999c; Plenz et al., 1999d). In vivo, in rats, both bFGF and PDGF-BB were found
to stimulate expression of type VIII collagen with PDGF-BB being the most potent
inducer of expression of type VIII collagen by vascular smooth muscle cells (Bendeck et
al., 1996b; Sibinga et al., 1997). To a lesser extent than PDGF-BB, prostaglandin E1,
angiotensin II, β-estradiol, and bFGF were also found to induce type VIII collagen
expression in vitro (Sibinga et al., 1997).
Due to the influence of bFGF and PDGF-BB on smooth muscle cell proliferation
(Lindner and Reidy, 1991) and migration (Jackson et al., 1993), respectively, and the
time course and localization of type VIII collagen upregulation following vascular injury,
it was hypothesized that type VIII collagen might serve as a chemotactic substrate,
promoting smooth muscle cell migration and adhesion. Subsequently, in vitro studies
have shown that type VIII collagen was able to independently stimulate chemotaxis of
44
vascular smooth muscle cells, in a dose-dependent manner (Hou et al., 2000). Type VIII
collagen is a less adhesive substrate than type I collagen to smooth muscle cells (Sibinga
et al., 1997; Hou et al., 2000) and smooth muscle cells are able to migrate faster through
wells coated with type VIII collagen compared to type I collagen (Sibinga et al., 1997).
Furthermore, plating cells on type VIII collagen caused formation of focal adhesions by
primarily α2β1 and secondarily α1β1 integrins (Hou et al., 2000).
During the completion of this thesis, an interesting study was published by Dr.
Gary Owens’ research group showing that an atherogenic stimulus, treatment with
oxidized phospholipids, stimulates smooth muscle cell type VIII collagen expression and
cell migration (Cherepanova et al., 2009). Oxidized phospholipids, by causing synthesis
and nuclear translocation of Krüppel-like transcription factor-4 (Klf4), are able to induce
the phenotypic switching of smooth muscle cells from a differentiated, contractile, and
quiescent phenotype found in normal arteries to a dedifferentiated, synthetic, and
migratory phenotype found in atherosclerosis (Pidkovka et al., 2007). Dr. Owens’ group
demonstrated that the active components of oxidized phospholipids induced the
expression and secretion of type VIII collagen in rat smooth muscle cells in vitro, as well
as suppression of gene markers of differentiated smooth muscle cells. The increase in
type VIII collagen was specific for oxidized phospholipids as there were no effects when
smooth muscle cells were treated with vehicle or nonoxidized phospholipids.
Application of a pluronic gel containing oxidized phospholipids to adventitia of rat
carotids for 24 hours was found to induce type VIII collagen mRNA in vivo as well.
Both in vitro and in vivo, the increase in type VIII collagen expression and production
was dependent on Klf4 binding to the Col8a1 promoter. Furthermore, smooth muscle
45
cell migration stimulated by oxidized phospholipids required both Klf4 and type VIII
collagen as treatment of smooth muscle cells with either siRNA directed towards Klf4 or
type VIII collagen or utilizing Klf4-deficient smooth muscle cells abolished enhanced
migration induced by oxidized phospholipids. In a collaboration, we provided the
Owens’ research group with type VIII collagen-deficient and normal smooth muscle
cells, which allowed them to further demonstrate that type VIII collagen was required for
this response, as the type VIII collagen-deficient cells were attenuated in chemotaxis
towards oxidized phospholipids. The study also demonstrated an increase in type VIII
collagen and Klf4 mRNA and decreased expression of smooth muscle cell differentiation
genes in aortas of apoE -/- mice fed a high-fat diet (Cherepanova et al., 2009).
Unfortunately, this work concentrates predominately on the Col8a1 gene without direct
examination of effects on the Col8a2 gene, and, as previously demonstrated (Hopfer et
al., 2005), the α1(VIII) and α2(VIII) chains may compensate for one another within the
vasculature. Nonetheless, expression of Klf4 is upregulated by PDGF-BB and
responsible for downregulation of smooth muscle cell differentiation markers both in
vitro and in vivo after rat balloon carotid injury (Liu et al., 2005). This suggests that type
VIII collagen may be a marker of synthetic and dedifferentiated smooth muscle cells and
that the upregulation of type VIII collagen by atherogenic factors such as PDGF-BB
(Bendeck et al., 1996b; Sibinga et al., 1997) and oxidized phospholipids (Cherepanova et
al., 2009) is mediated by Klf4.
46
1.7 Hypothesis and objectives
The majority of information on type VIII collagen has come from studies
describing increased production during development and disease. The few studies that
have studied cellular responses to exogenous type VIII collagen suggest that type VIII
collagen plays an important role in the regulation of smooth muscle cell migration during
atherogenesis. However, the functional role of type VIII collagen endogenously
produced by smooth muscle cells in the context of the complex multi-component
extracellular matrix found in blood vessels is not currently known. We hypothesize that
during atherogenesis, smooth muscle cells increase their production of type VIII collagen
and use it to remodel the existing extracellular matrix, providing a substrate more
favorable for rapid migration and proliferation of cells, and ultimately contributing to
extensive intimal hyperplasia.
In the first set of experiments described in Chapter 2, we examined the
contribution of endogenous type VIII collagen to smooth muscle cell behavior when
cultured on a pre-existing matrix rich in type I collagen, as normally found in vivo. We
compared wild-type Col8a1+/+/Col8a2+/+ (COL8+/+) aortic smooth muscle cells to type
VIII collagen-deficient Col8a1-/-/Col8a2-/- (COL8-/-) aortic smooth muscle cells cultured
on type I collagen in order to test the following hypothesis:
Hypothesis 1: Deletion of Col8a1/Col8a2 will result in decreased migration and
proliferation rates and MMP activity in smooth muscle cells within a type I collagen-rich
environment.
47
As mentioned previously, type VIII collagen upregulates the activity of MMPs -2
and -9 in smooth muscle cells (Hou et al., 2000), and as shown in Chapter 2, type VIII
collagen-deficient (COL8-/-) smooth muscle cells contain less MMP-2 activity. Because
MMP-2 is implicated in smooth muscle cell migration from the arterial media to the
intima in response to experimental arterial injury (Bendeck et al., 1994; Strauss et al.,
1996), the experiments described in Chapter 3 were performed to determine the role of
type VIII collagen in regulating MMP-2 by comparing COL8+/+ aortic smooth muscle
cells to COL8-/- aortic smooth muscle cells. We also utilized RNA interference to knock-
down MMP-2 production and treated cells with exogenous type VIII collagen to
stimulate matrix remodeling and test the following hypothesis:
Hypothesis 2: Deletion of Col8a1/Col8a2 results in decreased migration in smooth
muscle cells due to type VIII collagen-dependent regulation of MMP-2 expression.
In vivo, type VIII collagen is upregulated following experimental arterial injury
(Bendeck et al., 1996b; Sibinga et al., 1997; Plenz et al., 1999a), yet its involvement in
the proliferative and migratory response to injury in vivo has not been examined. In the
set of experiments described in Chapter 4, we determined the contribution of endogenous
type VIII collagen in arterial wound repair in vivo by comparing COL8+/+ mice to type
VIII collagen-deficient (COL8-/-) mice in order to test the following hypothesis:
Hypothesis 3: Deletion of Col8a1/Col8a2 will result in decreased migration and
proliferation rates, MMP activity, and intimal hyperplasia following wire injury of the
carotid or femoral artery in the mouse.
48
1.8 Tables
49
Table 1.8.1 Collagens in the vessel wall (Information adapted from(Prockop and Kivirikko, 1995; Plenz et al., 2003)
Molecular Length & Structure Collagen Type Change in Plaque
Fibrillar—300nm, assemble into bundles of staggered fibrils to form fibers
I ↑
III ↑
V ↑
Fibril-associated—240nm, short triple helical regions connected by short nonhelical regions
XIV -
XVI N/A Microfibrillar—150nm, form beaded filaments VI -
Basement membrane-associated—390nm, form net-like sheets
IV ↑
XV -
XVIII -
XIX - Membrane bound—150nm hinged ectodomain XIII -
Anchoring—450nm VII -
Short chain, network-forming—135nm VIII ↑
↑, increased in atherosclerosis; -, no change in atherosclerosis; N/A, information not available
50
Table 1.8.2 MMPs in the vasculature (Information adapted from(Sasamura et al., 2005; Nagase et al., 2006)
MMP Type Alternate Name Substrate
Collagenases
MMP-1 Interstitial collagenase-1
Types I-III, VII, VIII, X collagens, gelatin, proteoglycans
MMP-8 Neutrophil collagenase Types I-III, VII, VIII, X collagens
MMP-13 Interstitial collagenase-3 Types I-III, VII, VIII, X collagens, gelatin
Gelatinases
MMP-2 Gelatinase A Types IV, V, VII, X, XI collagens, gelatin, proteoglycans, fibronectin, elastin
MMP-9 Gelatinase B Types IV, V, VII collagens, gelatin, proteoglycans, elastin
Stromelysins
MMP-3 Stromelysin-1
Types IV, VII, IX collagens, laminin, elastin, gelatin, fibronectin, proteoglycans, proMMP-1
MMP-7 Matrilysin
Type IV collagen, gelatin, laminin, elastin, fibronectin, proteoglycans, proMMP-1, -7, -8, -9
MMP-10 Stromelysin-2 Elastin, fibronectin, proteoglycans
Matrix Metalloelastase
MMP-12 Metalloelastase Types IV, V, IX, X collagens, proteoglycans, fibronectin, laminin, elastin
Membrane type-MMPs
MMP-14 MT1-MMP Types I, II, III collagens, elastin proMMP-2,-13
MMP-16 MT3-MMP Type I collagen, elastin, proMMP-2
51
Chapter 2
Contribution of type VIII collagen to smooth muscle cell migration and proliferation
Portions of this chapter have been previously published (Adiguzel et al., 2006), for which
copyright permissions were obtained
52
2.1 Introduction
Investigating the interaction of smooth muscle cells with exogenous type VIII
collagen in vitro, we and others have shown that the protein acts as an attachment and
chemotactic factor for smooth muscle cells (Sibinga et al., 1997; Hou et al., 2000).
Smooth muscle cells attach to type VIII collagen, but it is a less adhesive substrate, and
promotes greater cell migration than type I collagen. In addition, type VIII collagen
stimulates smooth muscle cell MMP synthesis, while type I collagen does not (Hou et al.,
2000). These studies were performed using exogenous type VIII collagen coated on
tissue culture plates as a substrate for the smooth muscle cells. However, in the diseased
vessel wall, type VIII collagen is expressed and deposited by smooth muscle cells in the
presence of an existing matrix rich in type I collagen. The function of endogenously
expressed type VIII collagen in this more complex matrix microenvironment has not been
studied. Before generalizations can be made about the function of type VIII collagen in
vivo, we need to fully understand what roles it plays in vitro. We will attempt to
elucidate the role of endogenous type VIII collagen in smooth muscle cells in vitro. We
have hypothesized that following arterial injury in vivo, smooth muscle cells produce
type VIII collagen and use it to overlay existing extracellular matrix, providing a
substrate more favorable for rapid migration. To be able to address this hypothesis in
vitro, we have compared aortic smooth muscle cells isolated from Col8a1+/+/Col8a2+/+
mice (COL8+/+), to smooth muscle cells isolated from type VIII collagen-deficient mice,
Col8a1-/-/Col8a2-/- (COL8-/-), to examine different components of the migratory process
when the cells are plated on either uncoated or type I collagen-coated surfaces.
53
2.2 Materials and methods
2.2.1 Chemicals and reagents
All reagents were obtained from Sigma Chemical Co. (St. Louis, MO), except where
noted otherwise.
2.2.2 Animals
Mice with targeted deletion of both the Col8a1 and Col8a2 genes (COL8-/-) were
generated in the laboratory of Dr. Bjorn Olsen (Harvard Medical School) as described
(Hopfer et al., 2005) with wild-type littermate mice (COL8+/+) used as controls.
Genotypes were verified using extracted tail DNA and polymerase chain reaction (PCR)
for both the Col8a1 and Col8a2 alleles. The primers for wild-type Col8a1 were as
follows: sense, 5'-CGG GAG TAG GAA AAC CAG GAG TGA-3'; antisense, 5'-GGC
CCA AGA ACC CCA GGA ACA-3'. Total length of product is 313 bp. The primers for
the knockout Col8a1 were as follows: sense, 5'-GTG GGG GTG GGG TGG GAT TAG
ATA-3'; antisense, 5'-CTC GGC CCA AGA ACC CCA GGA AC-3'. Total length of
product is 503 bp. The primers for wild-type Col8a2 were as follows: sense, 5'-CCG
GTA AAG TAT GTG CAG C-3'; antisense, 5'-CAA GTC CAT TGG CAG CAT C-3'.
Total length of product is 690 bp. The primers for knockout Col8a2 were as follows:
sense, 5'-CAG CGC ATC GCC TTC TAT CGC-3'; antisense, same as the wild-type
Col8a2 antisense primer. Total length of product is 1200 bp.
54
2.2.3 Cell culture
Aortic vascular smooth muscle cells were isolated from the mice by enzymatic
dispersion (Hou et al., 2001). The descending aorta was removed distal to the left
subclavian artery and proximal to the renal arteries. The advential layer was peeled off,
aortas opened longitudinally, and endothelial cells scraped off with a scalpel blade in
dissection media consisting of 1% HEPES and 1% penicillin-streptomycin in Dulbecco’s
Modified Eagle’s Medium (DMEM). Aortas from six animals were pooled for each
isolation. Medial aortic layers were then minced into smaller pieces and placed in
dispersion media consisting of 1% HEPES, 1.8 mg/mL collagenase type I (Worthington
Biochemical Co.; Freehold, NJ), 0.3 mg/mL elastase type III, 0.44 mg/mL soybean
trypsin inhibitor type I, and 2 mg/mL bovine serum albumin (BSA) in DMEM at 37ºC.
After dispersion, cells were maintained in 10% fetal bovine serum and 2% penicillin-
streptomycin supplemented DMEM, (10% FBS-DMEM) at 37ºC with 5% CO2 and used
between passages 5-10 for experiments. Unless otherwise noted, all experiments were
performed with cells in 10% FBS-DMEM. For all experiments, tissue culture
plates/flasks were either left uncoated, or coated with a solution containing 50 µg/mL of
pepsin-solubilized bovine dermal type I collagen (Collagen Biomaterials; Mahwah, NJ).
Collagen stock solution was dissolved in phosphate-buffered saline (PBS) and neutralized
with NaOH. Unless otherwise described, plates/flasks were then incubated for 1 hour at
37°C and then blocked with 10 mg/mL BSA/PBS.
55
2.2.4 Cell morphology
5,000 smooth muscle cells/well were plated in 6 well plates (uncoated) and
allowed to attach for 16 hours. Cells were then fixed with 4% paraformaldehyde and
stained with the Dif-Quik Stain Set (Dade Behring; Newark, DE). Cells were imaged
using a Nikon Eclipse TE200 inverted microscope, Hamamatsu camera (model # C4742-
95), and Simple PCI software (Compix Inc.; Mars, PA). Simple PCI software was used
to measure cell area by tracing around the outside edge of the cell, and calculating the
area within. Roundness was calculated with Simple PCI software using the formula:
Roundness = 4πArea/√(perimeter)
For detection of cytoskeletal structures, 10,000 smooth muscle cells were plated
on 1 cm round glass coverslips in a 24 well tissue culture plate. The coverslips were
either uncoated or coated with 5 ug/mL type VIII collagen. After 24 hours, cultures were
fixed with 4% paraformaldehyde and stained with TRITC-phalloidin diluted 1:400 and
either acetylated tubulin mAb (#T6793, gift from Dr. L. Langille, University of Toronto)
diluted 1:100, or Paxillin mAb (#610051, BD Biosciences; Mississauga, ON) diluted
1:100, followed by incubation with 1:200 FITC-anti-mouse Ab (#F0257), and 1:1000 To-
Pro-3 to stain the cell nuclei (Invitrogen; Burlington, ON, gift from J. Trogadis, St.
Michael’s Hospital). Projection images were obtained using a BioRad Radiance 2100
confocal microscope and LaserSharp2000 imaging software (Carl Zeiss Advanced
Imaging Microscopy; Jena, Germany). Peak excitation wavelengths were 488nm,
543nm, and 637nm for detection of emission wavelengths of 520nm for FITC, 570nm for
TRITC, and 657nm for To-Pro-3 staining, respectively.
56
Projection images were analyzed by Simple PCI software. For quantification of
focal adhesion distribution, the percentage of smooth muscle cells containing paxillin
staining on the basal surface of the cells inward from cytoplasmic extensions was
determined in each image. For quantification of stress fiber formation, the percentage of
smooth muscle cells with prominent fibrillar actin staining throughout the cell cytoplasm
was determined in each image. To analyze the extent of the stable microtubule network,
the percentage of cell area occupied by positive acetylated tubulin staining was
determined in each image. For all three, measurements were performed on multiple
images with 7-10 cells each.
2.2.5 Immunocytochemistry
12 mm round glass coverslips (Fisher Scientific; Markham, ON) were placed in
24-well plates and 50,000 cells/well were seeded and grown to confluence. A scrape-
wound was created in the monolayer by dragging a 200 µL micropipette tip across the
coverslip. The cells were washed twice with Hanks’ Balanced Salt Solution (HBSS), and
1% FBS-DMEM was added. At 0 or 24 hours after wounding, cells were rinsed twice
with PBS, then fixed with 4% paraformaldehyde. Smooth muscle cells were stained with
anti-collagen α1(VIII) (Clone 8C, Seikagaku America; East Falmouth, MA) mAb at a
dilution of 1:500 using a monoclonal antibody detection kit with AEC Chromagen (R&D
Systems; Minneapolis, MN). Smooth muscle cells were then counterstained with
hemotoxylin Quick Stain (Vector; Burlington, ON), and mounted on slides under 1:1
PBS:glycerol. Slides were imaged with a Nikon Eclipse E600 microscope, Hamamatsu
camera and Simple PCI software.
57
To localize intracellular type VIII collagen, subconfluent cells fixed as above
were double-stained with anti-58K Golgi protein (#ab5820, Abcam; Cambridge, MA)
and a Cy3-conjugated anti-rabbit secondary (#111-166-046, Jackson Immunologicals;
West Grove, PA), and anti-collagen α1(VIII) and an FITC-conjugated anti-mouse
secondary antibody (#F0257), all at a dilution of 1:250, and mounted on slides under
Prolong Antifade Gold mounting medium (Molecular Probes; Eugene, OR). Serial
images were obtained using a BioRad Radiance 2100 confocal microscope and
LaserSharp2000 imaging software. Peak excitation wavelengths were 488nm and
543nm for detection of emission wavelengths of 520nm for FITC and 570nm for Cy3,
respectively.
To localize extracellular type VIII collagen, 22 mm square glass coverslips were
placed in 6-well plates and 3,000 cells/well were seeded, and grown to confluence for 21
days. The cells were rinsed twice with PBS, and incubated with 10 mM EDTA/EGTA
until adherent cells were lifted off. Plates were then incubated with 10µg/ml pepsin in
0.1 M acetic acid for 5 minutes at 37°C. The matrix was fixed with 4%
paraformaldehyde, then stained with the type VIII collagen antibody and an FITC-
conjugated anti-mouse secondary antibody, both at a dilution of 1:100, counterstained
with 1:1000 Hoescht nuclear stain, and mounted on slides under Prolong Antifade Gold.
Slides were imaged with a Nikon Eclipse E600 microscope, Hamamatsu camera and
Simple PCI software. Excitation wavelength ranges were 340-380nm, 465-495nm, and
510-560nm to detect emission wavelengths of 460nm for Hoescht, 520nm for FITC, and
570nm for Cy3 staining, respectively.
58
2.2.6 Adhesion assays
96-well plates were either uncoated or coated with type I collagen, then incubated
at 4°C for 16 hours. 60,000 cells/well for uncoated wells and 30,000 cells/well for type I
collagen-coated wells were seeded and incubated for 1 hour at 37°C and adhesion assays
were performed as previously described (Hou et al., 2000). Briefly, cells were allowed to
adhere for 1 hour at 37°C, after which non-adherent cells were washed off with PBS.
Adherent cells were fixed and stained with 0.5% toluidine blue dissolved in 4%
paraformaldehyde, then solubilized with 1% sodium dodecyl sulphate (SDS), and
absorbance was read on a spectrophotometer (Molecular Devices; Sunnyvale, CA) at OD
595 nm.
2.2.7 Spreading and migration assays
For spreading assays, 100,000 cells/flask were seeded onto 25 cm2 tissue culture
flasks and imaged using a Nikon Eclipse TE200 inverted microscope equipped with a
heated stage. A Hamamatsu digital camera was used to capture images every 10 minutes
for 4 hours after plating. 3-6 cells were analyzed for each experiment. Migration assays
were performed similar to the spreading assays, with the following modifications:
100,000 cells were seeded onto 6-well plates, then grown until 50% confluent (0.5-2
days). Subsequently, images were captured every 10 minutes for 8 hours. 6-8 cells were
analyzed in each experiment. Spreading was expressed as the percent change in cell area
relative to start, and migration was measured as the total distance traveled over time,
using Simple PCI software.
59
2.2.8 Gelatin zymography
50,000 smooth muscle cells/well were plated onto 6 well plates, and allowed to
attach for 16 hours. Wells were then washed with HBSS, followed by incubation for 24
hours with 1 mL of serum-free DMEM containing 2% BSA. The conditioned media was
collected, and used for MMP analysis by gelatin zymography. Conditioned media from
mouse embryonic fibroblasts (MEFs) stimulated with cytochalasin D was used as a
positive control (provided by Dr. Rama Khokha, University of Toronto). 10 μL of
conditioned media was loaded into separate wells and subjected to 8% SDS-
polyacrylamide gel electrophoresis (PAGE) containing 0.1% gelatin as a substrate for
MMP activity and zymograms. After eletrophoresis, gels were washed twice in 2.5%
Triton X-100. Gels were then incubated for 16 hours overnight in buffer consisting of
0.05M Tris, 2.5mM CaCl2, and 0.02% sodium azide. After incubation, gels were fixed
with 10% TCA. Gels were then stained with full strength Coomassie Blue solution for 30
minutes, and destained with a solution consisting of 10% acetic acid and 30% methanol
in distilled water until areas of lytic activity were visualized as clear bands on a dark blue
background.
2.2.9 Type VIII collagen rescue experiments
We attempted to rescue the COL8-/- smooth muscle cells by adding exogenous
type VIII collagen (isolated from bovine Descemet’s membrane as previously described
(Kapoor et al., 1986)) to the type I collagen coated plates. Wells were first coated with a
60
solution containing 37.5 µg/mL type I collagen/PBS, and incubated at 37º C for 1 hour.
The wells were then rinsed with PBS and coated with 6.6 µg/mL exogenous type VIII
collagen/PBS. This gives a coating composed of 75% type I collagen and 25% type VIII
collagen, with the same total molar concentration as the 50 µg/mL type I collagen used in
the first experiments. Adhesion, migration, and gelatin zymography were performed on
this mixed collagen substrate as described above.
2.2.10 Proliferation assays
To measure proliferation, smooth muscle cells were plated at a density of 10,000
cells per well and allowed to attach for 48 hours. To growth arrest and synchronize the
cells, the smooth muscle cells were then incubated for 24 hours with serum-free DMEM
containing 2% BSA, which was followed by replacement of the media with 10% FBS-
DMEM supplemented with 2 µCi/mL [3H]-thymidine (Amersham Biosciences;
Piscataway, NJ). 48 hours later, wells were washed with PBS, then fixed with 10%
trichloroacetic acid (TCA). The wells were washed with 10% TCA and 95% ethanol.
Wells were then incubated with 0.3M NaOH and subsequently neutralized with the
addition of 0.3N HCl. The contents of the well were transferred to scintillation vials and
Cytoscint (ICN Biomedicals; Irvine, CA) was added. Counts per minute (CPM) were
measured using a liquid scintillation counter (Fisher Scientific).
61
2.2.11 Immunoblotting
Confluent cultures of smooth muscle cells grown in 75 cm2 flasks with 10%-FBS-
DMEM were lysed with 50 mM Tris (pH 7.6), 0.1% SDS, 0.1 mM PMSF, and 10 µg/mL
leupeptin. Total protein levels in the cell lysates were determined using a protein assay
kit (BioRad; Mississauga, ON) and 10 µg of protein was subjected to 8% SDS-PAGE and
transferred to a nitrocellulose membrane (BioRad). Membranes were blocked in 0.5%
Tween-Tris buffered saline (TBS-T) solution containing 5% non-fat milk. Membranes
were incubated overnight with an anti-collagen α1(VIII) mAb diluted 1:500 in TBS-T
containing 2.5% non-fat milk. Membranes were then incubated in horseradish
peroxidase–coupled secondary sheep anti-mouse Ig Ab (#NXA931, GE Healthcare,
Buckinghamshire, UK) diluted 1:1000 and protein expression was detected using
enhanced chemiluminescence (Perkin Elmer Life Sciences; Boston, MA). Some cultures
were grown for 21 days, and treated with a mixture of 10mM EDTA and 10mM EGTA
in PBS to lift cells off the matrix. The matrix was then scraped off the plate in lysis
buffer, and 10 µg of matrix protein was subjected to gel electrophoresis and Western
blotting for type VIII collagen as described above.
2.2.12 Cell viability assays
Cell viability assays were performed using 96-well tissue culture plates. Cells
were plated at densities of 500, 1,000, 2,500, 5,000, 7,500, or 10,000 cells per well and
incubated at 37°C for 24 hours. Cell viability assays were performed using a
Colorimetric MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrasodium bromide)
62
Assay for Cell Survival (Chemicon International Inc; Temecula, CA) with four hours of
incubation between the addition of Solutions AB and C. Plates were read on a
spectrophotometer at a test wavelength of OD 570 nm and a reference wavelength of OD
630 nm.
2.2.13 Statistics
Each experiment was repeated in duplicate or triplicate. The Dif-Quik stained
cell morphology data were analyzed by Student’s t-test. Other data were analyzed by
ANOVA, with the exception of the spreading and migration data, which were analyzed
with repeated measures ANOVA. Student-Newman-Keuls post-hoc tests were used to
determine statistically-significant differences between groups, with a significance level of
p≤0.05 (SigmaStat v3.1, Systat Software Inc.; Point Richmond, CA).
2.3 Results
2.3.1 COL8-/- smooth muscle cells are phenotypically distinct
from COL8+/+ smooth muscle cells
Western blots probed with an antibody against type VIII collagen revealed a band
of 240 kDa in the lysates from COL8+/+ smooth muscle cells, while this band was absent
in COL8-/- smooth muscle cells (Figure 2.5.1A). Smooth muscle cells obtained from
COL8+/+ and COL8-/- mice exhibited distinct morphologies in culture. When plated on
uncoated wells, COL8+/+ smooth muscle cells appeared small and elongated, usually
63
displaying only one or two long cytoplasmic protrusions (Figure 2.5.1B). By contrast,
COL8-/- smooth muscle cells were larger and rounder, and they extended multiple stellate
processes (Figure 2.5.1C). Morphometric measurements demonstrated that COL8-/-
smooth muscle cells were indeed significantly larger (2830 ± 449 μm² vs. 649 ± 61μm²),
and rounder (0.153 ± 0.021 vs. 0.112 ± 0.008, with a value of 1 corresponding to a
perfect circle) than COL8+/+ smooth muscle cells.
Plating COL8+/+ smooth muscle cells on polymerized type I collagen caused them
to round up with no protrusions visible (Figure 2.5.1D), while COL8-/- smooth muscle
cells plated on type I collagen exhibited a similar morphology to those plated on plastic
(Figure 2.5.1E). There were no significant differences in viability between COL8-/- and
COL8+/+ smooth muscle cells, whether they were plated on uncoated (COL8-/- OD570-
630=0.162 ± 0.033 vs. COL8+/+ OD570-630=0.113 ± 0.028) or on type I collagen-coated
wells (COL8-/- OD570-630=0.212 ± 0.042 vs. COL8+/+ OD570-630=0.144 ± 0.038).
Immunocytochemistry was employed to examine differences in actin stress fibers,
tubulin, and focal adhesions between subconfluent COL8-/- and COL8+/+ smooth muscle
cells plated on uncoated glass coverslips. In COL8+/+ smooth muscle cells, phalloidin
staining for actin stress fibers demonstrated few stress fibers, and any stress fibers present
were aligned along extended cytoplasmic processes (Figure 2.5.2A). By contrast, in
COL8-/- smooth muscle cells, prominent stress fibers were evident and extended
throughout the entire cell, oriented either in parallel to cytoplasmic processes, or in
multiple directions across the smooth muscle cell (Figure 2.5.2B). When COL8+/+
smooth muscle cells were plated on exogenous type VIII collagen, staining patterns for
actin stress fibers did not change significantly (Figure 2.5.2C), whereas plating COL8-/-
64
smooth muscle cells on type VIII collagen markedly decreased stress fiber formation
(Figure 2.5.2D). Quantification of the percentage of cells containing prominent stress
fibers demonstrated a significant increase in stress fiber formation in COL8-/- smooth
muscle cells compared to COL8+/+ smooth muscle cells, while addition of exogenous
type VIII collagen to COL8-/- cells significantly decreased the percentage of cells with
stress fibers (Figure 2.5.2E).
Immunostaining for α-tubulin revealed a complex microtubule network that
appeared similar between COL8-/- and COL8+/+ smooth muscle cells (not shown). In
COL8+/+ smooth muscle cells, immunostaining for acetylated tubulin, a marker of stable
microtubules, demonstrated small numbers of acetylated microtubules in the perinuclear
region, with few microtubules radiating from this central region (Figure 2.5.3A). By
contrast, in COL8-/- smooth muscle cells, much of the microtubule network was
acetylated, with acetylated microtubules extending throughout all regions of the cell
(Figure 2.5.3B). When COL8+/+ smooth muscle cells were plated on exogenous type VIII
collagen, staining patterns for acetylated tubulin did not change significantly (Figure
2.5.3C), while COL8-/- smooth muscle cells plated on exogenous type VIII collagen
displayed marked reductions in the amount of acetylated tubulin (Figure 2.5.3D).
Quantification of the percentage of area of the smooth muscle cells occupied by stable
microtubules demonstrated a significant increase in the relative amount of stable
microtubules in COL8-/- smooth muscle cells compared to COL8+/+ smooth muscle cells,
while addition of exogenous type VIII collagen to COL8-/- smooth muscle cells resulted
in a decrease in stable microtubules (Figure 2.5.3E).
65
Smooth muscle cells were stained for paxillin, to determine if there were any
differences in localization of focal adhesions between COL8-/- and COL8+/+ smooth
muscle cells. COL8+/+ smooth muscle cells displayed small regions of paxillin-positive
staining located at the most distal portions of lamellipodia (Figure 2.5.4A). By contrast,
in COL8-/- smooth muscle cells, punctate dots of paxillin staining were distributed along
the cell periphery and underneath most of the cell body, on the basal surface of the cell
(Figure 2.5.4B). When plated on exogenous type VIII collagen, paxillin staining patterns
for COL8+/+ smooth muscle cells were similar to those on glass coverslips (Figure
2.5.4C). For COL8-/- smooth muscle cells, however, paxillin staining was reduced and
found only at the periphery of cytoplasmic extensions (Figure 2.5.4D) comparable to
COL8+/+ smooth muscle cells. Quantification of the percentage of cells demonstrating
paxillin staining on the basal surface of the cell and not at the periphery demonstrated
that a significantly greater percentage of COL8-/- smooth muscle cells had basal focal
adhesions compared to COL8+/+ smooth muscle cells, whereas a similar percentage of
cells with basal focal adhesions were observed in the COL8-/- and COL8+/+ smooth
muscle cells plated on type VIII collagen (Figure 2.5.4E).
Finally, images of phalloidin stained cells showed that addition of exogenous type
VIII collagen resulted in COL8-/- smooth muscle cells becoming smaller and more
elongated in shape, comparable to COL8+/+ smooth muscle cells (Figure 2.5.5)
66
2.3.2 The production of type VIII collagen was upregulated after
injury
Confluent layers of smooth muscle cells were subject to a scrape wound, then
immunostained with an antibody against type VIII collagen. Immediately after
wounding, COL8+/+ smooth muscle cells in the uninjured monolayer and in areas
adjacent to the wound stained for type VIII collagen (Figure 2.5.6A), while COL8-/-
smooth muscle cells did not stain (Figure 2.5.6B). A substantial increase in type VIII
collagen was evident in the COL8+/+ cells twenty four hours after wounding (Figure
2.5.6C). By contrast, COL8-/- smooth muscle cells did not stain for type VIII collagen at
24 hours (Figure 2.5.6D). Type VIII collagen was localized in the cytoplasm, with a
punctate staining pattern. Double staining with an antibody raised against a marker of the
Golgi complex (58K Golgi protein marker) revealed that most of the intracellular type
VIII collagen was localized in the Golgi (Figure 2.5.6E).
After treating confluent cultures with a mixture of EDTA/EGTA to lift off cells,
we were able to detect extracellular immunostaining for type VIII collagen (Figure
2.5.7A). Furthermore, using EDTA/EGTA to lift off cells, then lysing the underlying
matrix, we were also able to detect matrix–bound type VIII collagen produced by
COL8+/+ cells but not COL8-/- cells on a Western blot (Figure 2.5.7B).
67
2.3.3 The production of type VIII collagen decreased the
attachment of smooth muscle cells to type I collagen, and
facilitated spreading and migration
Smooth muscle cells must attach to substrate to gain traction for migration;
however too strong an attachment may fix the cells in place and prevent migration. To
determine the effect of type VIII collagen production on smooth muscle cell adhesion, we
measured adhesion to uncoated wells or to wells coated with type I collagen. The
COL8-/- smooth muscle cells adhered significantly more than COL8+/+ smooth muscle
cells to uncoated wells (Figure 2.5.8A), or to wells coated with type I collagen (Figure
2.5.8B). The difference in adhesion was especially large when the cells were plated on
type I collagen. Addition of exogenous type VIII collagen to the wells to rescue the
phenotype resulted in the decreased attachment of COL8-/- cells to a level which was not
significantly different from COL8+/+ cells (Figure 2.5.8B).
Cell spreading and protrusion occur during migration and are also affected by
adhesion strength, so we compared spreading in COL8+/+ and COL8-/- smooth muscle
cells. Our preliminary experiments revealed that most spreading occurred during the first
hour after plating. When plated on uncoated flasks, COL8-/- smooth muscle cells spread
and increased cell area by only two-fold in one hour, significantly less than COL8+/+
smooth muscle cells, which increased in area by almost 4-fold (Figure 2.5.9A). COL8+/+
smooth muscle cells also spread approximately 4-fold in one hour when plated on 50
µg/mL type I collagen, significantly more than COL8-/- smooth muscle cells, which did
not spread on type I collagen (Figure 2.5.9B).
68
We used time-lapse microscopy to measure cell migration. The total distance
migrated by individual cells over an 8 hour period was calculated. Whether plated on
uncoated or type I collagen-coated wells, the distance traveled by COL8-/- smooth muscle
cells was significantly less than that for COL8+/+ smooth muscle cells (Figure 2.5.10).
When plated on uncoated wells, COL8-/- smooth muscle cells traveled a total distance of
72.7 ± 3.8 µm compared to 115 ± 9 µm for COL8+/+ smooth muscle cells (Figure
2.5.10A). When plated on 50 µg/mL type I collagen, COL8-/- smooth muscle cells
traveled a total distance of 64.2 ± 4.4 µm compared to 80.6 ± 6.2 µm for COL8+/+ smooth
muscle cells (Figure 2.5.10B). Addition of type VIII collagen to the plates rescued the
COL8-/- smooth muscle cell migration, such that the COL8-/- smooth muscle cells
traveled a distance not significantly different from the distance traveled by COL8+/+
smooth muscle cells (79.5 ± 5.8 µm) (Figure 2.5.10B).
2.3.4 Type VIII collagen production increases MMP activity
Since MMPs facilitate migration of smooth muscle cells by allowing the
clearance of matrix barriers, gelatin zymograms were employed to measure MMP-2 and -
9 activity in conditioned media from COL8+/+ and COL8-/- smooth muscle cells (Figure
2.5.11A). Conditioned media from mouse embryonic fibroblasts (MEF) was used as a
positive control, and to identify the lytic bands on zymogram gels. MEF conditioned
media contained distinct lytic bands at 95 kDa (active MMP-9), 70 kDa (latent MMP-2),
and 61 kDa (active MMP-2). Media from COL8+/+ and COL8-/- mouse smooth muscle
cells contained lytic bands of 106 kDa (latent MMP-9), 95 kDa (active MMP-9), 84 kDa
(unknown), 77 kDa (unknown), and 70 kDa (latent MMP-2). There was decreased lysis
69
in the latent MMP-2 band in the conditioned media of COL8-/- smooth muscle cells,
compared to the conditioned media of COL8+/+ smooth muscle cells. Addition of
exogenous type VIII collagen to the plate led to an increase in latent MMP-2 production
by COL8-/- cells, showing a complete rescue of the COL8-/- phenotype. By contrast, there
were no apparent differences in the activity of MMP-9 or the unidentified bands,
comparing COL8+/+ and COL8-/- smooth muscle cells. Within each cell type, there were
minimal differences in MMP activity between cells plated on plastic or on type I collagen
(Figure 2.5.11B).
2.3.5 Type VIII collagen facilitates smooth muscle cell
proliferation
To assess cell proliferation, [3H]-thymidine incorporation was measured.
Thymidine incorporation was similar in COL8+/+ and COL8-/- smooth muscle cells plated
on plastic (Figure 2.5.12). By contrast, thymidine uptake was significantly decreased in
COL8-/- smooth muscle cells plated on type I collagen, compared to COL8+/+ smooth
muscle cells.
2.4 Discussion
Knowledge about the function of type VIII collagen is scarce. However it is
expressed at high levels in injured arteries, and in the atherosclerotic lesions of man.
Previous studies using differential display PCR described the upregulation of type VIII
collagen in injured compared to uninjured rat carotid arteries (Bendeck et al., 1996b;
70
Sibinga et al., 1997). Type VIII collagen was deposited in copious amounts by smooth
muscle cells immediately subjacent to the vessel lumen, and in smooth muscle cells
forming thickened intimal lesions after injury, a pattern which correlated with smooth
muscle cell migration (Sibinga et al., 1997). Previously, we performed in vitro
experiments to study the interactions of smooth muscle cells with exogenous type VIII
collagen, and demonstrated that the protein was an adhesive and chemotactic substrate,
and also stimulated MMP synthesis by intimal smooth muscle cells (Hou et al., 2000).
Taken together, these data suggested an important role for type VIII collagen in
promoting smooth muscle cell migration.
However, the vascular extracellular matrix is a complex mixture composed of
several different types of molecules, and it is particularly rich in type I collagen. In fact,
type I collagen and type VIII collagen are both upregulated and colocalized during plaque
development (Plenz et al., 1999d). In vitro studies have shown that smooth muscle cells
can attach to type I collagen; nonetheless, a substantial body of evidence shows that
intact polymerized type I collagen inhibits cell migration and proliferation, and
downregulates the expression of many genes (Koyama et al., 1996; Carragher et al.,
1999; Ichii et al., 2001). By contrast, type VIII collagen appears to stimulate opposite
responses. Although the effects of exogenous type VIII collagen on smooth muscle cells
have been studied, the importance of endogenously produced type VIII collagen is not
known. Furthermore, the effect of type VIII collagen in the presence of a polymerized
type I collagen matrix has not been examined. In the current study, we hypothesized that
smooth muscle cells produce type VIII collagen, lay it down on top of type I collagen,
and use this modified, less-adhesive matrix to facilitate migration. To investigate this, we
71
studied aortic smooth muscle cells from COL8+/+ and COL8-/- mice. We compared the
ability of these cells to migrate on dishes coated with polymerized type I collagen, which
was used to mimic the natural environment encountered in the vascular media.
COL8-/- smooth muscle cells displayed significantly stronger attachment than
COL8+/+ smooth muscle cells, to both tissue culture plastic and to wells coated with type
I collagen substrate. This suggests that cells which are able to produce type VIII collagen
adhere less, and these results are in accordance with our previous studies where we found
that smooth muscle cell attachment to type VIII collagen was less than attachment to type
I collagen (Hou et al., 2000). In fact, when COL8-/- smooth muscle cells were plated on a
mixture of type VIII and type I collagen, their levels of attachment were reduced to a
level comparable to COL8+/+ smooth muscle cells, showing a partial rescue of the
COL8-/- phenotype, indicating the presence of type VIII collagen may confer greater
mobility to cells due to their decreased attachment to substrate.
Cell migration is dependent upon the extension of lamellipodia and formation of
stable focal adhesions in the lamellopodia which generate contractile forces through the
actin-myosin cytoskeleton, while releasing adhesions at the rear of cells allows tail
retraction and net forward migration (Webb et al., 2002). The ability to spread and
subsequently migrate depends on a critical value of adhesive strength between cell and
substrate: high or low levels of substrate attachment inhibit spreading and migration,
while maximum migration occurs at intermediate adhesion strengths (DiMilla et al.,
1991; Palecek et al., 1997), since cells must both attach and detach from substrate to
migrate effectively. We found that COL8-/- smooth muscle cells migrated a lesser
distance than COL8+/+ smooth muscle cells on both uncoated and type I collagen-coated
72
wells. This suggests that the ability to produce type VIII collagen allowed the cells to
overcome strong adhesion to type I collagen, and thus enabled migration. The COL8+/+
smooth muscle cells extruded well-defined lamellopodia, and translocated efficiently on
the substrate. By contrast, the COL8-/- cells displayed membrane ruffling with repeated
extensions and retractions of stellate processes in all directions. Importantly, we were
able to rescue COL8-/- smooth muscle cell migration by adding exogenous type VIII
collagen to the wells. Interestingly, after treating cells with collagen synthesis inhibitors,
Rocnik et al. reported that new collagen synthesis is required for smooth muscle cell
spreading and migration on preformed polymerized type I collagen matrices (Rocnik et
al., 1998). While they noted that there was a decrease in type I collagen production after
inhibitor treatment, they did not investigate the other types of collagen produced during
these processes; however, we can speculate that type VIII collagen synthesis is involved
in stimulating migration.
Another important finding in the current study was that COL8-/- cells exhibited
lower proliferation rates than COL8+/+ cells when plated on type I collagen. This
suggests that endogenously produced type VIII collagen allows cells to overcome the
inhibitory effects of type I collagen on proliferation. Likewise, type VIII collagen has
recently been implicated in stimulation of the proliferation of corneal endothelial cells
(Biswas et al., 2001; Hopfer et al., 2005).
The production of matrix-degrading enzymes such as the MMPs is required for
smooth muscle cells to detach from matrix to migrate or proliferate, and to facilitate the
clearance of matrix barriers. Gelatin zymograms revealed MMP-2 activity in the media
from COL8+/+ smooth muscle cells, while there was less MMP-2 activity in the media
73
from the COL8-/- smooth muscle cells. However, addition of exogenous type VIII
collagen to the COL8-/- smooth muscle cells increased the MMP-2 production by these
cells. These results confirm previous studies in which we found that type VIII collagen
stimulated the production of both MMP-2 and MMP-9 by rat smooth muscle cells (Hou
et al., 2000). However, we did not see a difference in MMP-9 activity in the mouse cells,
suggesting that there may be species-specific differences.
Examination of the cells in culture demonstrated that COL8-/- smooth muscle cells
are larger and rounder than COL8+/+ smooth muscle cells. We then examined the
cytoskeletal architecture of the cells as the network of actin, microtubules and focal
adhesions contribute both to cell morphology and migration. We found that COL8-/-
smooth muscle cells plated on glass coverslips contained increased stress fibers,
increased stable microtubules and prominent focal adhesions throughout the basal surface
of the cell, compared to COL8+/+ smooth muscle cells. After plating cells on exogenous
type VIII collagen, COL8-/- smooth muscle cells reverted to a phenotype similar to that of
COL8+/+ smooth muscle cells; i.e., they contained fewer actin stress fibers, fewer stable
microtubules, and fewer basal focal adhesions. The phenotype of COL8-/- smooth
muscle cells is very similar to that of human smooth muscle cells treated with
hyaluronan, which inhibits migration and proliferation (Evanko et al., 1999).
Cells usually contain two subsets of microtubules: dynamic microtubules with a
half-life of just minutes, or stable microtubules that last for hours (Bulinski and
Gundersen, 1991). Since the process of migration requires dynamic microtubules
(Ballestrem et al., 2000) and pharmacological stabilization of microtubules was
demonstrated to decrease smooth muscle cell proliferation and migration (Axel et al.,
74
1997; Wiskirchen et al., 2004), it is not surprising that the COL8-/- smooth muscle cells
displayed decreased rates of proliferation and migration compared to COL8+/+ smooth
muscle cells. Previously we have shown that smooth muscle cells are able to form focal
adhesions on type VIII collagen (Hou et al., 2000). While COL8-/- smooth muscle cells
had increased numbers of focal adhesions, what is notable is that they also rearranged the
localization; focal adhesions were present throughout the basal cell surface, but evident
only at tips of cytoplasmic extensions when plated on type VIII collagen. No appreciable
changes were seen in COL8+/+ smooth muscle cells in either stress fibers, focal
adhesions, or acetylated microtubules when plated on glass compared to plating on
exogenous type VIII collagen. COL8+/+ smooth muscle cells in culture continuously
produce type VIII collagen, which is increased when they are migrating (Figure 2.5.6A-
D). Therefore, it is likely that the 5 µg/mL coating of exogenous type VIII collagen was
not enough to stimulate a greater effect in the COL8+/+ smooth muscle cells than the
endogenously produced type VIII collagen.
Our studies have concentrated on the smooth muscle cell as the source of type
VIII collagen, and focused on smooth muscle cell interactions with this protein.
However, other cell types in the vessel wall produce type VIII collagen in the
atherosclerotic plaque, including endothelial cells (Iruela-Arispe et al., 1991), and
macrophages (Weitkamp et al., 1999). It was recently shown that endothelial cells spread
more and demonstrated increased rates of retention on type VIII collagen coated-grafts,
compared to fibronectin-coated or uncoated grafts, implanted in a goat carotid artery
(Turner et al., 2006). Aside from this, very little is known about the interactions of
endothelial cells and macrophages with the protein, but these interactions are also likely
75
to be important in mediating the injury response in vascular disease. Furthermore, how
much each cell type contributes to the production of type VIII collagen is not known.
Here, weshow that vascular smooth muscle cells derived from mice with targeted
deletion of type VIII collagen exhibit critical defects in migration and proliferation.
Furthermore, the ability of these cells to express MMP-2 is reduced compared to normal
smooth muscle cells which produce type VIII collagen. This reduction in MMP-2
activity and migration over a type I collagen matrix was reversed with addition of
exogenous type VIII collagen. These studies demonstrate that smooth muscle cells are
able to produce type VIII collagen, and use it to overlay type I collagen, providing a
provisional substrate favorable for migration. Thus, type VIII collagen may be an
important mediator of smooth muscle cell responses in vascular diseases which involve
cell migration, including atherosclerosis, restenosis, vein graft and transplant
atherosclerosis.
Acknowledgements
We would like to thank Jane English in Dr. Khokha’s laboratory for providing the
cytochalasin D-stimulated MEF conditioned media.
76
2.5 Figures
A
77
Type VIII collagen
B C
COL8+/+ COL8-/-
D E
Figure 2.5.1 COL8-/- smooth muscle cells do not produce type VIII collagen and are morphologically different from COL8+/+ smooth muscle cells
A 240 kDa band representing type VIII collagen was present within cell lysates fromA 240 kDa band representing type VIII collagen was present within cell lysates from COL8+/+ SMCs, but absent in cell lysates from COL8-/- SMCs (A). Differences in cell morphology were evident between cultures of COL8+/+ (B) and COL8-/- SMCs (C) on uncoated surfaces and between cultures of COL8+/+ (D) and COL8-/- SMCs (E) plated on type I collagen. Scalebar represents 100 µm.
78A B
C D
100
s
Presence of prominent stress fibers
*E
20
40
60
80
% S
MC
s COL8+/+
COL8-/-
Figure 2.5.2 COL8-/- smooth muscle cells have increased prominent actin stress fibers compared to COL8+/+ smooth muscle cells, which are decreased in the presence of type VIII collagen
Ph ll idi t i i f ti t fib i COL8+/+ th l ll (A) d COL8-/-
0
Col 8 - - + +
Phalloidin staining for actin stress fibers in COL8+/+ smooth muscle cells (A) and COL8-/-
smooth muscle cells (B) on uncoated surfaces and COL8+/+ smooth muscle cells (C) and COL8-/- smooth muscle cells (D) plated on type VIII collagen. *p≤0.05, A greater percentage of COL8-/- SMCs contained abundant prominent stress fibers compared to COL8+/+ SMCs (E). Upon exposure to type VIII collagen (Col 8), COL8-/- SMCs contained reduced amounts of prominent stress fibers. Scalebar represents 50 μm.
79A B
C D
area
20Presence of stable microtubules
*E
% o
f SM
C a
5
10
15 COL8+/+
COL8-/-
Figure 2.5.3 COL8-/- smooth muscle cells have a large stable microtubule network compared to COL8+/+ smooth muscle cells, which is decreased in the presence of type VIII collagen
A t l t d t b li t i i f t bl i t b l i COL8+/+ th l ll (A)
0
Col 8 - - + +
Acetylated tubulin staining for stable microtubules in COL8+/+ smooth muscle cells (A) and COL8-/- smooth muscle cells (B) on uncoated surfaces and COL8+/+ smooth muscle cells (C) and COL8-/- smooth muscle cells (D) plated on type VIII collagen. * p≤0.05, A larger percentage of cell surface area in COL8-/- SMCs contained stable microtubules compared to COL8+/+ SMCs (E). Addition of exogenous to type VIII collagen resulted in similar relative amounts of stable microtubules in both SMCs. Scalebar represents 50 μm.
80A B
C D
80
Presence of basal focal adhesions*
E
% S
MC
s
0
20
40
60 COL8+/+
COL8-/-
Figure 2.5.4 COL8-/- smooth muscle cells contain more basal focal adhesions compared to COL8+/+ smooth muscle cells, which are decreased in the presence of type VIII collagen
Paxillin staining for focal adhesions in COL8+/+ smooth muscle cells (A) and COL8-/-
0Col 8 - - + +
g ( )smooth muscle cells (B) on uncoated surfaces and COL8+/+ smooth muscle cells (C) and COL8-/- smooth muscle cells (D) plated on type VIII collagen. * p≤0.05, A larger percentage of COL8-/- SMCs contained focal adhesions present on the basal surface of the cell (arrows), as opposed to at the periphery (arrowheads). Addition of exogenous to type VIII collagen reduced basal focal adhesions in the COL8-/- SMCs to levels similar to COL8+/+ SMCs (E). Scalebar represents 50 μm.
81
A B
C
Figure 2.5.5 COL8-/- smooth muscle cells revert to a size and shape similar to COL8+/+
smooth muscle cells in the presence of type VIII collagen
Images of SMCs stained for actin stress fibers COL8+/+ SMCs (A) appeared smaller andImages of SMCs stained for actin stress fibers. COL8+/+ SMCs (A) appeared smaller and more elongated than COL8-/- SMCs (B). After plating on type VIII collagen, COL8-/-
SMCs adopted a phenotype similar to that of COL8+/+ SMCs (C). Scalebar represents 50 μm.
A B
82
C D
E
Figure 2.5.6 Type VIII collagen is upregulated in COL8+/+ smooth muscle cells after woundingg
Immunostaining revealed type VIII collagen in COL8+/+ SMCs as punctate cytoplasmic immunostaining upon injury (A), that was upregulated 24 hours after injury (C). No staining was evident in COL8-/- SMCs neither upon injury (B), or 24 hours later (D). Type VIII collagen localized in the Golgi complex of COL8+/+ SMCs (E). W = wounded area. Scalebar in A-D represents 100 µm and in E represents 20 µm.
A
83
BB
Type VIII collagen
COL8+/+ COL8-/-
matrix matrix
Figure 2.5.7 Type VIII collagen is deposited into the extracellular matrix
P i d t di t th t i i COL8+/+ SMC lt d i flPepsin was used to digest the matrix in COL8+/+ SMC cultures, and immunoflourescence staining revealed the presence of type VIII collagen in the matrix (A). The presence of type VIII collagen was confirmed on western blots of matrix lysates of COL8+/+ SMCs, but COL8-/- SMC lysates were negative (B). Scalebar represents 100 µm.
A
2
3463
84
Adhesion
OD
595
.2
.1
*
B
0 n=3 3
.5
.6
B
*
Adhesion
.2
.3
.4
OD
595
†
COL8+/+
COL8 /
0
.1
n=3 3 3
COL8-/-
COL8-/- + type VIII collagen
Figure 2.5.8 COL8-/- smooth muscle cells display increased adhesion compared to COL8+/+ smooth muscle cells
Adhesion of COL8-/- and COL8+/+ SMCs to uncoated wells (A), or to wells coated with type I collagen (B). Gray bars represent rescue experiments with COL8-/- SMCs with addition of type VIII collagen to wells. Values are mean + SEM. * p≤0.05, Adhesion of COL8-/- SMCs was significantly greater than COL8+/+ SMCs. † p≤0.05, Adhesion of COL8-
/- SMCs + type VIII collagen was significantly less than adhesion of COL8-/- SMCs.
450 **A. 0 ug/ml type I collagen
85
250
300
350
400
in c
ell a
rea
*
50
100
150
200
% c
hang
e i
n=10 9
020 30 40 50 60
Time (min)10
500 *B. 50 ug/ml type I collagen
*
250300350400450
e in
cel
l are
a
*
*
*
050
100150200250
% c
hang
e
n=8 6
010 20 30 40 50 60
Time (min)
Figure 2.5.9 COL8-/- smooth muscle cells are impaired in their ability to spread after
COL8+/+
COL8-/-
plating
Spreading of COL8-/- and COL8+/+ SMCs after plating on either uncoated wells (A), or on wells coated with type I collagen (B). Values are mean + SEM. * p≤0.05, Spreading was decreased in the COL8-/- compared to COL8+/+ SMCs.
A. 0 ug/ml type I collagen140
**
86
60
80
100
120
tanc
e (u
m)
*
*
**
*
0
20
40Dis
t
0 1 2 3 4 5 6 7 8
B. 50 ug/ml type I collagen
Time (hr)
*8090
**
††
stan
ce (u
m)
30405060
70 **
*
Dis
Time (hr)
010
20
0 1 2 3 4 5 6 7 8( )
COL8+/+
COL8-/-
COL8-/- + type VIII collagen
Figure 2.5.10 Migration of COL8-/- smooth muscle cells is impaired compared to COL8+/+ th l llCOL8+/+ smooth muscle cells
Migration of COL8-/- and COL8+/+ SMCs plated on uncoated wells (A), or on type I collagen (B). Gray circles represents rescue experiments with addition of type VIII collagen to COL8-/- SMCs. Values are mean + SEM. * p≤0.05, COL8-/- SMCs migrated less than COL8+/+ SMCs. † p≤0.05, COL8-/- SMCs + type VIII collagen migrated more than COL8-/- SMCs. N=3
A.
87
MMP-9A
MMP-2L
MMP-2A
B.
MMP-9A
MMP-2LMMP-2A
Figure 2.5.11 COL8-/- smooth muscle cells have less MMP-2 activity than COL8+/+
smooth muscle cells
(A) Gelatin zymogram containing conditioned media samples obtained from COL8-/- and(A) Gelatin zymogram containing conditioned media samples obtained from COL8 and COL8+/+ SMCs and from COL8-/- SMCs with the addition of type VIII collagen (COL8-/-
+8). (B) Gelatin zymogram containing conditioned media samples obtained from COL8-/-
and COL8+/+ plated on uncoated surfaces or on type I collagen-coated surfaces (col). MEF lane contains conditioned media from mouse embryonic fibroblasts. N=3
88
[3H]-thymidine incorporation
140
160
180
200
103
*
40
60
80
100
120
Cou
nts/
min
x 1
0
20
40
0 50Type I collagen (ug/ml)
n=4 3 4 3
COL8+/+COL8
COL8-/-
Figure 2.5.12 COL8-/- smooth muscle cells proliferate less than COL8+/+ smooth muscle cells
Proliferation of COL8-/- and COL8+/+ SMCs plated on uncoated wells or wells coated withProliferation of COL8 and COL8 SMCs plated on uncoated wells, or wells coated with type I collagen. Values are mean + SEM. Numbers at the bottom indicate n. * p≤0.05, COL8-/- SMCs incorporated significantly less [3H]-thymidine than COL8+/+ SMCs when plated on type I collagen.
89
Chapter 3
Type VIII collagen-dependent regulation of smooth muscle cell MMP-2 production and migration
90
3.1 Introduction
Type VIII collagen, which is normally present in very low amounts in the media
and adventitia (Plenz et al., 2003), is significantly increased following experimental
balloon catheter arterial injury (Bendeck et al., 1996b; Sibinga et al., 1997; Sinha et al.,
2001) and in atherosclerotic lesions (Macbeath et al., 1996; Weitkamp et al., 1999; Plenz
et al., 1999a; Plenz et al., 1999d; Yasuda et al., 2000). Expression of type VIII collagen
mRNA is regulated by platelet-derived growth factor-BB (PDGF-BB) (Bendeck et al.,
1996b; Sibinga et al., 1997), an important factor in atherogenesis. Smooth muscle cells
exhibit greater chemotaxis towards PDGF-BB when plated on type VIII collagen
compared to type I collagen (Sibinga et al., 1997), suggesting the presence of PDGF-BB
on the luminal surface following arterial injury stimulates type VIII collagen expression
which then serves to enhance smooth muscle cell migration to the intima.
Furthermore, type VIII collagen stimulates the activity of matrix
metalloproteinases (MMPs) -2 and -9 in intimal smooth muscle cells (Hou et al., 2000),
considered critical for smooth muscle cell migration from the arterial media to the intima
after rat carotid balloon injury (Bendeck et al., 1994; Zempo et al., 1994; Bendeck et al.,
1996a; Bendeck et al., 2002). In Chapter 2, we demonstrated that Col8a1-/-/Col8a2-/-
(COL8-/-) smooth muscle cells displayed decreased MMP-2 activity and migration
compared to Col8a1+/+/Col8a2+/+ (COL8+/+) smooth muscle cells, which was restored
upon addition of exogenous type VIII collagen, suggesting that the stimulation of MMP-2
activity is a mechanism by which type VIII collagen stimulates cell migration. In fact,
MMP-2-deficient smooth muscle cells display decreased migration (Kuzuya et al., 2003;
Johnson and Galis, 2004). Furthermore, studies in MMP-2-deficient mice have
91
demonstrated reduced intimal formation in the carotid artery ligation model (Kuzuya et
al., 2003; Johnson and Galis, 2004) and reduced atherosclerosis in apoE- and MMP-2-
doubly deficient mice (Kuzuya et al., 2006). We therefore hypothesized that deletion of
Col8a1/Col8a2 would result in decreased migration of smooth muscle cells secondary to
reduced production of MMP-2. To test this hypothesis, we performed migration and gel
contraction experiments comparing COL8-/- to COL8+/+ smooth muscle cells following
RNA interference to knockdown MMP-2 levels, or treatment with exogenous type VIII
collagen to rescue the COL8-/- smooth muscle cells.
3.2 Materials and methods
3.2.1 Chemicals and reagents
All reagents were obtained from Sigma Chemical Co. (St. Louis, MO), except where
noted otherwise.
3.2.2 Aortic smooth muscle cells
Mice with targeted deletion of both the Col8a1 and Col8a2 genes (COL8-/-) were
generated in the laboratory of Dr. Bjorn Olsen (Harvard Medical School) as described
(Hopfer et al., 2005) with wild-type littermate mice (COL8+/+) used as controls.
Genotypes were verified as described in section 2.2.2 with the following change in
primers for increased accuracy in genotyping: the primers for wild-type Col8a2 were as
follows: sense, 5’-CCG GTA AAG TAT GTG CAG C-3’; antisense, 5’- ATC CTG GGA
92
ACA TTG CAG G -3’. Total length of product is 480 bp. The primers for knockout
Col8a2 were as follows: sense, 5’-CAG CGC ATC GCC TTC TAT CGC-3’; antisense,
same as the wild-type Col8a2 primer. Total length of product is 900 bp.
In some experiments, mice with a targeted deletion of the MMP-2 gene
(MMP-2-/-) were utilized (originally generated in Dr. S. Itohara’s laboratory (Itoh et al.,
1997)). Genotypes were verified using extracted tail DNA and PCR for the MMP-2
allele. The primers for wild-type MMP-2 were as follows: sense, 5’-CAA CGA TGG
AGG CAC GAG TG-3’; antisense, 5’-GCC GGG GAA CTT GAT CAT GG-3’. Total
length of product is 122 bp. The primers for the knockout MMP-2 were as follows: sense,
5’-CTT GGG TGG AGA GGC TAT TC-3’; antisense, 5’-AGG TGA GAT GAC AGG
AGA TC-3’. Total length of product is 351 bp.
Aortic vascular smooth muscle cells were isolated from the mice as previously
described in section 2.2.3.
3.2.3 Transwell migration assays
65,000 smooth muscle cells in 0.25 mL of DMEM containing 200 μg/mL bovine
serum albumin (BSA-DMEM) were plated into the upper chamber of a transwell assay
apparatus fitted with a membrane containing 8 μm pores placed into a 24 well plate.
Bottom chambers of the transwell contained either 0.5mL of BSA-DMEM or 10 ng/mL
murine PDGF-BB in BSA-DMEM. Plates were incubated for 4 hours, after which the
media in the upper chamber was aspirated and cells on the upper membrane were
removed with a cotton swab. Cells that had migrated to the lower membrane were fixed
in 4% paraformaldehyde and stained with Coomassie blue. Membranes were then cut
93
and mounted onto slides. Under 100x magnification, migrated cells were counted from
four fields per membrane selected at random and counts averaged to determine the
number of migrated cells. In some experiments, 100 μg/mL doxycycline or 5 ug/mL type
VIII collagen was added to both upper and lower chambers.
3.2.4 Gel contraction assays
Thick type I collagen gels were created by mixing 1.5 mg/mL neutralized type I
collagen solution and 600,000 smooth muscle cells/mL of phenol red-free 10% FBS-
DMEM to give a final solution volume of 0.5mL/well and a concentration of 1 mg/mL
type I collagen and 100,000 smooth muscle cells/well in a 24 well plate.
Collagen/smooth muscle cell solutions were incubated at 37ºC for 2 hours to allow
complete polymerization. Gels were then released from sides and bottom of the wells by
running a spatula around the sides of the well and rapidly pipetting 1mL phenol red-free
10% FBS-DMEM to release the bottoms of the gels. Gels were imaged at 0 and 24 hours
after release with a Nikon Coolpix digital camera. Total gel area was measured using
Simple PCI software and percent contraction was calculated as 100 x [1-(area at 24
hours)/(area at 0 hours)]. Media at 24 hours was also collected for subsequent gelatin
zymography. In some experiments, 200 μg/mL doxycycline was added to the media at
the 0 hour timepoint and cells were cultured in doxycycline for the duration of the
experiment. In other experiments, smooth muscle cells were plated on 5 ug/mL type VIII
collagen for 24 hours prior to embedding in a 3-dimensional type I collagen gel.
94
3.2.5 Quantitative real-time polymerase chain reaction (qRT-
PCR)
RNA was extracted from confluent smooth muscle cell monolayers by Trizol-
chloroform phase separation. RNA was precipitated from the aqueous phase by
isopropanol and collected by centrifugation at 12,000 x g for 10 minutes. RNA was
washed with 75% nuclease-free ethanol and resuspended in nuclease-free distilled water
and quantified. DNA-free RNA was prepared by incubating 1µg RNA with RNase-free
DNase I (Fermentas; Burlington, ON) for 30 minutes at 37ºC in a buffer containing 10
mM Tris-HCl pH 7.5, 2.5 mM MgCl2, and 0.1 mM CaCl2. 1µg of EDTA was added and
the DNase inactivated by incubation at 65ºC for 10 minutes. cDNA was then reverse-
transcribed from this 1 μg of RNA using 100 ng random hexamers, 200 nM dNTP, 10
mM DTT, 40 units of RNaseOUT and 50 units of Superscript II (Invitrogen; Burlington,
ON) in a buffer containing 50 mM Tris-HCl, pH 8.3, 75 mM KCl and 3 mM MgCl2.
Quantitative PCR reactions were performed using the ABI 7900 (Applied Biosystems;
Foster City, CA). Each reaction was performed in 10 μL and contained 8ng cDNA, 450
nM each of forward and reverse primers and 1X Sybr Green Reaction Buffer (Applied
Biosystems). Murine MMP-2 (mMMP-2) was measured and murine/human acidic
ribosomal protein (mhARP) used as an endogenous internal control to account for
differences in the extraction of RNA from cells and reverse transcription of total RNA.
The conditions for the PCR reaction were as follows: 95ºC for 10 minutes followed by
40 cycles of 95ºC for 15 seconds and 60ºC for 1 minute. Following PCR, a dissociation
step was performed by heating the samples to 95ºC for 15 seconds and then cooling them
to 60ºC for 15 seconds and then heating again to 95ºC for 15 seconds. Data collected
95
during this step was used to generate the dissociation curve of the PCR product. The
cycle number at which the sample enters the exponential phase of amplification where
the cDNA doubles with every cycle (termed the critical threshold, Ct) was determined
using SDS 2.1 software (Applied Biosystems). Sequences for mMMP-2 primers (Nuttall
et al., 2004) were as follows: sense AAC TAC GAT GAT GAC CGG AAG TG;
antisense TGG CAT GGC CGA ACT CA. Sequences for mhARP primers (gift from Dr.
P. Marsden, University of Toronto) were as follows: sense CAA GCT TGC TGG TGA
AAA GGA; antisense TGA AGT ACT CAT TAT AGT CAA GGG CAT ATC. Relative
quantification of MMP-2 was determined using the comparative Ct (2-∆∆Ct) method.
Briefly, this method first normalizes the Ct for each sample to a control gene (acidic
ribosomal protein: (ARP)) in the same sample by calculating ΔCt; by subtracting the Ct
of the control gene from the Ct of the target gene (mMMP-2) in each sample. The
difference between samples (ΔΔCt) is then calculated by subtracting the ΔCt of each
sample from the ΔCt of a reference sample. Finally, the relative expression of the target
gene is expressed as an exponential fold change using the ΔΔCt of each sample.
3.2.6 SiRNA experiments
A small interfering RNA (siRNA) construct targeted against murine MMP-2 was
obtained from Ambion/Applied Biosystems (#69929) and was used to silence
endogenous MMP-2 production in COL8+/+ and COL8-/- smooth muscle cells. A non-
targeting scrambled siRNA (All-Stars Negative Control, #1027280, Qiagen; Mississauga,
ON) to control for any nonspecific effects of transfection and previously demonstrated in
our laboratory not to affect MMP-2 levels (personal communication with Dr. P. Ahmad)
96
and a non-targeting Cy3-tagged transfection control siRNA (#21459668, IDT; Coralville,
IA; gift from Dr. M. Ohh, University of Toronto) to determine transfection efficiency,
were used as internal controls. siRNA was diluted in 500 μL of Opti-MEM media with
7.5 μL lipofectamine to give a final concentration of 20 nM siRNA transfected into
200,000 smooth muscle cells in suspension in 6 well plates in 2.5 mL penicillin-
streptomycin-free 10% FBS-DMEM. 24 hours after plating, media was changed to
normal 10% FBS-DMEM. 48 hours after plating, conditioned media from wells was
collected. RNA from transfected cells was extracted by Trizol-chloroform phase
separation as described in section 3.2.5. Protein was recovered from the organic phase by
dialysis in 0.1% SDS for subsequent analysis. Alternatively, smooth muscle cells were
used 48 hrs. after plating in various experiments.
3.2.7 Time-lapse migration assays
100,000 smooth muscle cells were seeded onto 10 cm tissue culture dishes and
imaged using a Nikon Eclipse TE200 inverted microscope equipped with a heated stage.
A Hamamatsu digital camera was used to capture images of 50% confluent monolayers
every 10 minutes for 8 hours after plating. 5-8 cells were analyzed for each experiment.
Migration was measured as the total distance traveled over time, using Simple PCI
software.
97
3.2.8 Immunoblotting
Total protein levels in protein extracts from section 3.2.6 were determined using a
protein assay kit (BioRad; Mississauga, ON) and 5 µg of protein was subjected to 8%
SDS-PAGE and transferred to a nitrocellulose membrane (BioRad). To verify equal
protein loading, membranes were stained with Ponceau S after transferring. Membranes
were blocked in 0.5% TBS-T containing 5% non-fat milk and incubated overnight with
anti-MMP-2 mAb (#63179, MP Biomedicals; Solon, OH) diluted 1:500 in TBS-T
containing 2.5% non-fat milk. Membranes were then incubated with horseradish
peroxidase–coupled secondary sheep anti-mouse Ig Ab (#NXA931, GE Healthcare;
Buckinghamshire, UK) diluted 1:5000 and protein expression detected using enhanced
chemiluminescence (Perkin-Elmer; Waltham, MA). For normalization, blots were
stripped in 62.5% Tris-buffer containing 2% SDS and 0.7% β-mercaptoethanol and
reprobed with anti-GAPDH Ab (#ab8227, Abcam; Cambridge, MA) diluted 1:10,000
followed by development with horseradish-peroxidase-coupled secondary donkey anti-
rabbit Ig Ab (#NA9340V, GE Healthcare), diluted 1:5000. MMP-2 levels were analyzed
by densitometry (Image J software, National Institutes of Health, available at
http://rsb.info.nih.gov/ij/), normalized to GAPDH levels, and expressed as fold change
comparing COL8-/- to COL8+/+ smooth muscle cells or COL8+/+ smooth muscle cells
transfected with MMP-2 siRNA to COL8+/+ smooth muscle cells transfected with
scrambled siRNA.
98
3.2.9 Gelatin zymography
Media from the gel contraction and siRNA experiments was used for MMP
analysis by gelatin zymography. 5 μL of conditioned media was loaded into separate
wells on an 8% SDS-polyacrylamide gel containing 0.1% gelatin as a substrate for MMP
activity and zymograms were processed as previously described in section 2.2.8. Lytic
bands were analyzed by densitometry and expressed as a fold change compared to
COL8+/+ smooth muscle cells or COL8+/+ smooth muscle cells transfected with scrambled
siRNA used as controls.
3.2.10 Statistics
Each experiment was repeated at least three times. Data were analyzed by either
Student’s t-test or ANOVA, with the exception of the time-lapse migration data, which
was analyzed by repeated measures ANOVA. Student-Newman-Keuls post-hoc tests
were used to determine statistically-significant differences between groups, with a
significance level of p≤0.05 (SigmaStat v3.1, Systat Software Inc.; Point Richmond, CA).
3.3 Results
3.3.1 There are decreased levels of MMP-2 mRNA, protein and
activity in the COL8-/- smooth muscle cells
Previously, we have shown decreased MMP-2 activity in COL8-/- smooth muscle
cells compared to COL8+/+ smooth muscle cells (Chapter 2 &(Adiguzel et al., 2006) and
99
we wished to determine whether this was due to alterations in MMP-2 mRNA and protein
content. Using qRT-PCR, we found that COL8-/- smooth muscle cells contained
significantly less MMP-2 mRNA than COL8+/+ smooth muscle cells (65% less relative to
COL8+/+ smooth muscle cells when MMP-2 mRNA levels were normalized to an internal
control, Figure 3.5.1). Furthermore, Western blotting followed by densitometric analysis
and normalization to GAPDH levels demonstrated a 45% reduction in MMP-2 protein
levels in COL8-/- smooth muscle cells compared to COL8+/+ smooth muscle cells (Figure
3.5.2A).
3.3.2 MMP-2 mRNA, protein, and activity was decreased after
treatment with siRNA in COL8+/+ smooth muscle cells
To attenuate MMP-2 production in smooth muscle cells, we utilized siRNA
targeted against murine MMP-2. Using Cy3-tagged siRNA, we determined that
transfection efficiency was 92.5 ± 2.5% at 24 hours and 86.5 ± 0.5% at 48 hours.
Utilizing qRT-PCR, we showed effective knock-down of MMP-2 mRNA levels in MMP-
2 siRNA transfected cells compared with scrambled siRNA transfected cells (Figure
3.5.2B). Furthermore, Western blotting, densitometry, and normalization to GAPDH
levels demonstrated that treatment with MMP-2 siRNA was able to effectively knock-
down ~70% of MMP-2 protein levels in smooth muscle cells compared to smooth muscle
cells treated with the scrambled siRNA (Figure 3.5.2C), while gelatin zymograms
demonstrated a 50% reduction in MMP-2 activity in MMP-2 siRNA transfected smooth
muscle cells compared to scrambled siRNA-transfected smooth muscle cells (Figure
100
3.5.2D). Transfection with MMP-2 siRNA also caused a slight decrease in MMP-9
activity compared to transfection with scrambled siRNA.
3.3.3 COL8-/- smooth muscle cells display impaired chemotaxis
Smooth muscle cells display increased chemotaxis towards PDGF-BB when
plated on exogenous type VIII collagen compared to type I collagen (Sibinga et al.,
1997), possibly due to type VIII collagen being a less adhesive substrate for smooth
muscle cells than type I collagen (Hou et al., 2000). To examine the role of
endogenously produced type VIII collagen in stimulatingchemotaxis, we performed
transwell migration assays using PDGF-BB as a chemotactic stimulus. We demonstrated
a significant decrease in COL8-/- compared to COL8+/+ smooth muscle cell chemotaxis
towards PDGF-BB (Figure 3.5.3). Next we added exogenous type VIII collagen on both
sides of the chambers, and while chemotactic migration to PDGF-BB was unaffected in
COL8+/+ smooth muscle cells with addition of exogenous type VIII collagen, chemotactic
migration to PDGF-BB in COL8-/- smooth muscle cells was significantly increased in the
presence of type VIII collagen, and was not significantly different from COL8+/+ smooth
muscle cell migration to PDGF-BB. To examine the contribution of MMPs to smooth
muscle cell migration, doxycycline, a broad spectrum MMP inhibitor, was used.
Treatment with doxycycline significantly inhibited chemotactic migration in both COL8-/-
and COL8+/+ smooth muscle cells. In the absence of PDGF-BB stimulation, the levels of
migration were very low, and there were no significant differences in migration between
cell types, indicating the need for a chemotactic substance to stimulate transwell
migration.
101
3.3.4 MMP-2 knockdown impairs the chemotactic migration of
smooth muscle cells
Using siRNA to knockdown MMP-2 levels in COL8+/+ smooth muscle cells
resulted in a significant decrease in chemotactic migration compared to COL8+/+ smooth
muscle cells transfected with a scrambled siRNA (Figure 3.5.4). Furthermore,
significantly fewer scrambled siRNA- and MMP-2 siRNA-transfected COL8-/- smooth
muscle cells migrated towards PDGF-BB than either scrambled siRNA or MMP-2
siRNA-transfected COL8+/+ smooth muscle cells. In the absence of PDGF-BB
stimulation, the levels of migration were very low and there were no differences noted in
any of the siRNA treatment groups.
3.3.5 COL8-/- smooth muscle cell migration deficiencies are due
to decreased MMP-2
Using time-lapse image microscopy, we found COL8+/+ smooth muscle cells
transfected with scrambled siRNA migrated fastest (Figure 3.5.5). By contrast, COL8-/-
smooth muscle cells transfected with scrambled siRNA migrated significantly slower.
Knockdown of MMP-2 expression in COL8+/+ smooth muscle cells attenuated migration
to a level equivalent to that observed in COL8-/- smooth muscle cells. MMP-2
knockdown in COL8-/- smooth muscle cells did not significantly alter migration of these
cells.
102
We also performed time-lapse migration assays comparing smooth muscle cells
from MMP-2-/- and MMP-2+/+ mice (Figure 3.5.6). We found that the MMP-2-/- smooth
muscle cells displayed decreased migration compared to MMP-2+/+ smooth muscle cells.
3.3.6 COL8-/- smooth muscle cells are significantly impaired in
contracting thick collagen gels
Contraction of 3-dimensional type I collagen gels by embedded smooth muscle
cells is frequently utilized as an in vitro assay to mimic the extracellular matrix
remodeling that occurs in vivo during atherosclerosis and restenosis. Furthermore, gel
contraction and cell migration are mediated by similar mechanisms as both involve cell
contraction and matrix degradation. COL8-/- smooth muscle cells displayed significantly
decreased contraction of thick type I collagen gels compared to the COL8+/+ smooth
muscle cells (Figure 3.5.7). We pre-treated cells with exogenous type VIII collagen by
plating COL8+/+ and COL8-/- smooth muscle cells on 5 μg/mL of type VIII collagen for
24 hours before performing gel contraction assays. We found both COL8-/- and COL8+/+
smooth muscle cells were able to contract gels significantly more than COL8-/- smooth
muscle cells without preincubation, and at the same levels of contraction as COL8+/+
smooth muscle cells without preincubation. Gel contraction assays were also performed
with the addition of doxycycline to examine the effect of MMPs on gel contraction. Both
COL8-/- and COL8+/+ smooth muscle cell gel contraction was inhibited in the presence of
doxycycline with no significant differences in contraction between cell types.
Conditioned media was collected from the COL8+/+ and COL8-/- cells after
incubation in the thick collagen gels, and subject to gelatin zymography. Gelatin
103
zymography demonstrated a decrease in both MMP-9 and MMP-2 activity in COL8-/-
compared to COL8+/+ smooth muscle cells (Figure 3.5.8). Levels of both MMP-9 and
MMP-2 activity were decreased in both COL8-/- and COL8+/+ smooth muscle cells in the
presence of doxycycline. Furthermore, there were minimal differences in MMP-2 levels
between COL8-/- and COL8+/+ smooth muscle cells in the presence of doxycycline.
Using siRNA to knockdown MMP-2, we found that COL8+/+ smooth muscle cells
transfected with MMP-2 siRNA were significantly impaired in their ability to contract
gels compared to COL8+/+ cells transfected with scrambled siRNA (Figure 3.5.9).
Furthermore, MMP-2 siRNA-transfected COL8-/- smooth muscle cells and scrambled
siRNA-transfected COL8-/- smooth muscle cells were all significantly impaired in their
ability to contract gels compared to scrambled siRNA transfected COL8+/+ smooth
muscle cells. There were no differences in contraction between MMP-2 siRNA
transfected COL8+/+ smooth muscle cells and scrambled siRNA transfected COL8-/-
smooth muscle cells.
3.4 Discussion
In the absence of type VIII collagen, smooth muscle cells were impaired in their
abilities to migrate and contract type VIII collagen gels. In support of the tenet that type
VIII collagen stimulation of MMP-2 is required for these processes, the silencing of
MMP-2 in COL8+/+ smooth muscle cells by treating with MMP-2 siRNA attenuated
smooth muscle cell migration and gel contraction. Conversely, the decreases in COL8-/-
smooth muscle cell migration and gel contraction were rescued upon exposure to
exogenous type VIII collagen, which stimulates MMP-2.
104
Given that our previous work showed a decrease in MMP-2 activity in COL8-/-
compared to COL8+/+ smooth muscle cells (Chapter 2 &(Adiguzel et al., 2006), we
sought to determine whether decreased MMP-2 activity in COL8-/- smooth muscle cells
was due to decreased mRNA or protein levels of MMP-2, or simply due to a decrease in
activation of latent MMP-2. Using qRT-PCR, we demonstrated that COL8-/- smooth
muscle cells contain significantly lower amounts of MMP-2 mRNA, which could in part
explain the decreased MMP-2 protein levels observed in Western blots and decreased
MMP-2 activity in gelatin zymograms compared to COL8+/+ smooth muscle cells.
Since there was a decrease in MMP-2 mRNA, protein level, and activity in the
COL8-/- compared to the COL8+/+ smooth muscle cells, we speculated that this difference
may have been the cause of the migratory deficits seen in the COL8-/- smooth muscle
cells. To test this, we utilized a siRNA construct directed against murine MMP-2 to
decrease the levels of MMP-2 in the COL8+/+ smooth muscle cells in an attempt to
recapitulate the COL8-/- smooth muscle cell phenotype. qRT-PCR demonstrated a 97%
decrease in MMP-2 RNA after silencing with MMP-2 siRNA. Western blots of cell
lysates revealed a 70% reduction in MMP-2 levels, while gelatin zymograms
demonstrated only 50% reduction in MMP-2 activity levels between Scr and MMP-2
siRNA transfected cells; however, these data must interpreted cautiously, as the media
contained 10% FBS, which might serve as a source of MMP protein and gelatinase
activity as well. Surprisingly, transfection of cells with MMP-2 siRNA resulted in a slight
decrease in MMP-9 activity as well. We attempted to examine mRNA levels of MMP-9
with qRT-PCR; however, we were not able to detect transcript despite using two distinct
sets of primers. Future work with validated mMMP-9 primers and Western blotting may
105
be required to determine if MMP-2 siRNA transfection has any effect on MMP-9 mRNA
and protein levels, respectively.
In the transwell migration experiments, PDGF-BB was used as it is a potent
chemotactic factor and has been shown to upregulate type VIII collagen expression in
smooth muscle cells (Bendeck et al., 1996b; Sibinga et al., 1997). Furthermore, previous
work has shown that PDGF-BB is able to stimulate MMP-2 expression in rat (Uzui et al.,
2000; Risinger, Jr. et al., 2006) and human smooth muscle cells (Borrelli et al., 2006),
suggesting increased migration, due to increased MMP-2 expression, will occur in the
presence of PDGF-BB. Indeed, we observed a significant attenuation of chemotaxis of
COL8-/- smooth muscle cells compared to COL8+/+ smooth muscle cells towards PDGF-
BB, which was reversed upon addition of exogenous type VIII collagen. Using
doxycycline to inhibit MMPs, we found that migration was inhibited in both COL8-/- and
COL8+/+ smooth muscle cells, suggesting that the migration was MMP-dependent.
However, doxycycline is a broad spectrum MMP inhibitor and can also have off-target
effects such as increased smooth muscle cell adhesion (Franco et al., 2006). Therefore,
we also silenced MMP-2 to determine the contribution of MMP-2 to the observed
differences in migration. We found that MMP-2 knockdown attenuated the chemotaxis
of COL8+/+ smooth muscle cells. In addition, both MMP-2 and scrambled siRNA-
transfected COL8-/- smooth muscle cells were inhibited in their chemotaxis compared to
COL8+/+ smooth muscle cells. Taken together, this data suggests that the deficit in
COL8-/- smooth muscle cell chemotaxis was due to the decreased amount of MMP-2
produced by these cells.
106
We also measured random migration of smooth muscle cells using time-lapse
image capture. We found that MMP-2 siRNA-transfected COL8+/+ smooth muscle cells
displayed significantly decreased migration rates compared to scrambled siRNA-
transfected COL8+/+ smooth muscle cells, suggesting that MMP-2 was required for cell
migration in this assay. Similar results were obtained in experiments using MMP-2-
deficient smooth muscle cells. These results are congruent with previous data
demonstrating impaired smooth muscle cell migration in the absence of MMP-2 (Kuzuya
et al., 2003; Johnson and Galis, 2004). Furthermore, knocking down MMP-2 in the
COL8+/+ smooth muscle cells reduced migration to levels equivalent to those of COL8-/-
smooth muscle cells and COL8-/- smooth muscle cells with MMP-2 knockdown. Taken
together, this data indicates that the deficits in migration of the COL8-/- smooth muscle
cells are most likely due to the decrease in MMP-2.
Since cell contraction is involved in migration and cell migration and extracellular
matrix degradation can also lead to constrictive arterial remodeling (Newby, 2005), we
assayed the ability of COL8-/- and COL8+/+ smooth muscle cells to contract a thick type I
collagen gel as an index of constrictive remodeling. COL8-/- smooth muscle cells were
significantly impaired in their ability to contract type I collagen gels compared to the
COL8+/+ smooth muscle cells. Consistent with previous experiments in our laboratory,
we showed that gel contraction was inhibited by doxycycline (Franco et al., 2006). It was
previously demonstrated that MMP-2 was required for endothelial cell migration within
3-dimensional collagen matrices and that both total levels and activation of MMP-2 were
increased in 3-dimensional culture (Haas et al., 1998). Furthermore, nonspecific
inhibition of MMPs resulted in a decrease in MMP-2 activity as well as disruption of
107
endothelial cell organization, suggesting a role for MMP-2 in endothelial migration
through 3-dimensional collagen. To eliminate the possibility of non-specific effects of
broad-spectrum MMP inhibitors and to examine the contribution of only MMP-2, RNA
interference was utilized to perform MMP-2 knockdown. MMP-2 siRNA-transfected
COL8+/+ smooth muscle cells, MMP-2 siRNA-transfected COL8-/- smooth muscle cells,
and scrambled siRNA-transfected COL8-/- smooth muscle cells were all significantly
impaired in their ability to contract gels and there were no differences in contraction
between these cell types, indicating that attenuating MMP-2 expression and activity in
COL8+/+ smooth muscle cells caused them to behave similarly to COL8-/- smooth muscle
cells. These results are in contrast to previous studies demonstrating that MMP-9, and
not MMP-2, is responsible for gel contraction by SMCs (Johnson and Galis, 2004).
However, it should be noted that this work was performed using MMP-2-/- SMCs, which
may have had compensatory increases in levels of other proteinases. While the authors
indicated no compensatory increase in MMP-9 levels in the MMP-2-/- SMCs in the
carotid artery ligation model (Johnson and Galis, 2004), the levels of the two gelatinases
were only examined in vivo and no other proteinase levels were assayed. In contrast, the
group that made the MMP-2-/- mice demonstrated increased levels of MMP-9 activity in
the MMP-2-/- mice compared to MMP-2+/+ mice at 3, 14, and 28 days after flow cessation
in the same carotid artery ligation model (Kuzuya et al., 2003), demonstrating a
discrepancy exists in the literature. We are confident, however, that MMP-2 does
contribute to our gel contraction model since gel contraction was inhibited in the
presence of MMP-2 siRNA. Furthermore, it was previously demonstrated that arterial
contraction in hypoxic conditions was impaired in both MMP-2-deficient aortic rings and
108
rat arterial segments exposed to a cyclic MMP-2, indicating a role for MMP-2 in vessel
contraction (He et al., 2007). There also appeared to be an effect of siRNA transfection
in that both COL8+/+ and COL8-/- smooth muscle cells transfected with scrambled siRNA
contracted about 20% more than COL8+/+ and COL8-/- smooth muscle cells that were not
transfected. However, any effect of transfection would be consistent in all groups,
therefore any differences observed between mMMP-2 siRNA and scrambled siRNA
transfection are attributable to silencing of MMP-2 and not due to a side effect of
transfection.
Exposing COL8-/- smooth muscle cells to type VIII collagen for 24 hours prior to
performing the gel contraction assays was sufficient to rescue the attenuation of gel
contraction. We have previously shown that addition of exogenous type VIII collagen for
24 hours increases MMP-2 activity in the COL8-/-cells to levels comparable to COL8+/+
cells (Chapter 2 &(Adiguzel et al., 2006), suggesting that differences between COL8-/-
and COL8+/+ smooth muscle cells in contracting the gels may have been due to
differences in MMP-2 levels. The fact that knock down of MMP-2 in the COL8+/+
smooth muscle cells increased their similarities to COL8-/- smooth muscle cells and
exposure to type VIII collagen increased COL8-/- smooth muscle cell similarity to
COL8+/+ smooth muscle cells lends further strength to our hypothesis that type VIII
collagen regulates gel contraction through regulation of MMP-2.
In conclusion, we have shown that COL8-/- smooth muscle cells are impaired in
their abilty to produce MMP-2 as well as their ability to migrate and contract type I
collagen gels. Addition of exogenous type VIII collagen to the COL8-/- smooth muscle
cells rescues this phenotype, while knockdown of MMP-2 in COL8+/+ smooth muscle
109
cells mimics the COL8-/- phenotype. Based on this data, we believe type VIII collagen
functions to regulate and enhance migration through its upregulation of MMP-2.
Acknowledgements
We would like to thank Brent Steer for technical assistance in the analysis of qRT-PCR
assays and Judy Trogadis for technical assistance with the confocal microscope. We
would also like to thank Dr. Michael Ward for supplying the MMP-2-deficient mice.
110
3.5 Figures
111
MMP 2 mRNA
1
1.2*
MMP-2 mRNA
0.6
0.8
1
Fold
cha
nge
0
0.2
0.4
n=3 3
COL8+/+
COL8-/-
Figure 3.5.1 COL8-/- smooth muscle cells contain less MMP-2 mRNA than COL8+/+gsmooth muscle cells
Quantitative real-time RCR was used to measure mRNA levels for MMP-2 in COL8-/-
and COL8+/+ SMCs. Values are mean ± SEM. * p≤0.05, MMP-2 mRNA is significantly reduced in COL8-/- SMCs compared to COL8+/+ SMCs.
A B
112
Scr siRNA
MMP-2 siRNA MMP-2 mRNA
0 4
0.6
0.8
1.0
1.2
old
Cha
nge
……*
MMP-2
1.00 0.55
0
0.2
0.4FoGAPDH
C D
GAPDH
MMP-2MMP-2
MMP-9
Densitometry 1.00 0.31
GAPDHMMP-9 1.00 0.71
MMP-2 1.00 0.50Figure 3.5.2 Levels of MMP-2 are effectively knocked-down after administration of MMP-2 siRNA
Representative Western blot demonstrating a decrease in MMP-2 protein levels in COL8-/-
SMCs compared to COL8+/+ SMCs (A). Numbers indicate densitometry normalized to GAPDH and expressed relative to COL8+/+ SMCs. Quantitative real-time RCR demonstrated a significant (* p≤0.05) reduction in MMP-2 mRNA after transfection of COL8+/+ SMCs with MMP-2 siRNA relative to scrambled siRNA (Scr siRNA) (B). Representative Western blot demonstrating a reduction in MMP-2 protein levels after transfection of COL8+/+ SMCs with MMP-2 siRNA (C). Numbers indicate densitometry
li d GA d d l i S i A inormalized to GAPDH and expressed relative to Scr siRNA treatment. Representative gelatin zymogram demonstrating reduced MMP-2 activity in conditioned media of COL8+/+ cells after transfection with MMP-2 siRNA (D). The same relative decreases for MMP-2 expression and activity were observed in COL8-/- SMCs transfected with MMP-2 siRNA compared to Scr siRNA. Numbers indicate densitometry normalized to Scr siRNA activity levels. N=3 for all experiments
113
Ch t i
100
120
100
120
Chemotaxis
†
60
80
umbe
r of
cel
ls
60
80
*
20
40
Nu
20
40
‡ ‡
PDGF-BB - - - - + + + + + +
DOX - - + + - - - - + +
COL 8 + +
0033 3 4 3 444 8 8n=
COL 8 - - - - - - + + - -
COL8+/+
COL8-/-
Figure 3.5.3 COL8-/- smooth muscle cells display less chemotaxis towards PDGF-BB than COL8+/+ smooth muscle cells
Number of SMCs migrating in response to PDGF-BB, in the presence or absence of doxycycline (DOX) or exogenous type VIII collagen (COL 8). * p≤0.05, COL8-/- SMCs were significantly impaired in their ability to migrate towards PDGF-BB compared COL8+/+ SMCs. † p≤0.05, Chemotaxis of COL8-/- SMCs to PDGF-BB was significantly † p , g yincreased upon exposure to exogenous type VIII collagen. ‡p ≤0.05, Doxycycline inhibited migration in both COL8-/- and COL8+/+ SMCs. Values are mean ± SEM.
114
Ch t i d RNA i t f
COL8+/+ -Scr siRNA
COL8+/+ -MMP-2 siRNA
120
*
Chemotaxis and RNA interference
COL8-/- -Scr siRNA
COL8-/- -MMP-2 siRNA
60
80
100
er o
f cel
ls
20
40Num
be † †
0
PDGF-BB - - - - + + + +
3 3 3 3444 4n=
Figure 3.5.4 MMP-2 siRNA inhibits chemotaxis in COL8+/+ smooth muscle cells
Number of SMCs migrating in response to PDGF-BB after transfection of scrambled, non-targeting siRNA (Scr siRNA) or MMP-2 targeted siRNA (MMP-2 siRNA). * p≤0.05, Th i ifi t d i i ti i MMP 2 iRNA t f t d COL8+/+ SMCThere was a significant decrease in migration in MMP-2 siRNA transfected COL8+/+ SMCs compared to scrambled siRNA transfected COL8+/+ SMCs. † p≤0.05, There was a significant decrease in migration in MMP-2 and scrambled siRNA transfected COL8-/-
SMCs compared to MMP-2 and scrambled siRNA transfected COL8+/+ SMCs. Values are mean ± SEM.
115
1 0
200*
*
*COL8+/+ -MMP-2 siRNACOL8+/+ -Scr siRNA
COL8 / S iRNA
Time-lapse migration
Dis
tanc
e (u
m)
100
150COL8-/- -MMP-2 siRNACOL8-/- -Scr siRNA
D
0
50
Time (hr)
0 1 2 3 4 5 6 7 8
Figure 3 5 5 MMP-2 siRNA reduces COL8+/+ migration levels to those of COL8-/-Figure 3.5.5 MMP-2 siRNA reduces COL8 migration levels to those of COL8smooth muscle cells
Distance migrated by COL8+/+ and COL8-/- SMCs over a period of 8 hours after transfection of non-targeting scrambled siRNA (Scr siRNA), or MMP-2 targeted siRNA (MMP-2 siRNA). Reducing MMP-2 in COL8+/+ SMCs significantly reduced their migration to levels comparable to COL8-/- SMCs. There were no differences at any ti i t i i ti b t bl d d MMP 2 t f t d COL8 / SMC dtimepoint in migration between scrambled and MMP-2 transfected COL8-/- SMCs and MMP-2 siRNA transfected COL8+/+ SMCs. * p≤0.05, Scrambled and MMP-2 transfected COL8-/- SMCs and MMP-2 transfected COL8+/+ SMCs displayed decreased migration compared to scrambled siRNA transfected COL8+/+ SMCs. Values are mean ± SEM. N=4
116
MMP-2+/+180 *
Time-lapse migration
MMP-2-/-
MMP 2
(um
)
120
140
160
*
**
Dis
tanc
e
40
60
80
100 *
0 1 2 3 4 5 6 7 8
0
20
Time (hr)
Figure 3.5.6 MMP-2-/- smooth muscle cells display reduced rates of migrationDistance migration by MMP-2+/+ and MMP-2-/- SMCs over a period of 8 hours. * p≤0.05, MMP-2-/- SMCs displayed significantly reduced migration compared to MMP-2+/+ SMCs. Values are mean ± SEM N=4Values are mean ± SEM. N=4
117
Gel contraction
60
70 * †
ent C
ontr
actio
n
30
40
50
Perc
e
0
10
20 ‡ ‡
DOX - - - - + +
COL 8 - - + + - -
0
COL8+/+
438 8n= 33
Figure 3.5.7 COL8-/- smooth muscle cells display attenuated collagen gel contraction compared to COL8+/+ smooth muscle cells
Percent collagen gel contraction by COL8-/- and COL8+/+ SMCs treated with doxycycline
COL8
COL8-/-
(DOX) or exogenous type VIII collagen (COL 8). * p≤0.05, COL8-/- SMCs exhibited decreased gel contraction compared to COL8+/+ SMCs. † p≤0.05, Addition of exogenous type VIII collagen significantly increased gel contraction in COL8-/- SMCs. ‡p ≤0.05, Doxycycline inhibited gel contraction of both COL8+/+ and COL8-/- SMC. Values are mean ± SEM.
118
MMP-9
Lane 1 2 3 4
MMP-2
Densitometry
MMP-9 1.00 0.55 0.52 0.27
MMP-2 1.00 0.42 0.50 0.34
Figure 3.5.8 In the 3-dimensional collagen gel assay, COL8-/- smooth muscle cells produce less MMP-2 and MMP-9 than COL8+/+ smooth muscle cells
Representative gelatin zymogram containing conditioned media obtained from gel contraction experiments. Lane 1 and 2 contain conditioned media from gels containing CO 8+/+ S C 3 d 4 i di i d di f l i i CO 8 /COL8+/+ SMCs. Lane 3 and 4 contain conditioned media from gels containing COL8-/-
SMCs. Lanes 2 and 4 were taken from gels incubated in the presence of doxycycline. Decreased MMP-2 and MMP-9 activity was present in the conditioned media of COL8-/-
SMCs compared to COL8+/+ SMCs. Activity of both MMP-9 and MMP-2 was inhibited in the presence of doxycycline. N=3
119
COL8+/+ -Scr siRNA
COL8+/+ -MMP-2 siRNA
COL8-/- -Scr siRNA
COL8 / MMP 2 iRNA
Gel contraction
COL8-/- -MMP-2 siRNA
80*
40
60
Con
trac
tion
20
40
Perc
ent C
0n=3 333
Figure 3.5.9 MMP-2 siRNA reduces COL8+/+ gel contraction to that of COL8-/-
smooth muscle cells
T I ll l t ti t 24 h i COL8+/+ d COL8 / SMC t f t d ithType I collagen gel contraction at 24 hours in COL8+/+ and COL8-/- SMCs transfected with scramble (Scr) or MMP-2 siRNA. Values are mean ± SEM. * p≤0.05, Both Scr and MMP-2 siRNA transfected COL8-/- SMCs and MMP-2 siRNA transfected COL8+/+ SMCs exhibited decreased gel contraction compared to scramble siRNA transfected COL8+/+
SMCs.
120
Chapter 4
The contribution of type VIII collagen in response to wire injury of mouse arteries
121
4.1 Introduction
In vivo, type VIII collagen is upregulated following experimental arterial injury in
rats (Bendeck et al., 1996b; Sibinga et al., 1997; Plenz et al., 1999a) and rabbits (Plenz et
al., 1999a), yet its contribution to intimal hyperplasia is currently not known. As a result
of our findings reported in Chapters 2 and 3, which suggest type VIII collagen plays a
critical role in regulating smooth muscle cell migration and proliferation and MMP
production in vitro, we sought to determine the contribution of type VIII collagen to
arterial wound repair in vivo. We hypothesized that arterial injury in Col8a1-/-/Col8a2-/-
(COL8-/-) mice would result in decreased intimal hyperplasia compared to
Col8a1+/+/Col8a2+/+ (COL8+/+) mice. To test this hypothesis, we injured the carotid and
femoral arteries of COL8+/+ and COL8-/- mice and assayed the cell proliferation, MMP
activity, and morphological changes in injured arteries.
4.2 Materials and methods
4.2.1 Chemicals and reagents
All reagents were obtained from Sigma-Aldrich Inc. (St. Louis, Missouri, USA), except
where noted otherwise.
122
4.2.2 Animals
Mice with targeted deletions of both the Col8a1 and Col8a2 genes (COL8-/-) were
generated in the laboratory of Dr. Bjorn Olsen (Harvard Medical School) as described
(Hopfer et al., 2005) with wild-type littermate mice (COL8+/+) used as controls.
Genotypes were verified using extracted tail DNA and polymerase chain reaction (PCR)
for both the Col8a1 and Col8a2 alleles as described in section 3.2.2.
4.2.3 Carotid artery wire injury
Mice were anesthetized with 5% isofluorane in 1.5L/min oxygen and maintained
at 1.5-2% isofluorane in oxygen for the duration of the surgery. Before surgery was
performed, mice were injected subcutaneously with 0.1mg/kg body weight of
buprenorphine. A midline incision was performed over the trachea and the left common
carotid artery was exposed by blunt dissection. Sutures were looped on the external
carotid artery and on the common carotid artery to stop blood flow during the procedure.
Mouse carotids were injured using a 0.3mm diameter wire containing 0.6 mm diameter
epoxy resin beads prepared as previously described (Zhu et al., 2000). Using a P-IBSS
dissecting microscope (Nikon Canada Inc.; Mississauga, ON) to visualize the arteries, the
beaded wire was advanced proximally through an arteriotomy in the left external carotid
artery into the common carotid down to the aortic arch. The wire was pulled through the
artery four times, rotating 90º with each pass, to denude the endothelium. The left
external carotid artery was ligated after withdrawal of the wire. Surgery was complete
after verifying the restoration of pulsatile blood flow in the common carotid.
123
4.2.4 Femoral artery wire injury
Mice were anesthesized and given analgesia as described in section 3.2.3. The
left femoral artery was exposed by blunt dissection. Sutures were looped onto a proximal
portion of the femoral artery and a small branch of the femoral artery located between the
rectus femoris and vastus medialis muscles. A 0.38mm diameter straight spring wire was
advanced proximally through the arteriotomy in the small branch artery into the femoral
and advanced over 5mm toward the iliac artery. The wire was left in place for 1 minute
to denude the endothelium and dilate the artery. The small branch artery was ligated after
withdrawal of the wire. Surgery was complete after verifying restoration of pulsatile
blood flow in the femoral artery.
4.2.5 Carotid and femoral artery processing
On the day of sacrifice, mice were euthanized by an intraperitoneal injection of
333mg/kg body weight ketamine (Ayerst Veterinary Laboratories; Guelph, ON) and
67mg/kg body weight xylazine (Bayer, Inc.; Toronto, ON). The entire circulatory system
was perfusion-fixed at constant physiologic pressure via a catheter placed in the left
ventricle. First the circulation was perfused with 0.9% saline solution (Baxter Inc.;
Mississauga, ON), then with 4% paraformaldehyde for 10 minutes. The entire left
common carotid from the aortic arch to the bifurcation or the femoral artery from the iliac
artery to the ligated small branch artery was removed, placed in 4% paraformaldehyde for
2 hours, then transferred to PBS and processed. Processing was performed by the Centre
124
for Modeling Human Disease (CMHD) Pathology Core, the Toronto Centre for
Phenogenomics (Toronto, ON).
For processing of carotid arteries, arteries were bisected, paraffin-embedded, and
4 µm thick cross-sections were cut from the midsection of both carotid halves towards
the proximal/distal ends. All subsequent morphometric analysis was performed on the
sixth cross-section from each half, thereby giving an accurate representation of the extent
of injury along the arterial length and ensuring consistency in the areas analyzed between
mice.
For processing of femoral arteries, vessels were bisected and paraffin embedded.
Four µm thick cross-sections were cut from each bisected half to get an accurate
representation of injury along the length of the femoral and analysis was performed on
cross-sections from the middle of the femoral arteries.
4.2.6 Determination of the extent of denudation and re-
endothelization
To ensure that there was equal denudation between the groups, percent
denudation was calculated with the four day post-injury group also used to calculate Ki67
index (described later in section 4.2.7). Percent denudation was measured at four days
after injury from cross-sections using Simple PCI digital imaging software by dividing
the length of the internal elastic lamina devoid of endothelial cells by the total length of
the internal elastic lamina x 100%.
Carotid arterial sections at twenty-one days only were processed and stained by
the CMHD Pathology Core to detect von Willebrand factor (vWF), which is produced
125
constituitively in endothelial cells. Briefly, after antigen retrieval and blocking, sections
were stained with a 1:200 dilution of rabbit anti-vWF antibody (#A-0082,
DakoCytomation; Mississauga, ON) overnight, then stained with a 1:200 dilution of
biotin-conjugated goat anti-rabbit IgG secondary antibody (#BA-1000, Vector
Laboratories; Burlingame, CA). Following avidin-biotin complex formation, staining
was visualized with 3,3’-diaminobenzidine and sections were then counterstained with
hematoxylin. Percent re-endothelization was measured at twenty-one days after injury
from vWF-stained cross-sections by dividing the length of the internal elastic lamina
containing endothelial cells by the total length of the internal elastic lamina x 100%.
4.2.7 Immunostaining for Ki67
Cell proliferation was assessed at four and seven days after carotid artery wire
injury and at seven days after femoral artery wire injury. Sections were stained by the
CMHD Pathology Core to detect Ki67. Briefly, after antigen retrieval and blocking,
sections were stained with a 1:200 dilution of rabbit anti-Ki67 antibody (#RM-9106–S,
LabVision; Fremont, CA) overnight, then stained with a 1:200 dilution of biotin-
conjugated goat anti-rabbit IgG secondary antibody (#BA-1000, Vector Laboratories;
Burlingame, CA). Following avidin-biotin complex formation, staining was visualized
with 3,3’-diaminobenzidine and sections were then counterstained with hematoxylin.
The percentage of Ki67-labeled nuclei was measured in the medial and intimal layers of
the vessel using a Nikon Eclipse E600 microscope, Hamamatsu camera (model #C4742-
95), and Simple PCI software (Compix Inc.; Mars, PA).
126
4.2.8 Gelatin zymography
Seven days after carotid artery injury, mice were euthanized and the carotid
arteries were flushed with saline to remove blood and snap-frozen in liquid nitrogen.
Three to five carotids were pooled, and 5 independent pooled samples for the COL8-/- and
COL8+/+ mice were prepared by pulverizing under liquid nitrogen, and lysing in buffer
containing 1% SDS, 1mM PMSF, 10μg/ml leupeptin in 50mM Tris (pH 7.6). To detect
MMP activity, 5μg of protein was loaded into separate wells on an 8% SDS-
polyacrylamide (BioRad Laboratories; Hercules, CA) gel containing 0.1% gelatin as a
substrate and electrophoresis was performed as previously described in section 2.2.8. For
TIMP activity, reverse gelatin zymography was performed using methods similar to those
described above, with the exception that 0.13mg/ml of recombinant human MMP-2
(Chemicon International; Temecula, CA) was added to a 12% SDS-polyacrylamide gel.
Areas of TIMP activity were visualized as dark bands on a clear background (Bendeck
and Nakada, 2001). For comparison, samples were run alongside uninjured COL8+/+ and
COL8-/- mouse carotid artery samples as well as an injured rat carotid artery sample.
Densitometry was performed on the zymograms and reverse zymograms to determine
differences in MMP and TIMP activity using ImageJ Software (freeware from the
Research Services Branch of the National Institute of Mental Health, available at
http://rsb.info.nih.gov/ij/). Data was normalized to densitometry values for the injured rat
carotid artery and expressed as the fold change in activity compared to the injured rat
carotid artery. The same rat carotid sample was run on each gel so this procedure
corrected for variability between gels.
127
4.2.9 Immunoblotting
Protein extracts from seven day injured and uninjured carotid arteries were used
to perform Western blots measuring type VIII collagen protein after carotid artery injury.
10µg of protein from the carotids, as well as smooth muscle cell lysates from section
2.2.11, were subjected to 8% SDS-PAGE and transferred to a nitrocellulose membrane
(BioRad). Immunoblotting was performed as described in section 2.2.11.
Immunoblotting was performed at least twice with multiple samples.
4.2.10 Intimal hyperplasia
Twenty-one days after injury, mice were euthanized and carotid and femoral
arteries were processed and sections were stained by the CMHD Pathology Core with
hematoxylin and eosin. Cross-sectional area of the intima, media, and lumen, and the
perimeters of media, intima, and lumen were measured using Simple PCI. Medial area
was determined by measuring total area inside the external elastic lamina and subtracting
the area inside the internal elastic lamina (IEL). Intimal area was determined by
subtracting the total lumen area from the area inside the internal elastic lamina. Total
vessel wall area was determined by adding the medial and intimal areas. Lumen area was
determined by measuring the total lumen perimeter and calculating the area from the
perimeter measurement; outward remodeling was determined by measuring the perimeter
of the external elastic lamina to determine the total vessel diameter using the following
equations:
128
P=perimeter; r=radius; A=area; D=diameter
P=2πr A=πr2 D=2r
Therefore, A=P2/4π and D= P/π
4.2.11 Statistics
All surgeries, experiments, and measurements were performed with the
experimenter blind to the genotype of the mice. Data were analyzed by Student’s t-test
(SigmaStat v3.1, Systat Software Inc.; Point Richmond, CA) to compare group means
between the two genotypes, with a significance level of p≤0.05.
4.3 Results
4.3.1 Type VIII collagen was increased in injured carotid arteries
of COL8+/+ mice
Western blotting of protein extracts from carotid arteries taken seven days after
injury demonstrated the presence of type VIII collagen in the COL8+/+ carotid arteries
(Figure 4.5.1). There was a clear increase in the amount of type VIII collagen present
within injured COL8+/+ carotid arteries, compared to COL8+/+ uninjured vessels, where
there was little or no type VIII collagen detectable. There was no type VIII collagen
present within COL8-/- uninjured and injured carotid arteries.
129
4.3.2 The extent of injury was the same in both COL8-/- and
COL8+/+ mice
Extent of injury was measured as the percent of endothelial denudation from
carotid artery cross-sections harvested four days after injury (Figure 4.5.2A). Percent
endothelial denudation was not significantly different between COL8-/- and COL8+/+
mice, indicating that the extent of injury was comparable between genotypes.
Furthermore, examination of re-endothelization at twenty-one days after injury (Figure
4.5.2B) demonstrated that both COL8-/- and COL8+/+ carotid arteries were equally re-
endothelized.
4.3.3 There were no significant differences in smooth muscle
cell proliferation in injured carotid arteries between the COL8-/-
and COL8+/+ mice
Smooth muscle cell proliferation was assessed by immunostaining for Ki67, a
nuclear antigen associated with proliferation and present during the cell cycle but absent
during the resting G0 phase. Examination of uninjured COL8-/- (n=6) and COL8+/+ (n=4)
carotid arteries demonstrated an equivalent number of medial (85 ± 7 vs. 76 ± 13
respectively) and intimal cells (23 ± 6 vs. 24 ± 5, respectively) in both mouse groups
(Figure 4.5.3). The percentage of Ki67 positive cells in the vessel wall was calculated at
four and seven days after injury (for representative images, see Figure 4.5.4). There were
no significant differences in the Ki67 labeling index in the media at four days after injury
(Figure 4.5.5A) or in the total number of cells in the media between the COL8-/- and
130
COL8+/+ vessels (Figure 4.5.5B). There were also no significant differences in the Ki67
labeling index in the intima at four days after injury between the COL8-/- and COL8+/+
vessels (Figure 4.5.5A). Intimal cell number was not significantly different between
groups (Figure 4.5.5B). Seven days after injury, there were no significant differences in
the Ki67 labeling index in the media (Figure 4.5.6A) or in the number of medial cells
(Figure 4.5.4B). There were also no significant differences in the Ki67 labeling index in
the intima (Figure 4.5.6A) or in the number of intimal cells (Figure 4.5.6B) seven days
after injury. In uninjured vessels, there was little (<1%) to no cell proliferation in COL8-
/- and COL8+/+ carotid arteries.
4.3.4 There were no significant differences between MMP and
TIMP activity in injured carotid arteries from COL8-/- and COL8+/+
mice
Carotid arteries were harvested seven days after injury for zymogram analysis. A
total of five pooled injured carotid artery samples for COL8+/+ and five pooled samples
for COL8-/- mice were analyzed. Densitometry was performed on bands corresponding to
MMP-9A (95 kDa), MMP-2L (70 kDa), and MMP-2A (61 kDa) on the zymograms and
normalized to the densitometry of the respective bands for the injured rat carotid artery
sample. Uninjured vessels contained no MMP-9A and no MMP-2A, and only a small
amount of MMP-2L (Figure 4.5.7A), which was not significantly different between
COL8-/- and COL8+/+ carotid arteries (Figure 4.5.8A). MMP-9A, MMP-2L, and MMP-
2A were all expressed in the injured carotids (Figure 4.5.7A). Comparing COL8-/- to
131
COL8+/+ injured carotid arteries, we noted small decreases in activity of all these MMPs
in the COL8-/- arteries; however, these differences were not statistically significant
(Figure 4.5.8B-D).
Densitometry was not performed on the reverse zymogram gels; however, we
observed that the TIMP-2 (18kDa) and TIMP-1 (27 & 30kDa) activities in the arteries
after injury were similar between the COL8+/+ and COL8-/- groups (Figure 4.5.7B).
4.3.5 There were no significant differences in intimal
hyperplasia in injured carotid arteries from COL8+/+ and COL8-/-
mice
Twenty-one days after carotid artery wire injury, intimal area, medial area, total
vessel wall area, intima-to-media ratio, lumen area, and outward remodeling were
measured. In uninjured COL8-/- (n=5) and COL8+/+ (n=5) carotid arteries (Figure 4.5.3),
there were no significant differences in medial area (16,200 ± 700 µm2 vs. 15,100 ±
1,600 µm2, respectively), lumen area (115,000 ± 5,100 µm2 vs. 95,500 ± 5,800 µm2,
respectively), or vessel diameter (413 ± 9 µm vs. 391 ± 16 µm, respectively) between
mouse groups. There were no significant differences in intimal area between COL8-/- and
COL8+/+ injured arteries; however intimal area was nearly doubled in the COL8-/- group
compared to COL8+/+ (Figure 4.5.9A). There were no significant differences in medial
area between injured COL8-/- and COL8+/+ carotid arteries (Figure 4.5.9B). Also, in
many instances, it was difficult to distinguish intima from media, as the IEL was not
continuous; therefore, we calculated total vessel wall (medial + intimal) cross-sectional
132
area as a measurement of injury. There were no significant differences between total
vessel wall area (Figure 4.5.9C) or intima-to-media ratio (Figure 4.5.9D) between COL8-
/- and COL8+/+ mice. There were no significant differences in cross-sectional area of the
lumen between injured COL8-/- and COL8+/+ carotid arteries (Figure 4.5.10A). There
were also no significant differences in outward remodeling, measured as vessel diameter,
between injured COL8-/- and COL8+/+ carotid arteries (Figure 4.5.10B). Figure 4.5.11
contains representative images of carotid artery cross-sections from mice twenty-one
days after injury.
4.3.6 COL8-/- mice had increased medial proliferation after
femoral artery wire injury
In uninjured COL8-/- (n=5) and COL8+/+ (n=1) femoral arteries (Figure 4.5.12A),
there were equivalent numbers of total medial (87 ± 28 vs. 83 ± 0, respectively) or
intimal cells (22 ± 3 vs. 18 ± 0, respectively). Seven days after injury of the femoral
artery with a wire, cell proliferation in the media and intima was assessed by
immunostaining for Ki67 (for representative images, see Figure 4.5.12B). Proliferation
was significantly increased in the media of COL8-/- mice compared to COL8+/+ mice,
while there were no significant differences in proliferation in the intima (Figure 4.5.13A).
There were no differences in total cell number in the media or intima between COL8-/-
and COL8+/+ mice (Figure 4.5.13B). In uninjured vessels, there was little (<2%) to no
cell proliferation in COL8-/- and COL8+/+ femoral arteries.
133
4.3.7 COL8-/- mice demonstrated reduced outward remodeling
after femoral artery wire injury
Twenty-one days after femoral artery wire injury, intimal area, medial area, total
vessel wall area, intima-to-media ratio, lumen area, and outward remodeling were
measured and analyzed. In uninjured COL8-/- (n=5) and COL8+/+ (n=5) femoral arteries
(Figure 4.5.12A), there were no significant differences in medial area (10,200 ± 600 µm2
vs. 11,400 ± 1,000 µm2, respectively), lumen area (74,700 ± 13,800 µm2 vs. 65,700 ±
6,600 µm2, respectively), or vessel diameter (313 ± 9 µm vs. 311 ± 11 µm, respectively)
between mouse groups. There was a 35% reduction in intimal area in COL8-/- compared
to COL8+/+ femoral arteries, although this was not significant (Figure 4.5.14A). There
were no significant differences in medial area between injured COL8-/- and COL8+/+ mice
(Figure 4.5.14B). There were no significant differences in total vessel wall area (Figure
4.5.14C) or intima-to-media ratio (Figure 4.5.14D). While there were also no significant
differences in cross-sectional area of the lumen (Figure 4.5.145), vessel diameter was
significantly decreased in COL8-/- compared to COL8+/+ mice after femoral arterial
injury, indicating a decreased outward remodeling response. Figure 4.5.16 contains
representative images of femoral artery cross-sections from mice twenty-one days after
injury.
4.4 Discussion
Previous work showed type VIII collagen was increased following balloon injury
of the rat carotid (Bendeck et al., 1996b; Sibinga et al., 1997) and porcine coronary
134
arteries (Sinha et al., 2001), and is present in atherosclerotic lesions in mice (Yasuda et
al., 2000), rabbits (Plenz et al., 1999a), and humans (Macbeath et al., 1996; Weitkamp et
al., 1999; Plenz et al., 1999d); however, there is no previous information on any
differences in type VIII collagen expression after mechanical injury in mice. Ours is the
first study to show that type VIII collagen was increased in mouse carotids after wire
injury.
We chose to examine proliferation rates at four days and seven days after injury
of COL8+/+ and COL8-/- mouse carotid arteries, which approximately correspond to the
timepoints when medial and intimal cell replication peak in the mouse carotid wire injury
model (Lindner et al., 1993). Our data show that intimal smooth muscle cell replication
increased between four and seven days after injury, also consistent with previous studies
in this model (Lindner et al., 1993). However, there were no significant differences
between proliferation rates in COL8+/+ and COL8-/- mice, in either layer of the vessel
wall, and at either timepoint after injury, indicating that the absence of type VIII collagen
did not significantly affect smooth muscle cell proliferation after carotid artery injury.
By contrast, seven days after femoral artery injury, we found the medial cell proliferation
was increased in COL8-/- mice compared to COL8+/+ mice. However, this data should be
interpreted cautiously; the femoral artery wire injury is characterized by a rapid onset of
medial apoptosis followed by medial proliferation at seven days (Sata et al., 2000),
therefore, it is possible that apoptosis rates were higher in the COL8-/- mice, offsetting the
increased proliferation with the net result of no change in medial cell number. Another
difference is that the carotid artery wire injury model has minimal inflammation (Lindner
et al., 1993), while the femoral artery wire injury model has infiltration of inflammatory
135
cells (Sata et al., 2000). In fact, the total number of intimal cells present at seven days
after injury is four-fold greater in the femoral arteries compared to the carotids,
suggesting that inflammatory cells do contribute to intimal hyperplasia. However, we
have not identified the proliferating cells in these sections. Analysis of the cellular
composition of the femoral artery lesions, as well as perhaps a time-course examining
femoral artery lesion development following wire injury including reassessment of both
proliferation and apoptosis, is required to interpret our results.
The production of matrix-degrading enzymes such as the MMPs is required for
smooth muscle cells to detach from matrix to migrate or proliferate, and to facilitate the
clearance of matrix barriers. Our in vitro experiments revealed that there was less MMP-
2 activity in the conditioned media from the COL8-/- smooth muscle cells, which was
increased upon addition of exogenous type VIII collagen (Chapter 2 &(Adiguzel et al.,
2006). In contrast, we found no significant differences in the latent band of MMP-2L
between uninjured COL8+/+ and COL8-/- carotids; however, our in vitro experiments were
performed with smooth muscle cells that were migrating and proliferating, while smooth
muscle cells within the normal carotid artery are quiescent, which may explain our
contrasting results. We have also previously shown that type VIII collagen stimulated the
production of both MMP-2 and MMP-9 by rat smooth muscle cells (Hou et al., 2000). In
vivo after carotid injury, MMP-9 and MMP-2 activities were increased. However, there
were no significant differences in MMP-9 or MMP-2 activity comparing the COL8+/+ and
COL8-/- injured carotids, indicating there was little contribution of type VIII collagen to
stimulate MMP activity, at least after carotid artery injury in vivo. While our in vitro
results (Chapter 2) had demonstrated decreased MMP-2 in COL8-/- compared to COL8+/+
136
smooth muscle cells, our in vivo results were in contrast to our in vitro data,
demonstrating no differences in MMP-2 activity. Nonetheless, activation of MMP-2 is
regulated by both TIMP-2 and MT1-MMP (Nagase et al., 2006). The reverse zymogram
data showed no significant changes in TIMP-2 activity in COL8+/+ compared to COL8-/-
injured carotid arteries; however, we did not measure MT1-MMP levels, which may also
be altered and should be examined in the future.
Based on our in vitro studies in Chapters 2 and 3 showing decreased migration,
proliferation, and MMP levels in COL8-/- smooth muscle cells, we hypothesized that
intimal hyperplasia would be reduced in COL8-/- compared to COL8+/+ mice subjected to
carotid arterial injury. However, we found that there were no significant differences in
intimal or medial area after injury of carotid arteries in COL8+/+ and COL8-/- mice,
indicating that the absence of type VIII collagen did not affect the response to injury in
this vessel. There were also no significant differences in vessel diameter or lumen area
of the carotid arteries between genotypes, indicating that outward remodeling was not
affected by the absence of type VIII collagen.
In the carotid artery injury experiments we denuded the endothelium using a
beaded wire. In COL8+/+ mice at 4 days after injury, ~10% of the intimal surface was re-
endothelized, and at 21 days after injury, ~80% of vessel intimal surface was re-
endothelized, which is consistent with previous studies showing 10% re-endothelization
at 5 days after wire injury and nearly complete re-endothelization within 21 days after
injury of the mouse carotids (Lindner et al., 1993). Furthermore, there were no
significant differences in endothelization in the COL8-/- mice. A healthy endothelium
serves to maintain smooth muscle cell quiescence and regeneration of endothelium in a
137
denuded vessel attenuates the underlying intimal hyperplastic response by smooth muscle
cells (Fingerle et al., 1990). While both COL8-/- and COL8+/+ injured carotid arteries
demonstrated equivalent endothelization at four and twenty-one days after injury, we
cannot be certain that the rate of re-endothelization was the same in both genotypes. It is
known that type VIII collagen serves as an attachment factor for (Turner et al., 2006) and
promotes the proliferation of endothelial cells (Hopfer et al., 2005). We can speculate
that perhaps type VIII collagen promoted greater endothelial cell proliferation and
migration leading to an increased rate of re-endothelization in the COL8+/+ group, thereby
blunting the intimal hyperplastic response. Injured COL8-/- carotid arteries, by contrast,
may have re-endothelized at a slower rate, allowing for an increased amount of time the
smooth muscle cells can contribute to intimal hyperplasia following injury, resulting in
an equivalent intimal area between COL8-/- and COL8+/+ mice.
In contrast to the carotid artery injury, our results from the femoral artery wire
injury experiments were more promising and demonstrated significantly decreased
outward remodeling and a trend towards decreased intimal area in COL8-/- mice
compared to COL8+/+ mice 21 days after injury. The response to femoral artery wire
injury is normally characterized by vessel dilation and outward remodeling following the
procedure and persisting for at least 21 days (Sata et al., 2000). We did observe this
response in COL8+/+ mice, yet we did not observe any outward remodeling in the COL8-/-
mice. It is not completely clear why the femoral arteries of COL8-/- mice did not
remodel; however, we can speculate that this could be due to reduced MMP production in
these vessels and/or decreased intimal hyperplasia. Type VIII collagen stimulates smooth
muscle cell production of MMP-9 (Hou et al., 2000), an MMP which has been implicated
138
in mediating outward vessel remodeling in response to increased blood flow (Lessner et
al., 2004). However, we have not measured MMP activity in the femoral arteries. The
other important trend that we observed is towards decreased intimal thickening in the
COL8-/- mice. Though the difference between genotypes failed to reach statistical
significance, increasing the sample size would likely result in a significant result.
Therefore, it is also possible that because the COL8-/- femoral arteries developed about
35% less intimal thickening, perhaps they did not need to undergo outward remodeling to
compensate for lumen loss.
In general, our results were disappointing because we did not find major
differences in the arterial response to injury between COL8+/+ and COL8-/- mice. When
examining the in vivo response to injury, it is possible that there are other factors which
compensate for the absence of type VIII collagen. For example the production of matrix
proteins like osteopontin or fibronectin may be increased in the absence of type VIII
collagen, as these two proteins are known to stimulate smooth muscle cell migration
(Naito et al., 1990; Liaw et al., 1994) and MMP expression (Bendeck et al., 2000).
Another possible limitation of our studies was the choice of arterial injury model.
There are no foolproof models of arterial injury; all have their advantages and limitations
(reviewed by(Wang et al., 2006b). In our model, the carotid artery wire injury or
mechanical injury model, vessels re-endothelize within three weeks and develop small
intimal lesions (Lindner et al., 1993). Its advantages are that it is most similar to balloon
angioplasty and allows for the study of smooth muscle cell responses with minimal
thrombosis, inflammation, and no change in blood flow. However, the limitations are
that it generates only a small intimal response and is less reproducible than other models.
139
In the partial or complete artery ligation or blood flow cessation model, the carotid or
one/both of its major branches are ligated to cause decreased blood flow, resulting in
lumen narrowing, reduced vessel diameter, and a smooth muscle cell-rich intima (Kumar
and Lindner, 1997; Korshunov and Berk, 2003). It is easy to perform and very
reproducible; however, one major limitation is that it does not mimic physiological
vascular interventions and results are also variable due to sudden changes in blood flow.
With the vein graft model, large veins from donor mice are grafted between two ends of a
cut common carotid, resulting in 80% lumen narrowing at 16 weeks with minimal
thrombosis (Zou et al., 1998). It is advantageous as it reproducibly results in intimas
similar to those seen in human vein grafts; however, the study is in veins and not arteries
and the initial vein diameters are much larger than those of the common carotid. Another
model is the electric injury model, where an electric current is passed through the arterial
wall, destroying all cells, and resulting in rapid cell growth and re-endothelization within
two weeks (Carmeliet et al., 1997). It is quick and easy to perform; however, this model
involves severe thrombosis, inflammation, and matrix remodeling.
Examining the differences between these models (summarized in Table 3.5.1), we
chose to use the carotid artery wire injury model as it is physiologically relevant, results
in an injury with minimal inflammation, thrombosis, and change in blood flow, and is
similar to other models used extensively in our laboratory. However, we did not find any
significant differences between COL8+/+ and COL8-/- mice with the carotid artery wire
injury while perhaps differences may have been noted between the mouse groups with
another mouse model of intimal hyperplasia. We believe the mice were adequately
injured, as the carotid arteries were nearly completely denuded early after injury, and
140
stained positive for Ki67 indicating that cells were proliferating. Also injured carotid
arteries expressed more MMP activity than uninjured carotid arteries, and there was
intimal hyperplasia evident in both injured mouse groups.
However, we observed a very large amount of variability in the amount of intimal
hyperplasia in injured carotid arteries from COL8-/- and COL8+/+ mice. In both groups,
we found some animals that had an extensive intimal growth response, (which may have
been attributable to disruptions of the IEL), whereas other animals appeared to have no
intimal growth. Later, we found evidence in the literature that the C57Bl/6 strain of mice
(the background strain of the COL8-/- and COL8+/+ mice) is resistant to intimal
hyperplasia in response to wire injury of the carotid arteries (Kuhel et al., 2002). Further
review of the literature demonstrated that in contrast to the carotid artery wire injury
model, the femoral artery wire injury model results in a concentric intimal hyperplasia
response that is induced to an equal extent in all strains of mice examined, including
C57Bl/6 (Sata et al., 2000). In collaboration with Dr. Scott Heximer (University of
Toronto), we performed femoral artery wire injuries on COL8-/- and COL8+/+ mice. We
observed a significant decrease in outward remodeling and consistent reduction in intimal
area in the COL8-/- arteries compared to COL8+/+ arteries without the large amount of
variability that was observed with the carotid arteries. Furthermore, the intima-to-media
ratio was three-fold higher in the femoral artery injury compared to the carotid artery
injury in COL8+/+ mice, suggesting that the femoral artery injury model is much more
robust. Therefore, we believe that with an increase in sample size, a statistically
significant difference in intimal area may be revealed. However, we cannot discount the
141
possibility that there is really no difference in response to injury between the COL8+/+
and COL8-/- mice.
Acknowledgements
We would like to thank Steven Gu from Dr. Scott Heximer’s laboratory for performing
the femoral artery wire injuries and Helen Su, an undergraduate student working in our
laboratory, for analysis of the femoral artery wire injury data. We would also like to
thank Jiwan Dhaliwal for the injured rat carotid artery samples used to identify MMPs.
142
4.5 Figures and tables
143
Type VIII Collagen
β-actinβ-actin
Figure 4.5.1 Type VIII collagen is increased in COL8+/+ mouse carotid arteries after wire injury
Representative Western blot demonstrating that type VIII collagen was increased 7 days after injury in COL8+/+ mouse carotid arteries compared to uninjured carotid arteries. Type j y p j ypVIII collagen was absent in arteries from the COL8-/- mice.
144
100
A
Four days after carotid artery injury
85
90
95
100he
lial D
enud
atio
n
70
75
80
% E
ndot
h
n=12 9
100
B
Twenty-one days after carotid artery injuryCOL8+/+
COL8-/-
60
80
ndot
heliz
atio
n
0
20
40
4n=5
% R
e-E
n
Figure 4.5.2 Extent of injury is the same in COL8-/- and COL8+/+ mice
Percent endothelial denudation was measured in vessel cross-sections at 4 days after injury as an indication of extent of injury (A). Percent re-endothelization was measured from vessel cross-sections at 21 days after injury (B). There were no differences in the extent of injury between COL8+/+ and COL8-/- mice. Values are mean + SEM.
145
COL8+/+ COL8-/-
Figure 4.5.3 Images of cross-sections from uninjured COL8-/- and COL8+/+ carotid arteries
Representative images of uninjured carotid arteries of COL8+/+ and COL8-/- mice. Scalebar is 100μm
146
A Four days after carotid artery injury
COL8+/+ COL8-/-
B Seven days after carotid artery injury
COL8+/+ COL8-/-
Figure 4.5.4 Images of cross-sections from COL8-/- and COL8+/+ carotid arteries four and seven days after injury
Representative images of Ki-67 stained carotid arteries of COL8+/+ and COL8-/- mice at four (A) and seven days (B) after injury. Scalebar is 100μm
25
A
Four days after carotid artery injury
147
posi
tive
cells
15
20
% K
i67
p
0
5
10
17 1617 16n=
Media Intima0
B
Four days after carotid artery injury COL8+/+
cells
150
200
250 COL8-/-
Num
ber
of
50
100
Figure 4.5.5 There were no differences in proliferation or total cell number between COL8-/- and COL8+/+ mice four days after carotid artery wire injury
017 16
Media Intima
17 16n=
There were no significant differences in the Ki67 labelling index in the media or intima between COL8+/+ and COL8-/- mice (A). There were no significant differences in total number of cells in the media or intima between COL8+/+ and COL8-/- mice (B). Values are mean +SEM.
60
A
Seven days after carotid artery injury
148
30
40
50
ositi
ve c
ells
10
20
30
% K
i67
p
6 56 5n=
Media Intima0
B
Seven days after carotid artery injuryCOL8+/+
200
250
300
of c
ells
COL8-/-
50
100
150
Num
ber
o
6 5 6 5
Figure 4.5.6 There were no differences in proliferation or total cell number between COL8-/- and COL8+/+ mice seven days after carotid artery wire injury
06 5 6 5
Media Intima
n=
There were no differences in the Ki67 labeling index in the media or intima between COL8+/+ and COL8-/- mice (A). There were no differences in total number of cells in the media or intima between COL8+/+ and COL8-/- mice (B). Values are mean + SEM.
A
149
MMP-9A
MMP-2LMMP-2A
B
TIMP-1
TIMP-2
Figure 4.5.7 There were no differences in MMP or TIMP activity between COL8-/-Figure 4.5.7 There were no differences in MMP or TIMP activity between COL8and COL8+/+ mice after carotid artery injury
Representative zymogram of pooled injured (COL8+/+ & COL8-/- ) & uninjured (COL8+/+
& COL8-/- ) carotid arteries (A). Rat Sample refers to a balloon-injured rat carotid artery used for the normalization of lytic bands. Representative reverse zymogram of pooled injured (COL8+/+ & COL8-/- ) & uninjured (COL8+/+ & COL8-/- ) carotid arteries (B). Rat Sample refers to a balloon injured rat carotid artery used for reference Three to fiveSample refers to a balloon-injured rat carotid artery used for reference. Three to five carotids were pooled to generate each COL8+/+ and COL8-/- sample, 5 pooled samples were analyzed in total.
A B
150
COL8+/+
COL8 /MMP-9A DensitometryUninjured MMP-2L Densitometry
COL8-/-y
4
5
6
zed
Valu
e
1 5
2
2.5
3
ized
Val
ue
0
1
2
3
Nor
mal
iz
5 5n=0
0.5
1
1.5
Nor
mal
i
2 2n=
C D
MMP-2L Densitometry
9
MMP-2A Densitometry
2
45678
mal
ized
Val
ue
0 81
1.21.41.61.8
mal
ized
Val
ue
0123
Nor
m
5 5n=0
0.20.40.60.8
Nor
m
5 5n=
Figure 4.5.8 There were no differences in gelatinase activity between COL8-/- and COL8+/+ mice after carotid artery injury
There were no significant differences in the bands for latent MMP-2 in uninjured carotid arteries (A) or the bands for active MMP-9 (B), latent MMP-2 (C), or active MMP-2 (D) in injured COL8-/- and COL8+/+ carotid arteries seven days after wire injury. Band density measurements were normalized to the density of band for the same MMP in a balloon-injured rat carotid sample included in each gel. Values are mean + SEM.
A BMedial AreaIntimal Area
151
COL8+/+
30000
40000
50000
60000
20000
25000
30000
35000
um2 )
um2 )
COL8-/-
0
10000
20000
16 220
5000
10000
15000
Are
a (u
16 22
Are
a (u
n= n=
C D
Total Vessel Wall Area
60000
700000 8
Intima to Media Ratio
20000
30000
40000
50000
60000
Are
a (u
m2 )
0 2
0.4
0.6
0.8
Rat
io
0
10000
20000
16 220
0.216 22n= n=
Figure 4.5.9 There were no significant differences in vessel wall hypertrophy after carotid artery injury between COL8-/- and COL8+/+ mice
There were no significant differences in cross sectional areas of the intima (A) mediaThere were no significant differences in cross-sectional areas of the intima (A), media (B), total vessel wall (C), or intima-to-media ratio (D) between injured COL8+/+ and COL8-/- mouse carotid arteries 21 days after injury. Values are mean + SEM.
A B
Outward RemodelingLumen Area
152
Outward Remodeling
400
500
600
met
er (u
m)
150000
200000
250000
(um
2 )
0
100
200
300
Vess
el D
iam
16 220
50000
100000
16 22
Are
a
n= n=
COL8+/+
COL8-/-
Figure 4.5.10 There were no differences in lumen size or outward remodeling between COL8-/- and COL8+/+ mice after carotid artery injury
There were no significant differences in cross-sectional area of the lumen between injured COL8+/+ and COL8-/- mice 21 days after carotid artery wire injury (A). There were no differences in outward remodeling between injured COL8+/+ and COL8-/- mice (B). Values are mean + SEM.
153
COL8+/+ COL8-/-
Figure 4.5.11 Images of cross-sections from COL8-/- and COL8+/+ carotid arteries twenty-one days after injury
Representative images of carotid arteries of COL8+/+ and COL8-/- twenty-one days after injury. Dashed lines indicate the internal elastic lamina. Scalebar is 100μm
154
COL8+/+ COL8-/-
A
COL8+/+ COL8-/-
B
Figure 4.5.12 Images of cross-sections from uninjured and injured COL8-/- and COL8+/+ femoral arteries
Representative images of uninjured femoral arteries of COL8+/+ and COL8-/- mice (A). Representative images of Ki-67 stained femoral arteries of COL8+/+ and COL8-/- mice at seven days after injury (B). Scalebar is 100μmy j y ( ) μ
A
Seven days after femoral artery injury
155
50
sitiv
e ce
lls
*30
40
50
% K
i67
pos
10
20
B
Seven days after femoral artery injury
Media
300
Intima0
5 555
IntimaMedia
COL8+/+
COL8 /
n=
mbe
r of
cel
ls
300
150
200
250
COL8-/-
Num
Intima
5 5
X Data
0
50
100
5 5
Media
n=
Figure 4.5.13 COL8-/- mice demonstrated increased proliferation in the media seven days after femoral artery injury compared to COL8+/+ mice
There was a significant (* p≤0.05) increase in the Ki67 labeling index in the media of COL8-/- mice compared to COL8+/+ mice but no significant difference in Ki67 labeling index in the intima (A). There were no differences in total number of cells in the media or intima between COL8+/+ and COL8-/- mice (B). Values are mean + SEM.
A B Medial AreaIntimal Area
156
COL8+/+
COL8 /
rea
(um
2 )
800010000120001400016000
15000
20000
25000
rea
(um
2 )
COL8-/-
Ar
0200040006000
0
5000
10000A
1515 1515n= n=
C DTotal Vessel Wall Area
30000
350001.0
1.2Intima to Media Ratio
Are
a (u
m2 )
5000
10000
15000
20000
25000
0.2
0.4
0.6
0.8
Rat
io
0
5000 15150
1515n= n=
Figure 4.5.14 There were no significant differences in vessel wall hypertrophy after femoral artery injury between COL8-/- and COL8+/+ mice
There were no significant differences in cross-sectional areas of the intima (A), media (B), total vessel wall (C), or intima-to-media ratio (D) between injured COL8-/- and COL8+/+ mouse femoral arteries 21 days after injury. Values are mean + SEM.
A BOutward RemodelingLumen Area
157
30000
40000
50000
60000
ea (u
m2 )
200
300
400
iam
eter
(um
)
*
0
10000
20000Are
15150
100
Vess
el D
i
1515n= n=
COL8+/+
COL8-/-
Figure 4.5.15 COL8-/- mice demonstrated attenuated outward remodeling after femoral injury
There were no significant differences in cross-sectional area of the lumen between injured COL8+/+ and COL8-/- mice 21 days after femoral artery wire injury (A). * p≤0.05, COL8-/- mice demonstrated significantly decreased outward remodeling compared to COL8+/+ mice 21 days after injury (B). Values are mean + SEM.
158
COL8+/+ COL8-/-
Figure 4.5.16 Images of cross sections from COL8-/- and COL8+/+ femoral arteries twenty-one days after injurytwenty one days after injury
Representative images of femoral arteries of COL8+/+ and COL8-/- mice twenty-one days after injury. Vessel diameter was decreased in COL8-/- compared to COL8+/+
femoral arteries after injury. Dashed lines indicate the internal elastic lamina. Scalebar is 100μm
159
Table 4.5.1 Comparison of different mouse injury models of intimal hyperplasia Adapted from (Wang et al., 2006b) Blood flow
cessation Mechanical (wire) injury
Vein graft Electric injury
Surgical Difficulty
+ ++ +++ +
Variation + +++ ++ ++
Physiological relevance
+ +++ +++ -
Endothelial injury
No Yes Maybe Yes
Medial injury No Yes Maybe Yes
Volume flow Changed Unchanged Unchanged Unchanged
Thrombotic occlusion
+ ++ + ++++
Intimal hyperplasia
+++ ++ + +++
160
Chapter 5
General discussion and future directions
161
5.1 The effects of type VIII collagen on migration and
proliferation
Type VIII collagen is upregulated in atherosclerosis in humans (Macbeath et al.,
1996; Weitkamp et al., 1999; Plenz et al., 1999d) and in animal models of atherosclerosis
and restenosis (Bendeck et al., 1996b; Sibinga et al., 1997; Plenz et al., 1999a; Plenz et
al., 1999b; Yasuda et al., 2000; Sinha et al., 2001). While collagen production in general
is greatly increased during atherosclerosis and following arterial injury, it was previously
noted that the increase in type VIII collagen mRNA and protein expression is far greater
than the modest increase in type I collagen (Sibinga et al., 1997). The study presented in
Chapter 2 is the first to examine the effects of type VIII collagen in a type I collagen-rich
environment, as would be found within the vessel wall. By utilizing COL8-/- aortic
smooth muscle cells and comparing them to COL8+/+ smooth muscle cells, we found that
the lack of endogenous type VIII collagen resulted in significant deficiencies in migration
and proliferation.
In accord with results presented in Chapter 2, COL8-/- corneal endothelial cells
also displayed decreased proliferation compared to COL8+/+ corneal endothelial cells
(Hopfer et al., 2005). Similarly, we demonstrated an increase in cell size of the COL8-/-
smooth muscle cells (Chapter 2), which was also described in COL8-/- compared to
COL8+/+ corneal endothelial cells (Hopfer et al., 2005). Type VIII collagen is
upregulated during angiogenesis (Sage et al., 1984; Sage and Iruela-Arispe, 1990; Iruela-
Arispe and Sage, 1991; Smith J et al., 1996) and can serve as an adhesive substrate for
vascular endothelial cells, ligating the same integrin receptors as smooth muscle cells
162
(Turner et al., 2006) . Furthermore, since type VIII collagen is an adhesive substrate for
astrocytes, stimulates their migration, and is also upregulated during glial scar formation
following cold brain injury (Hirano et al., 2004), it is tempting to infer that it serves as a
stimulatory substrate for several cell types during the response to biological injury.
For cell migration to occur, cells must first extend lamellipodia in one direction,
form new adhesions, undergo contraction, and release adhesions at the opposite end to
move forward (Lauffenburger and Horwitz, 1996). COL8-/- smooth muscle cells
demonstrated significantly increased adhesion to type I collagen compared to COL8+/+
smooth muscle cells and significantly decreased levels of cell migration on type I
collagen compared to COL8+/+ smooth muscle cells. Theoretically, if COL8-/- smooth
muscle cells adhere too strongly to a surface, they may not be able to migrate. In fact,
strong levels of adhesion were found to correlate with high levels of activity of the small
GTPase Rho (Cox et al., 2001). This leads to phosphorylation of MLCP, thereby
inhibiting it, resulting in increased myosin light chain activity, contraction, and decreased
migration (Somlyo and Somlyo, 2000). Increased activity of Rho could also cause
increased formation of actin stress fibers (Hall, 1998), similar to what we observed in the
COL8-/- smooth muscle cells.
These findings indicate the need for further investigation of the receptors for type
VIII collagen and the signaling pathways activated by type VIII collagen in smooth
muscle cells. Previous work has shown that there are three receptors for type VIII
collagen on smooth muscle cells: α2β1 and α1β1 integrins (Hou et al., 2000) and DDR1
(Hou et al., 2001). Treatment with integrin blocking antibodies decreased smooth muscle
cell adhesion to type VIII collagen (Hou et al., 2000), but DDR1-deficient smooth muscle
163
cells also displayed decreased adhesion to type VIII collagen and decreased proliferation
when stimulated with type VIII collagen. Furthermore, DDR1-deficient smooth muscle
cells had decreased levels of MMP-2 activity (Hou et al., 2001), similar to our COL8-/-
smooth muscle cells. Since DDR1 contains consensus sequences for the SH2 domains of
Nck (a tyrosine kinase adaptor protein), GTP-activating protein, and the p85 subunit of
PI3K (Vogel, 1999), which are proteins also involved in integrin signaling (Giancotti and
Ruoslahti, 1999), there is a possibility of cross-talk between the DDR1 and integrin
signaling pathways. However, in human mammary carcinoma cells, DDR1 is active even
in the presence of integrin blocking antibodies, indicating an integrin-independent
signaling pathway also exists (Vogel et al., 2000). The PI3K pathway can activate cyclin
D through the rac pathway, and therefore might influence cell proliferation mediated by
either integrins or DDRs (Giancotti and Ruoslahti, 1999).
First and foremost, it would be prudent to examine the absolute amounts and
proportions of the α2β1 and α1β1 integrins and DDR1 present on the COL8-/- and COL8+/+
smooth muscle cells to confirm that the noted discrepancies in the phenotypes of the
smooth muscle cells were not due to differences in the expression of type VIII collagen
receptors. Next, we cannot underestimate the value of the COL8-/- smooth muscle cells
in examining the cellular changes caused by type VIII collagen. We have shown that
administration of exogenous type VIII collagen decreased adhesion and increased
migration and MMP-2 activity in the COL8-/- smooth muscle cells. Utilizing this system,
future research should focus on elucidating the molecules involved in type VIII collagen
signaling. Our data demonstrating decreased MMP-2 in COL8-/- SMCs suggests multiple
signaling intermediates to examine, with the most ideal candidates being the Rho
164
GTPases involved in cell migration, Cdc42, Rac1, and RhoA. It has been proposed that
there is a hierarchy of signaling where Cdc42 is upstream of Rac1 which antagonizes
RhoA signaling (Nobes and Hall, 1995). Studies have shown that increased RhoA
activity decreases MMP-2 expression (Ispanovic et al., 2008) while increased Cdc42
(Ispanovic et al., 2008) and Rac1 (Westermarck and Kahari, 1999) activity increases
expression of MMP-2. The next steps would be to determine differences in the activation
of the cdc42, rac, and Rho GTPases in the COL8-/- and COL8+/+ smooth muscle cells,
which can easily be performed with commercially-available kits to assay the amount of
activated Rho GTPase compared to total levels of Rho GTPase. The necessity of
signaling pathways, such as the PI3K or Rho pathway, can then be assayed by utilizing
readily-available inhibitors of these molecules, such as wortmannin or dominant
negative/constituitively active Rho constructs, respectively.
5.2 Regulation of MMP-2 and migration by type VIII
collagen
Type VIII collagen is known to stimulate smooth muscle cell migration (Sibinga
et al., 1997; Hou et al., 2000) and upregulate MMP-2 and MMP-9 expression (Hou et al.,
2000) in vitro. In the first set of experiments with the COL8-/- smooth muscle cells
described in Chapter 2, we examined differences in MMP-2 activity between the COL8-/-
and COL8+/+ smooth muscle cells. The purpose of the experiments in Chapter 3 was to
determine the contribution of MMP-2 to the type VIII collagen-induced migratory
response. One novel finding from Chapter 3 is that decreasing the amount of endogenous
165
MMP-2 in COL8+/+ smooth muscle cells by RNA silencing was sufficient to recapitulate
the COL8-/- smooth muscle cell phenotype of decreased chemotaxis towards PDGF-BB,
decreased random cell migration in culture, and decreased contraction of type I collagen
gels. Another novel finding of this work is that addition of exogenous type VIII collagen
to COL8-/- smooth muscle cells is able to increase the levels of chemotaxis, migration,
and gel contraction to those of the COL8+/+ smooth muscle cells.
Several exciting areas of research are suggested by this work. One future
experiment would be to determine if increasing MMP-2 in the COL8-/- smooth muscle
cells, either by addition of recombinant MMP-2 or by transfection with an MMP-2
expression vector, would be sufficient to rescue the phenotype in a manner similar to the
addition of exogenous type VIII collagen. Secondly, we found that COL8-/- smooth
muscle cells were impaired in their chemotaxis towards PDGF-BB. PDGF-BB is able to
directly stimulate MMP-2 expression in smooth muscle cells (Uzui et al., 2000; Borrelli
et al., 2006; Risinger, Jr. et al., 2006), which suggests that increased migration due to
increased MMP-2 expression will occur in the presence of PDGF-BB. To clearly
eliminate any possibility of differential activation of the PDGF-BB receptor (PDGFRβ),
the phosphorylation status of the receptor upon ligation should be measured in both
COL8-/- and COL8+/+ smooth muscle cells. If activation of the PDGFRβ is equivalent in
both cell types, this would suggest that there is cross-talk between the PDGFRβ,
integrins, and possibly DDRs as well, acting downstream of the PDGFRβ. Type I
collagen has been found to synergistically enhance smooth muscle cell proliferation in
response to PDGF-BB through src-dependent cross-talk with the α2β1 integrin
(Hollenbeck et al., 2004). Since type VIII collagen is also a ligand for the α2β1 integrin,
166
it too may enhance signaling of the PDGFRβ and perhaps exert its effects on increased
proliferation or increased MMP-2 expression. For example, smooth muscle cells could
be stimulated with PDGF-BB and/or type VIII collagen and the effects on proliferation
examined. While both stimuli should increase smooth muscle cell proliferation, if the
combined effects resulted in an enhancement of proliferation greater than the addition of
the mitogenic effects together, this would indicate synergistic activation. The amount of
phosphorylation of intracellular signaling intermediates or the PDGFRβ should also be
examined to determine if there is synergy in signaling. Using functional antibodies to
block the α2β1 integrin and examining the activation status of the PDGFRβ would
determine whether activation of the integrin was required for the synergistic effect and
using inhibitors of signaling intermediates such as src or Erk1/2 would determine where
the two pathways converge. Furthermore, both the PDGFRβ (DeMali et al., 1999) and
DDR1 (Koo et al., 2006; Wang et al., 2006a) associate with the SH2 domain containing
phosphatase SHP-2, indicating another potential level of cross-talk that type VIII
collagen may induce. Due to the lack of DDR1 inhibitors, mutant DDR1 constructs
could be utilized to block DDR1 signaling to examine cross-talk pathways involving type
VIII collagen.
5.3 Contribution of type VIII collagen in the arterial
response to mechanical injury
Previous studies in the pig (Sinha et al., 2001) and rat (Bendeck et al., 1996b;
Sibinga et al., 1997) after carotid artery balloon catheter injury and during the
167
progression of atherosclerosis in cholesterol-fed rabbits (Plenz et al., 1999a; Plenz et al.,
1999b) and apoE-deficient mice (Yasuda et al., 2001) have demonstrated increases in
type VIII collagen expression. The in vivo experiments reported in Chapter 4 are the first
to show that type VIII collagen is increased after carotid artery wire injury in the mouse
and the first to examine the contribution of type VIII collagen to the response to arterial
wire injury.
We were not able to detect a significant difference in intimal formation, cell
proliferation, or MMP expression after wire injury of the carotid arteries between COL8-/-
and COL8+/+ mice. We speculate that this may have been due to the extreme variability
noted in intimal hyperplasia in both the COL8-/- and COL8+/+ mice, despite our large
sample size. Upon discovering that the background strain of the type VIII collagen
mouse, C57Bl/6, is very resistant to intimal hyperplasia following carotid artery injury
(Kuhel et al., 2002), we decided to perform mechanical injuries of the femoral artery,
which have been shown previously to cause a robust and reproducible intimal thickening
response in C57Bl/6 mice (Sata et al., 2000). COL8-/- injured femoral arteries underwent
significantly less outward remodeling and there was a consistent trend toward a decrease
in the extent of intimal hyperplasia in the COL8-/- compared to the COL8+/+ injured
femoral arteries. Future experiments to increase the sample size for these experiments
and to assess type VIII collagen expression and MMP/TIMP production in the injured
femoral arteries are now required. Furthermore, to test the hypothesis that type VIII
collagen stimulated outward remodeling of the vessel after injury, type VIII collagen
should be administered to COL8-/- femoral arteries immediately after injuring to
determine if there is an effect. However, in the event that the results remain insignificant,
168
then the conclusion would be that, in the context of restenosis, type VIII collagen does
not have a measurable effect.
While restenosis and atherosclerosis involve many similar mechanisms, the role
of type VIII collagen may be different in each disease. In light of the research
demonstrating increases in type VIII collagen expression and accumulation in animal
models of atherosclerosis (Plenz et al., 1999a; Plenz et al., 1999b; Yasuda et al., 2001), a
logical future direction for research would be to cross the COL8-/- mice with LDLR-
deficient mice and examine the extent of atherosclerosis development and the cellular and
extracellular composition of vascular lesions. COL8-/-; LDLR-/- mice and COL8+/+;
LDLR-/- mice mice would be placed on a high fat diet for 12 or 24 weeks.
Measurements would be made of total plaque burden and plaque area, percent of lesional
areas occupied by smooth muscle cells and macrophages, and the collagen and elastin
content of plaques would be measured.
Our studies have concentrated on smooth muscle cells as the source of type VIII
collagen, and focused on smooth muscle cell interactions with this protein. However,
other cell types in the vessel wall produce type VIII collagen in the atherosclerotic
plaque, including endothelial cells (Iruela-Arispe et al., 1991) and macrophages
(Weitkamp et al., 1999). Furthermore, the consensus of previous work in the mouse
model is that there are few inflammatory cells in the lesions of carotid artery wire injury
models (Lindner et al., 1993), while there is some infiltration of macrophages in the
femoral artery wire injury model (Roque et al., 2000). At this time, very little is known
about the relative contributions of each cell type to the amount of type VIII collagen
produced in the arterial wall, which would be an interesting avenue for further research.
169
Utilizing reciprocal bone marrow transplantation in COL8-/-; LDLR-/- mice and COL8+/+;
LDLR-/- mice, the contribution of macrophages and smooth muscle cells to total type VIII
collagen production in the vessel wall, as well as the influence of vessel wall
macrophage-derived type VIII collagen on lesion progression would be examined.
The findings from this thesis, combined with previous results from our laboratory
and the Owens’ laboratory (Hou et al., 2000; Hou et al., 2001; Pidkovka et al., 2007;
Cherepanova et al., 2009) suggest a role for type VIII collagen in the cascade of events
leading to development of vascular occlusive disease as follows: oxidized phospholipids
accumulate within the vascular wall and cause nuclear translocation of Klf4, leading to
both the phenotypic switching of smooth muscle cells and the upregulation of type VIII
collagen. In turn, type VIII collagen, signaling through the α2β1 and α1β1 integrins and
DDR1 receptor, upregulates MMP-2 expression and activity, which facilitates smooth
muscle cell migration and proliferation (Figure 5.5.1). The experiments demonstrating
that type VIII collagen is necessary for chemotactic migration towards oxidized
phospholipids were performed in vitro, utilizing our COL8+/+ and COL8-/- smooth muscle
cells (Cherepanova et al., 2009). Oxidized phospholipids were also demonstrated to
induce MMP-2 activity, which was required to induce smooth muscle cell proliferation,
as proliferation was inhibited in MMP-2 deficient smooth muscle cells (Auge et al.,
2004). It would be interesting to determine if type VIII collagen is necessary for the
immediate and direct effects of oxidized phospholipids in stimulating smooth muscle cell
growth and activation in vivo. Our laboratory is beginning experiments to examine this
by analyzing smooth muscle cell proliferation, MMP levels, and intimal formation
following periadventitial administration of oxidized phospholipids to the carotid and
170
femoral arteries of COL8+/+ and COL8-/- mice in a pluronic gel. Oxidized phospholipids
can activate the phenotypic switching of smooth muscle cells from a quiescent,
contractile state to a proliferative, synthetic state (Pidkovka et al., 2007) and also
upregulate type VIII collagen expression (Cherepanova et al., 2009). What is not known
is whether type VIII collagen is simply a marker of activated smooth muscle cells, or can
itself induce the phenotypic switching of smooth muscle cells. This could be examined
by analyzing the expression of smooth muscle cell differentiation markers in COL8-/-
compared to COL8+/+ smooth muscle cells, both with and without stimulation by
exogenous type VIII collagen. These experiments could be performed in vitro, and in
vivo with the administration of exogenous type VIII collagen to arteries in a pluronic gel.
5.4 Conclusion
Before completion of this thesis, very little was known about the functional role
of type VIII collagen in vascular occlusive disease. This thesis has made significant
advances in elucidating the biological functions of type VIII collagen. In summary, we
have shown that the production of type VIII collagen confers a migratory and mitogenic
phenotype to smooth muscle cells, and can dramatically affect their production of MMP-
2. While we have determined some of the functions of type VIII collagen, we have also
opened the door to much future research. Future experiments concentrating on
uncovering the intracellular signaling pathways activated by type VIII collagen in vitro,
and investigating the role of type VIII collagen in vivo, will further our understanding of
this molecule and perhaps serve as a gateway to the development of therapeutics for
atherosclerosis and restenosis. Also, extrapolating our results in the vascular system to
171
pathologies in other systems involving type VIII collagen to better our understanding is
exciting, but should be done with caution and carefully planned experiments to reveal all
functions of type VIII collagen within the body.
172
5.5 Figures
173
AMac
ROS?
OxPLs Type VIII Collagen
ROS
Phenotypic switching ?
Klf4
Klf4
yp g(Contractile→Synthetic)
B SMC proliferation & migration
↑↑ MMP-2
g
Figure 5.5.1 The role of type VIII collagen in smooth muscle cells in vascular occlusive disease
A simplified role for type VIII collagen in atherogenesis in SMCs is suggested as follows: oxidized phospholipids (OxPLs) accumulating within the vessel wall cause the nuclear translocation of the transcription factor Klf4, resulting in the phenotypic
it hi f SMC d th d ti f t VIII ll (A) Wh th t VIIIswitching of SMCs and the production of type VIII collagen (A). Whether type VIII collagen can stimulate macrophages or a possible link between SMC phenotypic transition and type VIII collagen is currently unknown (dashed arrows). Type VIII collagen then signals through integrins and DDR1 to upregulate MMP-2 expression and activity (B), which facilitates matrix degradation and SMC migration into the intima. Mac=macrophage, ROS=reactive oxygen species
174
References
Abedi, H and Zachary, I (1995) Signalling mechanisms in the regulation of vascular cell migration. Cardiovasc Res 30(4):544-556.
Adiguzel, E, Ahmad, PJ, Franco, C, and Bendeck, MP (2009) Collagens in the progression and complications of atherosclerosis. Vasc Med 14(1):73-89.
Adiguzel, E, Hou, G, Mulholland, D, Hopfer, U, Fukai, N, Olsen, B, and Bendeck, M (2006) Migration and growth are attenuated in vascular smooth muscle cells with type VIII collagen-null alleles. Arterioscler Thromb Vasc Biol 26(1):56-61.
Allen, WE, Zicha, D, Ridley, AJ, and Jones, GE (1998) A role for Cdc42 in macrophage chemotaxis. J Cell Biol 141(5):1147-1157.
Amano, M, Fukata, Y, and Kaibuchi, K (2000) Regulation and functions of Rho-associated kinase. Exp Cell Res 261(1):44-51.
Asakura, T and Karino, T (1990) Flow patterns and spatial distribution of atherosclerotic lesions in human coronary arteries. Circ Res 66(4):1045-1066.
Auge, N, Maupas-Schwalm, F, Elbaz, M, Thiers, JC, Waysbort, A, Itohara, S, Krell, HW, Salvayre, R, and Negre-Salvayre, A (2004) Role for matrix metalloproteinase-2 in oxidized low-density lipoprotein-induced activation of the sphingomyelin/ceramide pathway and smooth muscle cell proliferation. Circulation 110(5):571-578.
Avivi-Green, C, Singal, M, and Vogel, WF (2006) Discoidin domain receptor 1-deficient mice are resistant to bleomycin-induced lung fibrosis. Am J Respir Crit Care Med 174(4):420-427.
Axel, DI, Kunert, W, Goggelmann, C, Oberhoff, M, Herdeg, C, Kuttner, A, Wild, DH, Brehm, BR, Riessen, R, Koveker, G, and Karsch, KR (1997) Paclitaxel inhibits arterial smooth muscle cell proliferation and migration in vitro and in vivo using local drug delivery. Circulation 96(2):636-645.
Bailly, M, Wyckoff, J, Bouzahzah, B, Hammerman, R, Sylvestre, V, Cammer, M, Pestell, R, and Segall, JE (2000) Epidermal growth factor receptor distribution during chemotactic responses. Mol Biol Cell 11(11):3873-3883.
Ballestrem, C, Wehrle-Haller, B, Hinz, B, and Imhof, BA (2000) Actin-dependent lamellipodia formation and microtubule-dependent tail retraction control-directed cell migration. Mol Biol Cell 11(9):2999-3012.
Baron, JH, Moiseeva, EP, de Bono, DP, Abrams, KR, and Gershlick, AH (2000) Inhibition of vascular smooth muscle cell adhesion and migration by c7E3 Fab
175
(abciximab): a possible mechanism for influencing restenosis. Cardiovasc Res 48(3):464-472.
Belkin, VM, Belkin, AM, and Koteliansky, VE (1990) Human smooth muscle VLA-1 integrin: purification, substrate specificity, localization in aorta, and expression during development. J Cell Biol 111(5 Pt 1):2159-2170.
Bendeck, MP, Conte, M, Zhang, M, Nili, N, Strauss, BH, and Farwell, SM (2002) Doxycycline modulates smooth muscle cell growth, migration, and matrix remodeling after arterial injury. Am J Pathol 160(3):1089-1095.
Bendeck, MP, Irvin, C, Reidy, M, Smith, L, Mulholland, D, Horton, M, and Giachelli, CM (2000) Smooth muscle cell matrix metalloproteinase production is stimulated via alpha(v)beta(3) integrin. Arterioscler Thromb Vasc Biol 20(6):1467-1472.
Bendeck, MP, Irvin, C, and Reidy, MA (1996a) Inhibition of matrix metalloproteinase activity inhibits smooth muscle cell migration but not neointimal thickening after arterial injury. Circ Res 78(1):38-43.
Bendeck, MP and Nakada, MT (2001) The beta3 integrin antagonist m7E3 reduces matrix metalloproteinase activity and smooth muscle cell migration. J Vasc Res 38(6):590-599.
Bendeck, MP, Regenass, S, Tom, WD, Giachelli, CM, Schwartz, SM, Hart, C, and Reidy, MA (1996b) Differential expression of alpha 1 type VIII collagen in injured platelet-derived growth factor-BB--stimulated rat carotid arteries. Circ Res 79(3):524-531.
Bendeck, MP, Zempo, N, Clowes, AW, Galardy, RE, and Reidy, MA (1994) Smooth muscle cell migration and matrix metalloproteinase expression after arterial injury in the rat. Circ Res 75(3):539-545.
Benya, PD and Padilla, SR (1986) Isolation and characterization of type VIII collagen synthesized by cultured rabbit corneal endothelial cells. A conventional structure replaces the interrupted-helix model. J Biol Chem 261(9):4160-4169.
Bergers, G, Brekken, R, McMahon, G, Vu, TH, Itoh, T, Tamaki, K, Tanzawa, K, Thorpe, P, Itohara, S, Werb, Z, and Hanahan, D (2000) Matrix metalloproteinase-9 triggers the angiogenic switch during carcinogenesis. Nat Cell Biol 2(10):737-744.
Bilato, C, Curto, KA, Monticone, RE, Pauly, RR, White, AJ, and Crow, MT (1997) The inhibition of vascular smooth muscle cell migration by peptide and antibody antagonists of the alphavbeta3 integrin complex is reversed by activated calcium/calmodulin- dependent protein kinase II. J Clin Invest 100(3):693-704.
Bingley, JA, Hayward, IP, Campbell, GR, and Campbell, JH (2001) Relationship of glycosaminoglycan and matrix changes to vascular smooth muscle cell phenotype modulation in rabbit arteries after acute injury. J Vasc Surg 33(1):155-164.
176
Bingley, JA, Hayward, IP, Campbell, JH, and Campbell, GR (1998) Arterial heparan sulfate proteoglycans inhibit vascular smooth muscle cell proliferation and phenotype change in vitro and neointimal formation in vivo. J Vasc Surg 28(2):308-318.
Biswas, S, Munier, FL, Yardley, J, Hart-Holden, N, Perveen, R, Cousin, P, Sutphin, JE, Noble, B, Batterbury, M, Kielty, C, Hackett, A, Bonshek, R, Ridgway, A, McLeod, D, Sheffield, VC, Stone, EM, Schorderet, DF, and Black, GC (2001) Missense mutations in COL8A2, the gene encoding the alpha2 chain of type VIII collagen, cause two forms of corneal endothelial dystrophy. Hum Mol Genet 10(21):2415-2423.
Borrelli, V, di, ML, Sapienza, P, Colasanti, M, Moroni, E, and Cavallaro, A (2006) Role of platelet-derived growth factor and transforming growth factor beta1 the in the regulation of metalloproteinase expressions. Surgery 140(3):454-463.
Brown, SL, Lundgren, CH, Nordt, T, and Fujii, S (1994) Stimulation of migration of human aortic smooth muscle cells by vitronectin: implications for atherosclerosis. Cardiovasc Res 28(12):1815-1820.
Bulinski, JC and Gundersen, GG (1991) Stabilization of post-translational modification of microtubules during cellular morphogenesis. Bioessays 13(6):285-293.
Campbell, JH and Campbell, GR (1993) Culture techniques and their applications to studies of vascular smooth muscle. Clin Sci (Lond) 85(5):501-513.
Campbell, JH and Campbell, GR (1994a) Cell biology of atherosclerosis. J Hypertens Suppl 12(10):S129-S132.
Campbell, JH and Campbell, GR (1994b) The role of smooth muscle cells in atherosclerosis. Curr Opin Lipidol 5(5):323-330.
Carmeliet, P, Moons, L, Stassen, JM, De, MM, Bouche, A, van den Oord, JJ, Kockx, M, and Collen, D (1997) Vascular wound healing and neointima formation induced by perivascular electric injury in mice. Am J Pathol 150(2):761-776.
Carragher, NO, Levkau, B, Ross, R, and Raines, EW (1999) Degraded collagen fragments promote rapid disassembly of smooth muscle focal adhesions that correlates with cleavage of pp125(FAK), paxillin, and talin. J Cell Biol 147(3):619-630.
Cheng, L, Mantile, G, Pauly, R, Nater, C, Felici, A, Monticone, R, Bilato, C, Gluzband, YA, Crow, MT, Stetler-Stevenson, W, and Capogrossi, MC (1998) Adenovirus-mediated gene transfer of the human tissue inhibitor of metalloproteinase-2 blocks vascular smooth muscle cell invasiveness in vitro and modulates neointimal development in vivo. Circulation 98(20):2195-2201.
Cherepanova, OA, Pidkovka, NA, Sarmento, OF, Yoshida, T, Gan, Q, Adiguzel, E, Bendeck, MP, Berliner, J, Leitinger, N, and Owens, GK (2009) Oxidized Phospholipids Induce Type VIII Collagen Expression and Vascular Smooth Muscle Cell Migration. Circ Res
177
Choi, ET, Engel, L, Callow, AD, Sun, S, Trachtenberg, J, Santoro, S, and Ryan, US (1994) Inhibition of neointimal hyperplasia by blocking alpha V beta 3 integrin with a small peptide antagonist GpenGRGDSPCA. J Vasc Surg 19(1):125-134.
Clemmons, DR, Horvitz, G, Engleman, W, Nichols, T, Moralez, A, and Nickols, GA (1999) Synthetic alphaVbeta3 antagonists inhibit insulin-like growth factor-I-stimulated smooth muscle cell migration and replication. Endocrinology 140(10):4616-4621.
Clowes, AW, Clowes, MM, and Reidy, MA (1986) Kinetics of cellular proliferation after arterial injury. III. Endothelial and smooth muscle growth in chronically denuded vessels. Lab Invest 54(3):295-303.
Clowes, AW, Reidy, MA, and Clowes, MM (1983a) Mechanisms of stenosis after arterial injury. Lab Invest 49(2):208-215.
Clowes, AW, Reidy, MA, and Clowes, MM (1983b) Kinetics of cellular proliferation after arterial injury. I. Smooth muscle growth in the absence of endothelium. Lab Invest 49(3):327-333.
Corjay, MH, Diamond, SM, Schlingmann, KL, Gibbs, SK, Stoltenborg, JK, and Racanelli, AL (1999) alphavbeta3, alphavbeta5, and osteopontin are coordinately upregulated at early time points in a rabbit model of neointima formation. J Cell Biochem 75(3):492-504.
Cox, EA and Huttenlocher, A (1998) Regulation of integrin-mediated adhesion during cell migration. Microsc Res Tech 43(5):412-419.
Cox, EA, Sastry, SK, and Huttenlocher, A (2001) Integrin-mediated adhesion regulates cell polarity and membrane protrusion through the Rho family of GTPases. Mol Biol Cell 12(2):265-277.
Dadlani, H, Ballinger, ML, Osman, N, Getachew, R, and Little, PJ (2008) Smad and p38 MAP kinase-mediated signaling of proteoglycan synthesis in vascular smooth muscle. J Biol Chem 283(12):7844-7852.
Degani, S, Balzac, F, Brancaccio, M, Guazzone, S, Retta, SF, Silengo, L, Eva, A, and Tarone, G (2002) The integrin cytoplasmic domain-associated protein ICAP-1 binds and regulates Rho family GTPases during cell spreading. J Cell Biol 156(2):377-387.
DeMali, KA, Balciunaite, E, and Kazlauskas, A (1999) Integrins enhance platelet-derived growth factor (PDGF)-dependent responses by altering the signal relay enzymes that are recruited to the PDGF beta receptor. J Biol Chem 274(28):19551-19558.
Dilley, RJ, McGeachie, JK, and Prendergast, FJ (1987) A review of the proliferative behaviour, morphology and phenotypes of vascular smooth muscle. Atherosclerosis 63(2-3):99-107.
178
DiMilla, PA, Barbee, K, and Lauffenburger, DA (1991) Mathematical model for the effects of adhesion and mechanics on cell migration speed. Biophys J 60(1):15-37.
Dollery, CM and Libby, P (2006) Atherosclerosis and proteinase activation. Cardiovasc Res 69(3):625-635.
Dufourcq, P, Louis, H, Moreau, C, Daret, D, Boisseau, MR, Lamaziere, JM, and Bonnet, J (1998) Vitronectin expression and interaction with receptors in smooth muscle cells from human atheromatous plaque. Arterioscler Thromb Vasc Biol 18(2):168-176.
Eliceiri, BP (2001) Integrin and growth factor receptor crosstalk. Circ Res 89(12):1104-1110.
Evanko, SP, Angello, JC, and Wight, TN (1999) Formation of hyaluronan- and versican-rich pericellular matrix is required for proliferation and migration of vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 19(4):1004-1013.
Ezratty, EJ, Partridge, MA, and Gundersen, GG (2005) Microtubule-induced focal adhesion disassembly is mediated by dynamin and focal adhesion kinase. Nat Cell Biol 7(6):581-590.
Ferri, N, Carragher, NO, and Raines, EW (2004) Role of discoidin domain receptors 1 and 2 in human smooth muscle cell-mediated collagen remodeling: potential implications in atherosclerosis and lymphangioleiomyomatosis. Am J Pathol 164(5):1575-1585.
Fingerle, J, Au, YP, Clowes, AW, and Reidy, MA (1990) Intimal lesion formation in rat carotid arteries after endothelial denudation in absence of medial injury. Arteriosclerosis 10(6):1082-1087.
Flamant, M, Placier, S, Rodenas, A, Curat, CA, Vogel, WF, Chatziantoniou, C, and Dussaule, JC (2006) Discoidin domain receptor 1 null mice are protected against hypertension-induced renal disease. J Am Soc Nephrol 17(12):3374-3381.
Foehr, ED, Tatavos, A, Tanabe, E, Raffioni, S, Goetz, S, Dimarco, E, De, LM, and Bradshaw, RA (2000) Discoidin domain receptor 1 (DDR1) signaling in PC12 cells: activation of juxtamembrane domains in PDGFR/DDR/TrkA chimeric receptors. FASEB J 14(7):973-981.
Forough, R, Koyama, N, Hasenstab, D, Lea, H, Clowes, M, Nikkari, ST, and Clowes, AW (1996) Overexpression of tissue inhibitor of matrix metalloproteinase-1 inhibits vascular smooth muscle cell functions in vitro and in vivo. Circ Res 79(4):812-820.
Franco, C, Ho, B, Mulholland, D, Hou, G, Islam, M, Donaldson, K, and Bendeck, MP (2006) Doxycycline alters vascular smooth muscle cell adhesion, migration, and reorganization of fibrillar collagen matrices. Am J Pathol 168(5):1697-1709.
Franco, C, Hou, G, Ahmad, PJ, Fu, EY, Koh, L, Vogel, WF, and Bendeck, MP (2008) Discoidin domain receptor 1 (ddr1) deletion decreases atherosclerosis by accelerating
179
matrix accumulation and reducing inflammation in low-density lipoprotein receptor-deficient mice. Circ Res 102(10):1202-1211.
Fukata, M, Nakagawa, M, and Kaibuchi, K (2003) Roles of Rho-family GTPases in cell polarisation and directional migration. Curr Opin Cell Biol 15(5):590-597.
Garl, PJ, Wenzlau, JM, Walker, HA, Whitelock, JM, Costell, M, and Weiser-Evans, MC (2004) Perlecan-induced suppression of smooth muscle cell proliferation is mediated through increased activity of the tumor suppressor PTEN. Circ Res 94(2):175-183.
Gerth, J, Cohen, CD, Hopfer, U, Lindenmeyer, MT, Sommer, M, Grone, HJ, and Wolf, G (2007) Collagen type VIII expression in human diabetic nephropathy. Eur J Clin Invest 37(10):767-773.
Giancotti, FG and Ruoslahti, E (1999) Integrin signaling. Science 285(5430):1028-1032.
Gilmore, AP and Romer, LH (1996) Inhibition of focal adhesion kinase (FAK) signaling in focal adhesions decreases cell motility and proliferation. Mol Biol Cell 7(8):1209-1224.
Gottsch, JD, Sundin, OH, Liu, SH, Jun, AS, Broman, KW, Stark, WJ, Vito, EC, Narang, AK, Thompson, JM, and Magovern, M (2005a) Inheritance of a novel COL8A2 mutation defines a distinct early-onset subtype of fuchs corneal dystrophy. Invest Ophthalmol Vis Sci 46(6):1934-1939.
Gottsch, JD, Zhang, C, Sundin, OH, Bell, WR, Stark, WJ, and Green, WR (2005b) Fuchs corneal dystrophy: aberrant collagen distribution in an L450W mutant of the COL8A2 gene. Invest Ophthalmol Vis Sci 46(12):4504-4511.
Gotwals, PJ, Chi-Rosso, G, Lindner, V, Yang, J, Ling, L, Fawell, SE, and Koteliansky, VE (1996) The alpha1beta1 integrin is expressed during neointima formation in rat arteries and mediates collagen matrix reorganization. J Clin Invest 97(11):2469-2477.
Greenhill, NS, Ruger, BM, Hasan, Q, and Davis, PF (2000) The alpha1(VIII) and alpha2(VIII) collagen chains form two distinct homotrimeric proteins in vivo. Matrix Biol 19(1):19-28.
Guerin, C, Laterra, J, Masnyk, T, Golub, LM, and Brem, H (1992) Selective endothelial growth inhibition by tetracyclines that inhibit collagenase. Biochem Biophys Res Commun 188(2):740-745.
Haas, TL, Davis, SJ, and Madri, JA (1998) Three-dimensional type I collagen lattices induce coordinate expression of matrix metalloproteinases MT1-MMP and MMP-2 in microvascular endothelial cells. J Biol Chem 273(6):3604-3610.
Hall, A (1998) Rho GTPases and the actin cytoskeleton. Science 279(5350):509-514.
180
Hansson, GK and Libby, P (2006) The immune response in atherosclerosis: a double-edged sword. Nat Rev Immunol 6(7):508-519.
Hauck, CR, Hsia, DA, and Schlaepfer, DD (2000) Focal adhesion kinase facilitates platelet-derived growth factor-BB-stimulated ERK2 activation required for chemotaxis migration of vascular smooth muscle cells. J Biol Chem 275(52):41092-41099.
Hayward, IP, Bridle, KR, Campbell, GR, Underwood, PA, and Campbell, JH (1995) Effect of extracellular matrix proteins on vascular smooth muscle cell phenotype. Cell Biol Int 19(10):839-846.
He, JZ, Quan, A, Xu, Y, Teoh, H, Wang, G, Fish, JE, Steer, BM, Itohara, S, Marsden, PA, Davidge, ST, and Ward, ME (2007) Induction of matrix metalloproteinase-2 enhances systemic arterial contraction after hypoxia. Am J Physiol Heart Circ Physiol 292(1):H684-H693.
Hedin, U, Bottger, BA, Forsberg, E, Johansson, S, and Thyberg, J (1988) Diverse effects of fibronectin and laminin on phenotypic properties of cultured arterial smooth muscle cells. J Cell Biol 107(1):307-319.
Hillis, GS, Mlynski, RA, Simpson, JG, and MacLeod, AM (1998) The expression of beta 1 integrins in human coronary artery. Basic Res Cardiol 93(4):295-302.
Hinek, A, Boyle, J, and Rabinovitch, M (1992) Vascular smooth muscle cell detachment from elastin and migration through elastic laminae is promoted by chondroitin sulfate-induced "shedding" of the 67-kDa cell surface elastin binding protein. Exp Cell Res 203(2):344-353.
Hirano, S, Yonezawa, T, Hasegawa, H, Hattori, S, Greenhill, NS, Davis, PF, Sage, EH, and Ninomiya, Y (2004) Astrocytes express type VIII collagen during the repair process of brain cold injury. Biochem Biophys Res Commun 317(2):437-443.
Hollenbeck, ST, Itoh, H, Louie, O, Faries, PL, Liu, B, and Kent, KC (2004) Type I collagen synergistically enhances PDGF-induced smooth muscle cell proliferation through pp60src-dependent crosstalk between the alpha2beta1 integrin and PDGFbeta receptor. Biochem Biophys Res Commun 325(1):328-337.
Hopfer, U, Fukai, N, Hopfer, H, Wolf, G, Joyce, N, Li, E, and Olsen, BR (2005) Targeted disruption of Col8a1 and Col8a2 genes in mice leads to anterior segment abnormalities in the eye. FASEB J 19(10):1232-1244.
Hou, G, Mulholland, D, Gronska, MA, and Bendeck, MP (2000) Type VIII collagen stimulates smooth muscle cell migration and matrix metalloproteinase synthesis after arterial injury. Am J Pathol 156(2):467-476.
Hou, G, Vogel, W, and Bendeck, MP (2001) The discoidin domain receptor tyrosine kinase DDR1 in arterial wound repair. J Clin Invest 107(6):727-735.
181
Huang, R, Merrilees, MJ, Braun, K, Beaumont, B, Lemire, J, Clowes, AW, Hinek, A, and Wight, TN (2006) Inhibition of versican synthesis by antisense alters smooth muscle cell phenotype and induces elastic fiber formation in vitro and in neointima after vessel injury. Circ Res 98(3):370-377.
Humphries, JD, Byron, A, and Humphries, MJ (2006) Integrin ligands at a glance. J Cell Sci 119(Pt 19):3901-3903.
Hwang, JY, Johnson, PY, Braun, KR, Hinek, A, Fischer, JW, O'Brien, KD, Starcher, B, Clowes, AW, Merrilees, MJ, and Wight, TN (2008) Retrovirally mediated overexpression of glycosaminoglycan-deficient biglycan in arterial smooth muscle cells induces tropoelastin synthesis and elastic fiber formation in vitro and in neointimae after vascular injury. Am J Pathol 173(6):1919-1928.
Hynes, RO (2002) Integrins: bidirectional, allosteric signaling machines. Cell 110(6):673-687.
Ichii, T, Koyama, H, Tanaka, S, Kim, S, Shioi, A, Okuno, Y, Raines, EW, Iwao, H, Otani, S, and Nishizawa, Y (2001) Fibrillar collagen specifically regulates human vascular smooth muscle cell genes involved in cellular responses and the pericellular matrix environment. Circ Res 88(5):460-467.
Ikari, Y, Yee, KO, and Schwartz, SM (2000) Role of alpha5beta1 and alphavbeta3 integrins on smooth muscle cell spreading and migration in fibrin gels. Thromb Haemost 84(4):701-705.
Illidge, C, Kielty, C, and Shuttleworth, A (1998) The alpha1(VIII) and alpha2(VIII) chains of type VIII collagen can form stable homotrimeric molecules. J Biol Chem 273(34):22091-22095.
Illidge, C, Kielty, C, and Shuttleworth, A (2001) Type VIII collagen: heterotrimeric chain association. Int J Biochem Cell Biol 33(5):521-529.
Iozzo, RV (1998) Matrix proteoglycans: from molecular design to cellular function. Annu Rev Biochem 67:609-652.
Iruela-Arispe, ML, Diglio, CA, and Sage, EH (1991) Modulation of extracellular matrix proteins by endothelial cells undergoing angiogenesis in vitro. Arterioscler Thromb 11(4):805-815.
Iruela-Arispe, ML and Sage, EH (1991) Expression of type VIII collagen during morphogenesis of the chicken and mouse heart. Dev Biol 144(1):107-118.
Ishibashi, S, Goldstein, JL, Brown, MS, Herz, J, and Burns, DK (1994) Massive xanthomatosis and atherosclerosis in cholesterol-fed low density lipoprotein receptor-negative mice. J Clin Invest 93(5):1885-1893.
182
Ishizaki, T, Morishima, Y, Okamoto, M, Furuyashiki, T, Kato, T, and Narumiya, S (2001) Coordination of microtubules and the actin cytoskeleton by the Rho effector mDia1. Nat Cell Biol 3(1):8-14.
Ispanovic, E, Serio, D, and Haas, TL (2008) Cdc42 and RhoA have opposing roles in regulating membrane type 1-matrix metalloproteinase localization and matrix metalloproteinase-2 activation. Am J Physiol Cell Physiol 295(3):C600-C610.
Itoh, T, Ikeda, T, Gomi, H, Nakao, S, Suzuki, T, and Itohara, S (1997) Unaltered secretion of beta-amyloid precursor protein in gelatinase A (matrix metalloproteinase 2)-deficient mice. J Biol Chem 272(36):22389-22392.
Jackson, CL, Raines, EW, Ross, R, and Reidy, MA (1993) Role of endogenous platelet-derived growth factor in arterial smooth muscle cell migration after balloon catheter injury. Arterioscler Thromb 13(8):1218-1226.
Jenkins, G (2008) The role of proteases in transforming growth factor-beta activation. Int J Biochem Cell Biol 40(6-7):1068-1078.
Johnson, C and Galis, ZS (2004) Matrix metalloproteinase-2 and -9 differentially regulate smooth muscle cell migration and cell-mediated collagen organization. Arterioscler Thromb Vasc Biol 24(1):54-60.
Johnson, JL, George, SJ, Newby, AC, and Jackson, CL (2005) Divergent effects of matrix metalloproteinases 3, 7, 9, and 12 on atherosclerotic plaque stability in mouse brachiocephalic arteries. Proc Natl Acad Sci U S A 102(43):15575-15580.
Jones, JI, Prevette, T, Gockerman, A, and Clemmons, DR (1996) Ligand occupancy of the alpha-V-beta3 integrin is necessary for smooth muscle cells to migrate in response to insulin-like growth factor. Proc Natl Acad Sci U S A 93(6):2482-2487.
Kapoor, R, Bornstein, P, and Sage, EH (1986) Type VIII collagen from bovine Descemet's membrane: structural characterization of a triple-helical domain. Biochemistry 25(13):3930-3937.
Kapoor, R, Sakai, LY, Funk, S, Roux, E, Bornstein, P, and Sage, EH (1988) Type VIII collagen has a restricted distribution in specialized extracellular matrices. J Cell Biol 107(2):721-730.
Katsuda, S, Okada, Y, Minamoto, T, Oda, Y, Matsui, Y, and Nakanishi, I (1992) Collagens in human atherosclerosis. Immunohistochemical analysis using collagen type-specific antibodies. Arterioscler Thromb 12(4):494-502.
Kimura, T, Abe, K, Shizuta, S, Odashiro, K, Yoshida, Y, Sakai, K, Kaitani, K, Inoue, K, Nakagawa, Y, Yokoi, H, Iwabuchi, M, Hamasaki, N, Nosaka, H, and Nobuyoshi, M (2002) Long-term clinical and angiographic follow-up after coronary stent placement in native coronary arteries. Circulation 105(25):2986-2991.
183
Kittelberger, R, Davis, PF, Flynn, DW, and Greenhill, NS (1990) Distribution of type VIII collagen in tissues: an immunohistochemical study. Connect Tissue Res 24(3-4):303-318.
Konitsiotis, AD, Raynal, N, Bihan, D, Hohenester, E, Farndale, RW, and Leitinger, B (2008) Characterization of high affinity binding motifs for the discoidin domain receptor DDR2 in collagen. J Biol Chem 283(11):6861-6868.
Koo, DH, McFadden, C, Huang, Y, Abdulhussein, R, Friese-Hamim, M, and Vogel, WF (2006) Pinpointing phosphotyrosine-dependent interactions downstream of the collagen receptor DDR1. FEBS Lett 580(1):15-22.
Korshunov, VA and Berk, BC (2003) Flow-induced vascular remodeling in the mouse: a model for carotid intima-media thickening. Arterioscler Thromb Vasc Biol 23(12):2185-2191.
Koyama, H, Raines, EW, Bornfeldt, KE, Roberts, JM, and Ross, R (1996) Fibrillar collagen inhibits arterial smooth muscle proliferation through regulation of Cdk2 inhibitors. Cell 87(6):1069-1078.
Krylyshkina, O, Anderson, KI, Kaverina, I, Upmann, I, Manstein, DJ, Small, JV, and Toomre, DK (2003) Nanometer targeting of microtubules to focal adhesions. J Cell Biol 161(5):853-859.
Kuhel, DG, Zhu, B, Witte, DP, and Hui, DY (2002) Distinction in genetic determinants for injury-induced neointimal hyperplasia and diet-induced atherosclerosis in inbred mice. Arterioscler Thromb Vasc Biol 22(6):955-960.
Kumar, A and Lindner, V (1997) Remodeling with neointima formation in the mouse carotid artery after cessation of blood flow. Arterioscler Thromb Vasc Biol 17(10):2238-2244.
Kundra, V, Escobedo, JA, Kazlauskas, A, Kim, HK, Rhee, SG, Williams, LT, and Zetter, BR (1994) Regulation of chemotaxis by the platelet-derived growth factor receptor-beta. Nature 367(6462):474-476.
Kuzuya, M, Kanda, S, Sasaki, T, Tamaya-Mori, N, Cheng, XW, Itoh, T, Itohara, S, and Iguchi, A (2003) Deficiency of gelatinase a suppresses smooth muscle cell invasion and development of experimental intimal hyperplasia. Circulation 108(11):1375-1381.
Kuzuya, M, Nakamura, K, Sasaki, T, Cheng, XW, Itohara, S, and Iguchi, A (2006) Effect of MMP-2 deficiency on atherosclerotic lesion formation in apoE-deficient mice. Arterioscler Thromb Vasc Biol 26(5):1120-1125.
Kvansakul, M, Bogin, O, Hohenester, E, and Yayon, A (2003) Crystal structure of the collagen alpha1(VIII) NC1 trimer. Matrix Biol 22(2):145-152.
184
Labrador, JP, Azcoitia, V, Tuckermann, J, Lin, C, Olaso, E, Manes, S, Bruckner, K, Goergen, JL, Lemke, G, Yancopoulos, G, Angel, P, Martinez, C, and Klein, R (2001) The collagen receptor DDR2 regulates proliferation and its elimination leads to dwarfism. EMBO Rep 2(5):446-452.
Lafont, A, Durand, E, Samuel, JL, Besse, B, Addad, F, Levy, BI, Desnos, M, Guerot, C, and Boulanger, CM (1999) Endothelial dysfunction and collagen accumulation: two independent factors for restenosis and constrictive remodeling after experimental angioplasty. Circulation 100(10):1109-1115.
Langholz, O, Rockel, D, Mauch, C, Kozlowska, E, Bank, I, Krieg, T, and Eckes, B (1995) Collagen and collagenase gene expression in three-dimensional collagen lattices are differentially regulated by alpha 1 beta 1 and alpha 2 beta 1 integrins. J Cell Biol 131(6 Pt 2):1903-1915.
Lauffenburger, DA and Horwitz, AF (1996) Cell migration: a physically integrated molecular process. Cell 84(3):359-369.
Legate, KR, Wickstrom, SA, and Fassler, R (2009) Genetic and cell biological analysis of integrin outside-in signaling. Genes Dev 23(4):397-418.
Leitinger, B (2003) Molecular analysis of collagen binding by the human discoidin domain receptors, DDR1 and DDR2. Identification of collagen binding sites in DDR2. J Biol Chem 278(19):16761-16769.
Leitinger, B and Kwan, AP (2006) The discoidin domain receptor DDR2 is a receptor for type X collagen. Matrix Biol 25(6):355-364.
Lemire, JM, Merrilees, MJ, Braun, KR, and Wight, TN (2002) Overexpression of the V3 variant of versican alters arterial smooth muscle cell adhesion, migration, and proliferation in vitro. J Cell Physiol 190(1):38-45.
Lessner, SM, Martinson, DE, and Galis, ZS (2004) Compensatory vascular remodeling during atherosclerotic lesion growth depends on matrix metalloproteinase-9 activity. Arterioscler Thromb Vasc Biol 24(11):2123-2129.
Li, D, Zhang, C, Song, F, Lubenec, I, Tian, Y, and Song, QH (2009) VEGF regulates FGF-2 and TGF-beta1 expression in injury endothelial cells and mediates smooth muscle cells proliferation and migration. Microvasc Res 77(2):134-142.
Li, DY, Brooke, B, Davis, EC, Mecham, RP, Sorensen, LK, Boak, BB, Eichwald, E, and Keating, MT (1998a) Elastin is an essential determinant of arterial morphogenesis. Nature 393(6682):276-280.
Li, DY, Faury, G, Taylor, DG, Davis, EC, Boyle, WA, Mecham, RP, Stenzel, P, Boak, B, and Keating, MT (1998b) Novel arterial pathology in mice and humans hemizygous for elastin. J Clin Invest 102(10):1783-1787.
185
Li, S, Chow, LH, and Pickering, JG (2000) Cell surface-bound collagenase-1 and focal substrate degradation stimulate the rear release of motile vascular smooth muscle cells. J Biol Chem 275(45):35384-35392.
Li, S, Lao, J, Chen, BP, Li, YS, Zhao, Y, Chu, J, Chen, KD, Tsou, TC, Peck, K, and Chien, S (2003) Genomic analysis of smooth muscle cells in 3-dimensional collagen matrix. FASEB J 17(1):97-99.
Liang, J, Liu, E, Yu, Y, Kitajima, S, Koike, T, Jin, Y, Morimoto, M, Hatakeyama, K, Asada, Y, Watanabe, T, Sasaguri, Y, Watanabe, S, and Fan, J (2006) Macrophage metalloelastase accelerates the progression of atherosclerosis in transgenic rabbits. Circulation 113(16):1993-2001.
Liaw, L, Almeida, M, Hart, CE, Schwartz, SM, and Giachelli, CM (1994) Osteopontin promotes vascular cell adhesion and spreading and is chemotactic for smooth muscle cells in vitro. Circ Res 74(2):214-224.
Liaw, L, Skinner, MP, Raines, EW, Ross, R, Cheresh, DA, Schwartz, SM, and Giachelli, CM (1995) The adhesive and migratory effects of osteopontin are mediated via distinct cell surface integrins. Role of alpha v beta 3 in smooth muscle cell migration to osteopontin in vitro. J Clin Invest 95(2):713-724.
Lijnen, HR, Van, HB, Vanlinthout, I, Verstreken, M, Rio, MC, and Collen, D (1999) Accelerated neointima formation after vascular injury in mice with stromelysin-3 (MMP-11) gene inactivation. Arterioscler Thromb Vasc Biol 19(12):2863-2870.
Lindner, V, Fingerle, J, and Reidy, MA (1993) Mouse model of arterial injury. Circ Res 73(5):792-796.
Lindner, V and Reidy, MA (1991) Proliferation of smooth muscle cells after vascular injury is inhibited by an antibody against basic fibroblast growth factor. Proc Natl Acad Sci U S A 88(9):3739-3743.
Liu, B, Itoh, H, Louie, O, Kubota, K, and Kent, KC (2004) The role of phospholipase C and phosphatidylinositol 3-kinase in vascular smooth muscle cell migration and proliferation. J Surg Res 120(2):256-265.
Liu, Y, Sinha, S, McDonald, OG, Shang, Y, Hoofnagle, MH, and Owens, GK (2005) Kruppel-like factor 4 abrogates myocardin-induced activation of smooth muscle gene expression. J Biol Chem 280(10):9719-9727.
Luttun, A, Lutgens, E, Manderveld, A, Maris, K, Collen, D, Carmeliet, P, and Moons, L (2004) Loss of matrix metalloproteinase-9 or matrix metalloproteinase-12 protects apolipoprotein E-deficient mice against atherosclerotic media destruction but differentially affects plaque growth. Circulation 109(11):1408-1414.
186
Macbeath, JR, Kielty, CM, and Shuttleworth, CA (1996) Type VIII collagen is a product of vascular smooth-muscle cells in development and disease. Biochem J 319 ( Pt 3):993-998.
Majesky, MW, Lindner, V, Twardzik, DR, Schwartz, SM, and Reidy, MA (1991) Production of transforming growth factor beta 1 during repair of arterial injury. J Clin Invest 88(3):904-910.
Matsuno, H, Stassen, JM, Vermylen, J, and Deckmyn, H (1994) Inhibition of integrin function by a cyclic RGD-containing peptide prevents neointima formation. Circulation 90(5):2203-2206.
Mayne, R (1986) Collagenous proteins of blood vessels. Arteriosclerosis 6(6):585-593.
Merrilees, MJ, Lemire, JM, Fischer, JW, Kinsella, MG, Braun, KR, Clowes, AW, and Wight, TN (2002) Retrovirally mediated overexpression of versican v3 by arterial smooth muscle cells induces tropoelastin synthesis and elastic fiber formation in vitro and in neointima after vascular injury. Circ Res 90(4):481-487.
Mitchison, TJ and Cramer, LP (1996) Actin-based cell motility and cell locomotion. Cell 84(3):371-379.
Mochizuki, S, Brassart, B, and Hinek, A (2002) Signaling pathways transduced through the elastin receptor facilitate proliferation of arterial smooth muscle cells. J Biol Chem 277(47):44854-44863.
Muragaki, Y, Jacenko, O, Apte, S, Mattei, MG, Ninomiya, Y, and Olsen, BR (1991a) The alpha 2(VIII) collagen gene. A novel member of the short chain collagen family located on the human chromosome 1. J Biol Chem 266(12):7721-7727.
Muragaki, Y, Mattei, MG, Yamaguchi, N, Olsen, BR, and Ninomiya, Y (1991b) The complete primary structure of the human alpha 1 (VIII) chain and assignment of its gene (COL8A1) to chromosome 3. Eur J Biochem 197(3):615-622.
Nagase, H, Visse, R, and Murphy, G (2006) Structure and function of matrix metalloproteinases and TIMPs. Cardiovasc Res 69(3):562-573.
Naito, M, Hayashi, T, Kuzuya, M, Funaki, C, Asai, K, and Kuzuya, F (1990) Effects of fibrinogen and fibrin on the migration of vascular smooth muscle cells in vitro. Atherosclerosis 83(1):9-14.
Newby, AC (2005) Dual role of matrix metalloproteinases (matrixins) in intimal thickening and atherosclerotic plaque rupture. Physiol Rev 85(1):1-31.
Nili, N, Zhang, M, Strauss, BH, and Bendeck, MP (2002) Biochemical analysis of collagen and elastin synthesis in the balloon injured rat carotid artery. Cardiovasc Pathol 11(5):272-276.
187
Nobes, CD and Hall, A (1995) Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell 81(1):53-62.
Nuttall, RK, Sampieri, CL, Pennington, CJ, Gill, SE, Schultz, GA, and Edwards, DR (2004) Expression analysis of the entire MMP and TIMP gene families during mouse tissue development. FEBS Lett 563(1-3):129-134.
O'Brien, ER, Garvin, MR, Stewart, DK, Hinohara, T, Simpson, JB, Schwartz, SM, and Giachelli, CM (1994) Osteopontin is synthesized by macrophage, smooth muscle, and endothelial cells in primary and restenotic human coronary atherosclerotic plaques. Arterioscler Thromb 14(10):1648-1656.
Okada, Y, Katsuda, S, Matsui, Y, and Nakanishi, I (1992) The modulation of collagen synthesis in cultured arterial smooth muscle cells by platelet-derived growth factor. Cell Biol Int Rep 16(10):1015-1022.
Okada, Y, Katsuda, S, Matsui, Y, Watanabe, H, and Nakanishi, I (1990) Collagen synthesis by cultured arterial smooth muscle cells during spontaneous phenotypic modulation. Acta Pathol Jpn 40(3):157-164.
Palecek, SP, Huttenlocher, A, Horwitz, AF, and Lauffenburger, DA (1998) Physical and biochemical regulation of integrin release during rear detachment of migrating cells. J Cell Sci 111 ( Pt 7):929-940.
Palecek, SP, Loftus, JC, Ginsberg, MH, Lauffenburger, DA, and Horwitz, AF (1997) Integrin-ligand binding properties govern cell migration speed through cell-substratum adhesiveness. Nature 385(6616):537-540.
Panda, D, Kundu, GC, Lee, BI, Peri, A, Fohl, D, Chackalaparampil, I, Mukherjee, BB, Li, XD, Mukherjee, DC, Seides, S, Rosenberg, J, Stark, K, and Mukherjee, AB (1997) Potential roles of osteopontin and alphaVbeta3 integrin in the development of coronary artery restenosis after angioplasty. Proc Natl Acad Sci U S A 94(17):9308-9313.
Pasterkamp, G, de Kleijn, DP, and Borst, C (2000) Arterial remodeling in atherosclerosis, restenosis and after alteration of blood flow: potential mechanisms and clinical implications. Cardiovasc Res 45(4):843-852.
Paulus, W, Sage, EH, Liszka, U, Iruela-Arispe, ML, and Jellinger, K (1991) Increased levels of type VIII collagen in human brain tumours compared to normal brain tissue and non-neoplastic cerebral disorders. Br J Cancer 63(3):367-371.
Pickering, JG, Chow, LH, Li, S, Rogers, KA, Rocnik, EF, Zhong, R, and Chan, BM (2000) alpha5beta1 integrin expression and luminal edge fibronectin matrix assembly by smooth muscle cells after arterial injury. Am J Pathol 156(2):453-465.
188
Pidkovka, NA, Cherepanova, OA, Yoshida, T, Alexander, MR, Deaton, RA, Thomas, JA, Leitinger, N, and Owens, GK (2007) Oxidized phospholipids induce phenotypic switching of vascular smooth muscle cells in vivo and in vitro. Circ Res 101(8):792-801.
Plenz, G, Dorszewski, A, Breithardt, G, and Robenek, H (1999a) Expression of type VIII collagen after cholesterol diet and injury in the rabbit model of atherosclerosis. Arterioscler Thromb Vasc Biol 19(5):1201-1209.
Plenz, G, Dorszewski, A, Volker, W, Ko, YS, Severs, NJ, Breithardt, G, and Robenek, H (1999b) Cholesterol-induced changes of type VIII collagen expression and distribution in carotid arteries of rabbit. Arterioscler Thromb Vasc Biol 19(10):2395-2404.
Plenz, G, Koenig, C, Reichenberg, S, and Robenek, H (1999c) Colony stimulating factors modulate the transcription of type VIII collagen in vascular smooth muscle cells. Atherosclerosis 144(1):25-32.
Plenz, G, Reichenberg, S, Koenig, C, Rauterberg, J, Deng, MC, Baba, HA, and Robenek, H (1999d) Granulocyte-macrophage colony-stimulating factor (GM-CSF) modulates the expression of type VIII collagen mRNA in vascular smooth muscle cells and both are codistributed during atherogenesis. Arterioscler Thromb Vasc Biol 19(7):1658-1668.
Plenz, GA, Deng, MC, Robenek, H, and Volker, W (2003) Vascular collagens: spotlight on the role of type VIII collagen in atherogenesis. Atherosclerosis 166(1):1-11.
Plump, AS, Smith, JD, Hayek, T, alto-Setala, K, Walsh, A, Verstuyft, JG, Rubin, EM, and Breslow, JL (1992) Severe hypercholesterolemia and atherosclerosis in apolipoprotein E-deficient mice created by homologous recombination in ES cells. Cell 71(2):343-353.
Pozzi, A, Wary, KK, Giancotti, FG, and Gardner, HA (1998) Integrin alpha1beta1 mediates a unique collagen-dependent proliferation pathway in vivo. J Cell Biol 142(2):587-594.
Prockop, DJ and Kivirikko, KI (1995) Collagens: molecular biology, diseases, and potentials for therapy. Annu Rev Biochem 64:403-434.
Raines, EW (2000) The extracellular matrix can regulate vascular cell migration, proliferation, and survival: relationships to vascular disease. Int J Exp Pathol 81(3):173-182.
Ravanti, L, Heino, J, Lopez-Otin, C, and Kahari, VM (1999) Induction of collagenase-3 (MMP-13) expression in human skin fibroblasts by three-dimensional collagen is mediated by p38 mitogen-activated protein kinase. J Biol Chem 274(4):2446-2455.
Reddick, RL, Zhang, SH, and Maeda, N (1994) Atherosclerosis in mice lacking apo E. Evaluation of lesional development and progression. Arterioscler Thromb 14(1):141-147.
189
Richardson, A and Parsons, T (1996) A mechanism for regulation of the adhesion-associated proteintyrosine kinase pp125FAK. Nature 380(6574):538-540.
Riessen, R, Wight, TN, Pastore, C, Henley, C, and Isner, JM (1996) Distribution of hyaluronan during extracellular matrix remodeling in human restenotic arteries and balloon-injured rat carotid arteries. Circulation 93(6):1141-1147.
Risinger, GM, Jr., Hunt, TS, Updike, DL, Bullen, EC, and Howard, EW (2006) Matrix metalloproteinase-2 expression by vascular smooth muscle cells is mediated by both stimulatory and inhibitory signals in response to growth factors. J Biol Chem 281(36):25915-25925.
Rocnik, EF, Chan, BM, and Pickering, JG (1998) Evidence for a role of collagen synthesis in arterial smooth muscle cell migration. J Clin Invest 101(9):1889-1898.
Roque, M, Fallon, JT, Badimon, JJ, Zhang, WX, Taubman, MB, and Reis, ED (2000) Mouse model of femoral artery denudation injury associated with the rapid accumulation of adhesion molecules on the luminal surface and recruitment of neutrophils. Arterioscler Thromb Vasc Biol 20(2):335-342.
Rosenbloom, J, Abrams, WR, and Mecham, R (1993) Extracellular matrix 4: the elastic fiber. FASEB J 7(13):1208-1218.
Saelman, EU, Nieuwenhuis, HK, Hese, KM, de Groot, PG, Heijnen, HF, Sage, EH, Williams, S, McKeown, L, Gralnick, HR, and Sixma, JJ (1994) Platelet adhesion to collagen types I through VIII under conditions of stasis and flow is mediated by GPIa/IIa (alpha 2 beta 1-integrin). Blood 83(5):1244-1250.
Sage, EH and Bornstein, P (1991) Extracellular proteins that modulate cell-matrix interactions. SPARC, tenascin, and thrombospondin. J Biol Chem 266(23):14831-14834.
Sage, H, Balian, G, Vogel, AM, and Bornstein, P (1984) Type VIII collagen. Synthesis by normal and malignant cells in culture. Lab Invest 50(2):219-231.
Sage, H and Iruela-Arispe, ML (1990) Type VIII collagen in murine development. Association with capillary formation in vitro. Ann N Y Acad Sci 580:17-31.
Sage, H, Pritzl, P, and Bornstein, P (1980) A unique, pepsin-sensitive collagen synthesized by aortic endothelial cells in culture. Biochemistry 19(25):5747-5755.
Saito, Y, Mori, S, Yokote, K, Kanzaki, T, Saito, Y, and Morisaki, N (1996) Phosphatidylinositol 3-kinase activity is required for the activation process of focal adhesion kinase by platelet-derived growth factor. Biochem Biophys Res Commun 224(1):23-26.
Sander, EE, ten Klooster, JP, van, DS, van der Kammen, RA, and Collard, JG (1999) Rac downregulates Rho activity: reciprocal balance between both GTPases determines cellular morphology and migratory behavior. J Cell Biol 147(5):1009-1022.
190
Sasamura, H, Shimizu-Hirota, R, and Saruta, T (2005) Extracellular Matrix Remodeling in Hypertension. Current Hypertension Reviews 1:51-60.
Sata, M, Maejima, Y, Adachi, F, Fukino, K, Saiura, A, Sugiura, S, Aoyagi, T, Imai, Y, Kurihara, H, Kimura, K, Omata, M, Makuuchi, M, Hirata, Y, and Nagai, R (2000) A mouse model of vascular injury that induces rapid onset of medial cell apoptosis followed by reproducible neointimal hyperplasia. J Mol Cell Cardiol 32(11):2097-2104.
Sawada, H, Konomi, H, and Hirosawa, K (1990) Characterization of the collagen in the hexagonal lattice of Descemet's membrane: its relation to type VIII collagen. J Cell Biol 110(1):219-227.
Shrivastava, A, Radziejewski, C, Campbell, E, Kovac, L, McGlynn, M, Ryan, TE, Davis, S, Goldfarb, MP, Glass, DJ, Lemke, G, and Yancopoulos, GD (1997) An orphan receptor tyrosine kinase family whose members serve as nonintegrin collagen receptors. Mol Cell 1(1):25-34.
Sibinga, NE, Foster, LC, Hsieh, CM, Perrella, MA, Lee, WS, Endege, WO, Sage, EH, Lee, ME, and Haber, E (1997) Collagen VIII is expressed by vascular smooth muscle cells in response to vascular injury. Circ Res 80(4):532-541.
Sieg, DJ, Hauck, CR, Ilic, D, Klingbeil, CK, Schaefer, E, Damsky, CH, and Schlaepfer, DD (2000) FAK integrates growth-factor and integrin signals to promote cell migration. Nat Cell Biol 2(5):249-256.
Sieg, DJ, Hauck, CR, and Schlaepfer, DD (1999) Required role of focal adhesion kinase (FAK) for integrin-stimulated cell migration. J Cell Sci 112 ( Pt 16):2677-2691.
Silence, J, Lupu, F, Collen, D, and Lijnen, HR (2001) Persistence of atherosclerotic plaque but reduced aneurysm formation in mice with stromelysin-1 (MMP-3) gene inactivation. Arterioscler Thromb Vasc Biol 21(9):1440-1445.
Siljander, PR, Hamaia, S, Peachey, AR, Slatter, DA, Smethurst, PA, Ouwehand, WH, Knight, CG, and Farndale, RW (2004) Integrin activation state determines selectivity for novel recognition sites in fibrillar collagens. J Biol Chem 279(46):47763-47772.
Sinha, S, Kielty, CM, Heagerty, AM, Canfield, AE, and Shuttleworth, CA (2001) Upregulation of collagen VIII following porcine coronary artery angioplasty is related to smooth muscle cell migration not angiogenesis. Int J Exp Pathol 82(5):295-302.
Skinner, MP, Raines, EW, and Ross, R (1994) Dynamic expression of alpha 1 beta 1 and alpha 2 beta 1 integrin receptors by human vascular smooth muscle cells. Alpha 2 beta 1 integrin is required for chemotaxis across type I collagen-coated membranes. Am J Pathol 145(5):1070-1081.
Slepian, MJ, Massia, SP, Dehdashti, B, Fritz, A, and Whitesell, L (1998) Beta3-integrins rather than beta1-integrins dominate integrin-matrix interactions involved in postinjury smooth muscle cell migration. Circulation 97(18):1818-1827.
191
Smith J, Ponting JM, Kumar S, and Rooney R (1996) Type VIII collagen promotes angiogenesis. J Vasc Res 33(51):94-94.
Somlyo, AP and Somlyo, AV (2000) Signal transduction by G-proteins, rho-kinase and protein phosphatase to smooth muscle and non-muscle myosin II. J Physiol 522 Pt 2:177-185.
Srivatsa, SS, Fitzpatrick, LA, Tsao, PW, Reilly, TM, Holmes, DR, Jr., Schwartz, RS, and Mousa, SA (1997) Selective alpha v beta 3 integrin blockade potently limits neointimal hyperplasia and lumen stenosis following deep coronary arterial stent injury: evidence for the functional importance of integrin alpha v beta 3 and osteopontin expression during neointima formation. Cardiovasc Res 36(3):408-428.
Stary, HC, Chandler, AB, Dinsmore, RE, Fuster, V, Glagov, S, Insull, W, Jr., Rosenfeld, ME, Schwartz, CJ, Wagner, WD, and Wissler, RW (1995) A definition of advanced types of atherosclerotic lesions and a histological classification of atherosclerosis. A report from the Committee on Vascular Lesions of the Council on Arteriosclerosis, American Heart Association. Circulation 92(5):1355-1374.
Stephan, S, Sherratt, MJ, Hodson, N, Shuttleworth, CA, and Kielty, CM (2004) Expression and supramolecular assembly of recombinant alpha1(viii) and alpha2(viii) collagen homotrimers. J Biol Chem 279(20):21469-21477.
Stone, GW, Moses, JW, Ellis, SG, Schofer, J, Dawkins, KD, Morice, MC, Colombo, A, Schampaert, E, Grube, E, Kirtane, AJ, Cutlip, DE, Fahy, M, Pocock, SJ, Mehran, R, and Leon, MB (2007) Safety and efficacy of sirolimus- and paclitaxel-eluting coronary stents. N Engl J Med 356(10):998-1008.
Strauss, BH, Robinson, R, Batchelor, WB, Chisholm, RJ, Ravi, G, Natarajan, MK, Logan, RA, Mehta, SR, Levy, DE, Ezrin, AM, and Keeley, FW (1996) In vivo collagen turnover following experimental balloon angioplasty injury and the role of matrix metalloproteinases. Circ Res 79(3):541-550.
Tanner, FC, Yang, ZY, Duckers, E, Gordon, D, Nabel, GJ, and Nabel, EG (1998) Expression of cyclin-dependent kinase inhibitors in vascular disease. Circ Res 82(3):396-403.
Thyberg, J, Blomgren, K, Roy, J, Tran, PK, and Hedin, U (1997) Phenotypic modulation of smooth muscle cells after arterial injury is associated with changes in the distribution of laminin and fibronectin. J Histochem Cytochem 45(6):837-846.
Tran, PK, Tran-Lundmark, K, Soininen, R, Tryggvason, K, Thyberg, J, and Hedin, U (2004) Increased intimal hyperplasia and smooth muscle cell proliferation in transgenic mice with heparan sulfate-deficient perlecan. Circ Res 94(4):550-558.
Tran-Lundmark, K, Tran, PK, Paulsson-Berne, G, Friden, V, Soininen, R, Tryggvason, K, Wight, TN, Kinsella, MG, Boren, J, and Hedin, U (2008) Heparan sulfate in perlecan
192
promotes mouse atherosclerosis: roles in lipid permeability, lipid retention, and smooth muscle cell proliferation. Circ Res 103(1):43-52.
Tulla, M, Pentikainen, OT, Viitasalo, T, Kapyla, J, Impola, U, Nykvist, P, Nissinen, L, Johnson, MS, and Heino, J (2001) Selective binding of collagen subtypes by integrin alpha 1I, alpha 2I, and alpha 10I domains. J Biol Chem 276(51):48206-48212.
Turner, NJ, Murphy, MO, Kielty, CM, Shuttleworth, CA, Black, RA, Humphries, MJ, Walker, MG, and Canfield, AE (2006) Alpha2(VIII) collagen substrata enhance endothelial cell retention under acute shear stress flow via an alpha2beta1 integrin-dependent mechanism: an in vitro and in vivo study. Circulation 114(8):820-829.
Urban, Z, Riazi, S, Seidl, TL, Katahira, J, Smoot, LB, Chitayat, D, Boyd, CD, and Hinek, A (2002) Connection between elastin haploinsufficiency and increased cell proliferation in patients with supravalvular aortic stenosis and Williams-Beuren syndrome. Am J Hum Genet 71(1):30-44.
Uzui, H, Lee, JD, Shimizu, H, Tsutani, H, and Ueda, T (2000) The role of protein-tyrosine phosphorylation and gelatinase production in the migration and proliferation of smooth muscle cells. Atherosclerosis 149(1):51-59.
Vaina, S, Ong, AT, and Serruys, PW (2005) New drug-eluting stents, optimizing technique, and the problem of drug-eluting stent restenosis. Minerva Cardioangiol 53(5):341-360.
Valenzuela, R, Shainoff, JR, DiBello, PM, Urbanic, DA, Anderson, JM, Matsueda, GR, and Kudryk, BJ (1992) Immunoelectrophoretic and immunohistochemical characterizations of fibrinogen derivatives in atherosclerotic aortic intimas and vascular prosthesis pseudo-intimas. Am J Pathol 141(4):861-880.
van der, RM and Garrone, R (1991) Collagen family of proteins. FASEB J 5(13):2814-2823.
van Hennik, PB, ten Klooster, JP, Halstead, JR, Voermans, C, Anthony, EC, Divecha, N, and Hordijk, PL (2003) The C-terminal domain of Rac1 contains two motifs that control targeting and signaling specificity. J Biol Chem 278(40):39166-39175.
van Hinsbergh, V and Koolwijk, P (2008) Endothelial sprouting and angiogenesis: matrix metalloproteinases in the lead. Cardiovasc Res 78(2):203-212.
Vogel, W (1999) Discoidin domain receptors: structural relations and functional implications. FASEB J 13 Suppl:S77-S82.
Vogel, W, Brakebusch, C, Fassler, R, Alves, F, Ruggiero, F, and Pawson, T (2000) Discoidin domain receptor 1 is activated independently of beta(1) integrin. J Biol Chem 275(8):5779-5784.
193
Vogel, W, Gish, GD, Alves, F, and Pawson, T (1997) The discoidin domain receptor tyrosine kinases are activated by collagen. Mol Cell 1(1):13-23.
Vogel, WF, Aszodi, A, Alves, F, and Pawson, T (2001) Discoidin domain receptor 1 tyrosine kinase has an essential role in mammary gland development. Mol Cell Biol 21(8):2906-2917.
Walker-Caprioglio, HM, Hunter, DD, McGuire, PG, Little, SA, and McGuffee, LJ (1995) Composition in situ and in vitro of vascular smooth muscle laminin in the rat. Cell Tissue Res 281(1):187-196.
Wang, CZ, Su, HW, Hsu, YC, Shen, MR, and Tang, MJ (2006a) A discoidin domain receptor 1/SHP-2 signaling complex inhibits alpha2beta1-integrin-mediated signal transducers and activators of transcription 1/3 activation and cell migration. Mol Biol Cell 17(6):2839-2852.
Wang, X, Chai, H, Lin, PH, Lumsden, AB, Yao, Q, and Chen, C (2006b) Mouse models of neointimal hyperplasia: techniques and applications. Med Sci Monit 12(9):RA177-RA185.
Wang, Y and Kovanen, PT (1999) Heparin proteoglycans released from rat serosal mast cells inhibit proliferation of rat aortic smooth muscle cells in culture. Circ Res 84(1):74-83.
Ward, MR, Pasterkamp, G, Yeung, AC, and Borst, C (2000) Arterial remodeling. Mechanisms and clinical implications. Circulation 102(10):1186-1191.
Webb, DJ, Parsons, JT, and Horwitz, AF (2002) Adhesion assembly, disassembly and turnover in migrating cells -- over and over and over again. Nat Cell Biol 4(4):E97-100.
Weitkamp, B, Cullen, P, Plenz, G, Robenek, H, and Rauterberg, J (1999) Human macrophages synthesize type VIII collagen in vitro and in the atherosclerotic plaque. FASEB J 13(11):1445-1457.
Westermarck, J and Kahari, VM (1999) Regulation of matrix metalloproteinase expression in tumor invasion. FASEB J 13(8):781-792.
Wight, TN, Raugi, GJ, Mumby, SM, and Bornstein, P (1985) Light microscopic immunolocation of thrombospondin in human tissues. J Histochem Cytochem 33(4):295-302.
Wiskirchen, J, Schober, W, Schart, N, Kehlbach, R, Wersebe, A, Tepe, G, Claussen, CD, and Duda, SH (2004) The effects of paclitaxel on the three phases of restenosis: smooth muscle cell proliferation, migration, and matrix formation: an in vitro study. Invest Radiol 39(9):565-571.
194
Xiong, JP, Li, R, Essafi, M, Stehle, T, and Arnaout, MA (2000) An isoleucine-based allosteric switch controls affinity and shape shifting in integrin CD11b A-domain. J Biol Chem 275(49):38762-38767.
Yamaguchi, N, Benya, PD, van der, RM, and Ninomiya, Y (1989) The cloning and sequencing of alpha 1(VIII) collagen cDNAs demonstrate that type VIII collagen is a short chain collagen and contains triple-helical and carboxyl-terminal non-triple-helical domains similar to those of type X collagen. J Biol Chem 264(27):16022-16029.
Yamaguchi, N, Mayne, R, and Ninomiya, Y (1991) The alpha 1 (VIII) collagen gene is homologous to the alpha 1 (X) collagen gene and contains a large exon encoding the entire triple helical and carboxyl-terminal non-triple helical domains of the alpha 1 (VIII) polypeptide. J Biol Chem 266(7):4508-4513.
Yamamoto, M, Yamato, M, Aoyagi, M, and Yamamoto, K (1995) Identification of integrins involved in cell adhesion to native and denatured type I collagens and the phenotypic transition of rabbit arterial smooth muscle cells. Exp Cell Res 219(1):249-256.
Yao, CC, Breuss, J, Pytela, R, and Kramer, RH (1997) Functional expression of the alpha 7 integrin receptor in differentiated smooth muscle cells. J Cell Sci 110 ( Pt 13):1477-1487.
Yasuda, O, Fukuo, K, Maeda, N, and Ogihara, T (2001) Elevated expression of type VIII collagen gene in the atherosclerotic plaque of the ApoE-deficient mouse. Ann N Y Acad Sci 947:312-315.
Yasuda, O, Zhang, SH, Miyamoto, Y, and Maeda, N (2000) Differential expression of the alpha1 type VIII collagen gene by smooth muscle cells from atherosclerotic plaques of apolipoprotein-E-deficient mice. J Vasc Res 37(3):158-169.
Yue, TL, McKenna, PJ, Ohlstein, EH, Farach-Carson, MC, Butler, WT, Johanson, K, McDevitt, P, Feuerstein, GZ, and Stadel, JM (1994) Osteopontin-stimulated vascular smooth muscle cell migration is mediated by beta 3 integrin. Exp Cell Res 214(2):459-464.
Zempo, N, Kenagy, RD, Au, YP, Bendeck, M, Clowes, MM, Reidy, MA, and Clowes, AW (1994) Matrix metalloproteinases of vascular wall cells are increased in balloon-injured rat carotid artery. J Vasc Surg 20(2):209-217.
Zempo, N, Koyama, N, Kenagy, RD, Lea, HJ, and Clowes, AW (1996) Regulation of vascular smooth muscle cell migration and proliferation in vitro and in injured rat arteries by a synthetic matrix metalloproteinase inhibitor. Arterioscler Thromb Vasc Biol 16(1):28-33.
Zhang, J, Nie, L, Razavian, M, Ahmed, M, Dobrucki, LW, Asadi, A, Edwards, DS, Azure, M, Sinusas, AJ, and Sadeghi, MM (2008) Molecular imaging of activated matrix metalloproteinases in vascular remodeling. Circulation 118(19):1953-1960.
195
Zhang, SH, Reddick, RL, Piedrahita, JA, and Maeda, N (1992) Spontaneous hypercholesterolemia and arterial lesions in mice lacking apolipoprotein E. Science 258(5081):468-471.
Zheng, DQ, Woodard, AS, Fornaro, M, Tallini, G, and Languino, LR (1999) Prostatic carcinoma cell migration via alpha(v)beta3 integrin is modulated by a focal adhesion kinase pathway. Cancer Res 59(7):1655-1664.
Zhu, B, Kuhel, DG, Witte, DP, and Hui, DY (2000) Apolipoprotein E inhibits neointimal hyperplasia after arterial injury in mice. Am J Pathol 157(6):1839-1848.
Zou, Y, Dietrich, H, Hu, Y, Metzler, B, Wick, G, and Xu, Q (1998) Mouse model of venous bypass graft arteriosclerosis. Am J Pathol 153(4):1301-1310.
![RESEARCH ARTICLE Manipulation of Developing Juvenile Structures in Purple Sea … · 2015-04-09 · pentameral symmetry, a water vascular system, mutable collagen [2,3] and larval](https://img.pdfslide.us/doc/110x75/5f7f44a36eecb57bd7243cef/research-article-manipulation-of-developing-juvenile-structures-in-purple-sea-2015-04-09.jpg)
