ENGINEERING POLY(ETHYLENE GLYCOL) HYDROGEL SCAFFOLDS TO
MODULATE SMOOTH MUSCLE CELL PHENOTYPE
by
JEFFREY ALAN BEAMISH
Submitted in partial fulfillment of the requirements
For the degree of Doctor of Philosophy
Dissertation Advisors: Dr. Roger E. Marchant and Dr. Kandice Kottke-Marchant
Department of Biomedical Engineering
CASE WESTERN RESERVE UNIVERSITY
August, 2009 CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of
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candidate for the ______degree *.
(signed)______(chair of the committee)
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(date) ______
*We also certify that written approval has been obtained for any proprietary material contained therein.
To my loving wife, Susan, and my parents, who have provided unwavering support for all my endeavors Table of Contents
Table of Contents ...... i List of Tables ...... iii List of Figures...... iv Acknowledgements ...... vi List of Abbreviations ...... ix Abstract...... xiv
CHAPTER 1: Cardiovascular Disease, Bypass Grafting, and Intimal Hyperplasia. 1 1.1. Summary of Cardiovascular Disease in the United States...... 1 1.2. Atherosclerosis...... 1 1.3. Treatment Approaches for Cardiovascular Disease...... 2 1.4. Anastomotic Intimal Hyperplasia...... 5 1.5. References ...... 15
CHAPTER 2: Regulation of SMC Phenotype: Implications for Vascular Tissue Engineering...... 20 2.1. Introduction ...... 20 2.2. The Continuum of SMC Phenotypes ...... 21 2.3. Markers of Contractile SMC Phenotype ...... 24 2.4. Mediators of SMC Phenotype...... 30 2.5. Molecular Regulation of SMC Gene Expression...... 48 2.6. Engineered Biomaterial Approaches to Regulate SMC Phenotype...... 54 2.7. Conclusions ...... 58 2.8. References ...... 61
CHAPTER 3: Vascular Tissue Engineering...... 83 3.1. Introduction ...... 83 3.2. Normal Blood Vessel Histology ...... 83 3.3. Design Criteria ...... 84 3.4. Tissue Engineered Modification of Existing Graft Materials ...... 84 3.5. Fully Tissue Engineered Blood Vessels (TEBVs) ...... 87 3.6. Discussion ...... 99 3.7. PEG-based Hydrogels for Tissue Engineering...... 103 3.8. Specific Aims ...... 108 3.9. References ...... 111
CHAPTER 4: The Effects of Monoacrylated Poly(Ethylene Glycol) on the Properties of Poly(Ethylene Glycol) Diacrylate Hydrogels Used for Tissue Engineering...... 119 4.1. Introduction ...... 119 4.2. Materials and Methods...... 123 4.3. Results ...... 129 4.4. Discussion ...... 136 4.5. Conclusions ...... 142
i 4.6. Acknowledgements ...... 142 4.7. References ...... 143
CHAPTER 5: The Influence of RGD-Bearing Hydrogels on the Re-expression of Contractile Vascular Smooth Muscle Cell Phenotype...... 146 5.1. Introduction ...... 146 5.2. Materials and Methods...... 148 5.3. Results ...... 156 5.4. Discussion ...... 167 5.5. Conclusions ...... 172 5.6. Acknowledgements ...... 172 5.7. References ...... 174
CHAPTER 6: The Effects of Heparin Releasing Hydrogels on Vascular Smooth Muscle Cell Phenotype ...... 178 6.1. Introduction ...... 178 6.2. Materials and Methods...... 180 6.3. Results ...... 189 6.4. Discussion ...... 201 6.5. Conclusions ...... 207 6.6. Acknowledgments...... 207 6.7. References ...... 208
CHAPTER 7: Conclusions and Future Directions ...... 213 7.1. Summary and Conclusions of Completed Work...... 213 7.2. Engineering of Improved Scaffold Systems...... 215 7.3. Modulation of Cultured SMCs Toward a Contractile Phenotype...... 226 7.4. References ...... 233
Bibliography ...... 238
ii List of Tables
Table 3.1-Design criteria for tissue engineered blood vessels 86
Table 4.1-Chemical structures of PEGDA and PEGMA 124
Table 5.1-Primers used for real-time RT-PCR 153
Table 6.1-Summary of heparin release from PEGDA hydrogels 193
Table 6.2-Network properties of PEGDA hydrogels 196
Table 6.3-Heparin concentration in medium during transwell insert study 199
iii List of Figures
Figure 1.1-Typical distribution of intimal hyperplasia 7
Figure 2.1-Summary of characteristics of SMC phenotypes 23
Figure 2.2-Mechanisms of SMC phenotype modulation 32
Figure 2.3-Chemical composition and structure of heparin 34
Figure 2.4-Molecular regulation of smooth muscle α-actin transcription 53
Figure 3.1-Structure of a muscular artery 85
Figure 3.2-Scaffolds used for tissue engineered blood vessels 101
Figure 3.3-Overview of PEGDA polymerization 103
Figure 3.4-Schematic overview of project specific aims 110
Figure 4.1-Schematic representation of PEGDA networks 122
Figure 4.2-Properties PEGDA-co-PEGMA hydrogels (mole fraction) 131
Figure 4.3-Mass swelling ratios of PEGDA-co-PEGMA hydrogels 132
Figure 4.4-Mass swelling ratios of hydrogels with low PEGDA concentration 132
Figure 4.5-Shear moduli of PEGDA-co-PEGMA hydrogels 134
* Figure 4.6-The effect of junction functionality on ve 134
* Figure 4.7-ve of PEGDA-co-PEGMA hydrogels calculated from swelling data 135
* Figure 4.8-ve of PEGDA-co-PEGMA hydrogels calculated from mechanical data 135
Figure 4.9-The effect of PEGMA concentration on chain length 137
Figure 5.1-Contractile marker mRNA expression after seeding 158
Figure 5.2-Contractile marker mRNA expression during differentiation 160
Figure 5.3-Western blot of contractile marker proteins during differentiation 162
Figure 5.4-Immunofluorescent stains of contractile marker proteins 163
iv Figure 5.5-Intracellular organization of contractile marker proteins 164
Figure 5.6-Attachement to peptide modified hydrogels 166
Figure 6.1-Schematic diagram of heparin release experiments 188
Figure 6.2-Effects of heparin on SMC proliferation 190
Figure 6.3-Effects of heparin on SMC contractile marker expression 192
Figure 6.4-Heparin release profiles for PEGDA hydrogels 194
Figure 6.5-Effect of released heparin on contractile marker expression 198
Figure 6.6-Effect of released heparin on myocardin expression 200
Figure 6.7-Contractile marker expression on heparin releasing PEGDA scaffolds 202
Figure 7.1-Schematic overview of future directions for this project 216
Figure 7.2-Effect of TGF-β1 on smooth muscle α-actin expression 220
Figure 7.3-Structure and activity of degradable PEGDA derivatives 222
Figure 7.4-Calculated compliance of vascular graft materials 225
Figure 7.5-Coating of the adventitial surface of an ePTFE graft material 227
Figure 7.6-Time dependent focal adhesion (FA) development 230
v Acknowledgements
This work would not have been possible without the contributions of a great many people. The path that led me toward science began long before I arrived in Cleveland.
I’d like to thank my math and science teachers at Troy High School who, despite limited resources, inspired my interest in science. I am also thankful for my experiences as an undergraduate at Northwestern University, where I found a fertile ground to nurture my scientific curiosity. During my undergraduate years, I am also indebted to the Dartmouth molecular materials REU program who provided my first research experience and to Dr.
E. T. Papoutsakis who offered me a position in his lab as an undergraduate. In particular,
I am grateful for the mentoring of Dr. Christopher Tomas, my graduate student advisor in the Papoutsakis lab. I have tried to emulate his excellent mentorship with the undergraduates that have worked with me over the course of my PhD training. Without the foundation these experiences provided, my PhD work would not have been possible.
The MSTP program at CWRU has provided a superb training environment for me, and I sincerely appreciate the oil that Donna McIlwain, Deidre Gruning, Kathy
Schultz and Bart Jarmusch have applied to keep the gears of this excellent program turning. I also appreciate the assistance of Angie Bracanovic guiding my orders, reimbursements, and numerous other forms through the BME bureaucracy. Holly Jones,
Pat Matkovic, and Alison Holzfaster have been invaluable liaisons in Dr. Kottke-
Marchant’s office.
The Marchant lab has proved to be an ideal training environment for me. At its helm, Dr. Roger Marchant and Dr. Kandice Kottke-Marchant have been excellent mentors and role models. I appreciate very much the latitude they have granted me to
vi develop this project and their patience while I “spun my wheels” for the first couple
years. It was through these early failures that I learned the most about how to design and
conduct experiments. It is only in hindsight that I see how their subtle guidance helped to
nudge me finally toward a viable project and successful experiments, while still
providing me with an opportunity to learn how to do my own research. For this
experience, I will always be grateful.
The members of the Marchant group, past and present, have been a daily source of support and encouragement. Some have shared my entire journey. I appreciate Dr.
Faina Kligman’s help with chemical synthesis, Dr. Junmin Zhu’s penchant for lively
scientific discussions, and Chris Hofmann’s help with using and maintaining a wide
range of equipment in our lab. The old crew of Dr. Eric Anderson, Dr. Coby Larsen, and
Dr. Arya Kumar helped me get my feet wet, and put up with my daily inquiries about the
procedures, equipment, and supplies in the lab. In particular, Coby and Arya have been
role models and advisors as they navigated the MSTP just two years ahead of me.
Although we have worked together for less than two years, Lynn Dudash has proven to be a valuable scientific sounding board and an even better friend. I also have been lucky
to have the assistance of an excellent team of hard-working undergraduates: Alex Fu,
Leah Geyer, Nada Haq, and Ae-jin “Jeannie” Choi. Much of my work, literally, would
not have been possible without their help.
I also appreciate the support that has been given to me from outside my lab. Dr.
Eben Alsberg and Dr. George Dubyak have acted as much more than faces at the table
during my thesis committee meetings. Outside of this formal setting, they have
generously volunteered their advice, laboratories, and students to help me with my
vii project. I also appreciate the assistance of Domenick “Tony” Prosdocimo, in Dr.
Dubyak’s lab, and Meghan Pennini, a former student Dr. Clifford Harding’s lab, for
teaching me western blotting and real time RT-PCR, respectively. I also am grateful for
the numerous scientific discussions with my colleagues Ken Rys, Paul Lin, and Melissa
Krebs.
I would also like to acknowledge the funding of the National Institutes of Health
for supporting my training indirectly through the Case MSTP and Dr. Marchant’s research program. I also am thankful for the predoctoral fellowship provided to me by the American Heart Association over the last two years.
Most importantly of all, I would like to thank my family for their unwavering support in all of my endeavors. My parents, Mike and Ginny Beamish, have always unselfishly provided to me anything within their means that could help me along my
path. My lovely wife Susan has been my most important daily source of support and motivation through this process. She has always been ready to give me a push when my
momentum faltered or provide a helping hand when my goals exceeded what I could
accomplish on my own. Having played such an important role along the way, I hope that
my family can now equally share in the happiness and sense of accomplishment afforded
by the completion of this work.
viii List of Abbreviations
2D Two dimensional 3D Three dimensional ε Mesh size λ Deformation ratio, h/h0 ρp Bulk density of polymer ρs Density of solvent σ Stress μg Microgram μl Microliter μm Micrometer μM Micromolar χ Flory-Huggins interaction parameter °C Degrees Celsius Å Angstrom ACLP Aortic carboxypeptidase-like protein ACRL Acrylate ANOVA Analysis of variance ATP Adenosine triphosphate BCA Bicinchoninic acid bFGF Basic fibroblast growth factor BMC Bone marrow derived cell BSA Bovine serum albumin CABG Coronary artery bypass grafting CArG CC(A/T)6GG promoter sequence, SRF binding site CD Cluster of differentiation cDNA Complementary deoxyribonucleic acid cELISA Cell based enzyme-linked immunosorbent assay CHD Coronary heart disease cm Centimeter Cn Characteristic ratio CS-PEGDA Collagenase sensitive PEGDA Ct Threshold cycle CVD Cardiovascular disease d Day(s) D Diameter DAIH Distal anastomotic intimal hyperplasia DAPI 4’,6-diamidino-2-phenylindole DCM Dichloromethane DMEM Dulbecco’s Modified Eagle Medium DMMB Dimethylmethylene blue dn/dc Derivative of refractive index with respect to concentration DNA Deoxyribonucleic acid EC Endothelial cell
ix ECM Extracellular matrix EGF Epidermal growth factor ELISA Enzyme-linked immunosorbent assay EPC Endothelial progenitor cell ePTFE Expanded poly(tetrafluoroethylene) ER Endoplasmic reticulum ERK Extracellular signal regulated kinase f Mass fraction of polymer F Junction functionality FA Focal adhesion FAK Focal adhesion kinase FBS Fetal bovine serum fmoc Fluorenylmethyloxycarbonyl FN Fibronectin fPEGDA Mass fraction of PEGDA at the time of polymerization FRNK Focal adhesion kinase related nonkinase G Equilibrium shear modulus g Gram GAG Glycosaminoglycan GAPDH Glyceraldehyde-3-phosphate dehydrogenase GlcA D-glucuronic acid GlcN D-glucosamine GMP Guanosine monophosphate GPC Gel permeation chromatography GPCR G-protein coupled receptor h Hour H, h Height h0 Predeformation height HCASMC Human coronary artery smooth muscle cell HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HERP Hairy-and enhancer of split-like-related repressor protein HPLC High performance liquid chromatography HSPG Heparan sulfate proteoglycan Hz Hertz ICAM Intercellular adhesion molecule-1 ID Inner diameter IdoA L-iduronic acid IEL Internal elastic lamina IF Immunofluorescence IGF Insulin-like growth factor IH Intimal hyperplasia IL-8 Interleukin-8 IND Internodal distance ITS Insulin-transferrin-selenium JNK c-Jun N-terminal kinase K Kelvin
x kD Kilodalton KLF-4 Kruppel-like factor 4 kPa Kilopascals l Weighted average bond length l Liter LDL Low density lipoprotein LN Laminin LPA Lysophosphatidic acid LSM Low serum medium LSM+H Low serum medium plus heparin (DMEM, 2% FBS, 400 μg/ml heparin) M Molar, moles per liter MALDI-MS Matrix assisted laser desorption/ionization mass spectroscopy MAPK Mitogen activated protein kinase Mc Molecular weight between cross-links MCP-1 Monocyte chemotactic protein-1 mg Milligram MHC-II Major histocompatibility complex, class II MI Myocardial infarction ml Milliliter mm Millimeter mM Millimolar mmHg Millimeters of mercury MMP Matrix metalloproteinase mN Millinewton Mn Number averaged molecular weight mp Mass of polymer MPa Megapascals Mr Repeat unit molecular weight mRNA Messenger RNA MRTF Myocardin related transcription factor ms Mass of swollen hydrogel MSC Mesenchymal stem cell MTS 3-(4,5-dimethylthiazol-2-yl)-5-(3-cardoxymethoxyphenyl)-2-(4- sulfophenyl)-2H-tetraxolium, inner salt mW Milliwatt MW Molecular weight Mw Weight averaged molecular weight MWCO Molecular weight cutoff MWPEGDA PEGDA molecular weight N Number of replicates NHS N-hydroxysuccinimide NIH National Institutes of Health nm Nanometer NMHC Non-muscle myosin heavy chain NMR Nuclear magnetic resonance NO Nitric oxide
xi OD Outer diameter Pa Pascal PAGE Polyacrylamide gel electrophoresis PBS Phosphate buffered saline PCI Percutaneous coronary intervention PCL Poly(ε-caprolactone) PCR Polymerase chain reaction PDGF Platelet derived growth factor PEG Poly(ethylene glycol) PEGDA Poly(ethylene glycol) diacrylate PEGMA Poly(ethylene glycol) monoacrylate PET Poly(ethylene terephthalate) PFA Paraformaldehyde PGA Poly(glycolic acid) PHA Poly(hydroxyalkanoate) PIAS Protein inhibitor of activated STAT PKB Protein kinase B (Akt) PKC Protein kinase C PKR Double stranded RNA protein kinase PLGA Poly(lactic-co-glycolic acid) PLLA Poly(L-lactic acid) q Mass swelling ratio qPCR Quantitative PCR R Universal gas constant, 8.413J K-1mol-1 r2 Pearson product moment correlation coefficient RF Radio frequency RGD Arginine-glycine-aspartic acid peptide RGD-gel RGD containing PEGDA hydrogel RIPA Radioimmunoprecipitation assay RNA Ribonucleic acid RPM Revolutions per minute RTK Receptor tyrosine kinase RT-PCR Reverse transcription polymerase chain reaction SDS Sodium dodecyl sulfate SFAM Serum-free attachment medium SFM Serum-free medium siRNA Small interfering RNA SMαA Smooth muscle α-actin SMC Smooth muscle cell SmGM Smooth muscle growth medium (Lonza) SM-MHC Smooth muscle myosin heavy chain SM-MLCK Smooth muscle myosin light chain kinase SRF Serum response factor STAT Signal transducer and activator of transcription T Temperature TBS Tris-buffered saline
xii TCPS Tissue culture polystyrene TEBV Tissue engineered blood vessel TGF-β Transforming growth factor-β TGFβR TGF-β receptor UV Ultraviolet
ve Number of effective cross-links per gram polymer * ve Number of effective cross-links per PEGDA molecule
vp Volume fraction of polymer
vp,r Volume fraction of polymer, relaxed state
vp,s Volume fraction of polymer, swollen state
Vs Molar volume of the solvent v/v Volume per volume VCAM-1 Vascular cell adhesion molecule-1 w/v Weight per volume w/w Weight per weight wk Week(s) wt Weight
Peptide sequences are listed from amino- to carboxy-terminus using standard single letter abbreviations for each amino acid.
Nucleic acid sequences are listed from 5’ to 3’ using standard single letter abbreviations for each nucleotide.
xiii Engineering Poly(Ethylene Glycol) Hydrogel Scaffolds to Modulate Smooth Muscle Cell Phenotype
Abstract
by
JEFFREY ALAN BEAMISH
This work investigated the hypothesis that smooth muscle cell (SMC) phenotype
can be modulated by a cell-instructive hydrogel scaffold. By modulating SMCs toward a quiescent contractile phenotype, such a scaffold may facilitate the regeneration of functional vascular tissue as part of a tissue engineered blood vessel (TEBV) and may mitigate SMC intimal hyperplasia, which is one of the common modes of vascular prosthesis failure. A photopolymerizable poly(ethylene glycol) diacrylate (PEGDA) hydrogel system was employed because this scaffold material can be engineered quantitatively to meet a broad range of physical and biological design specifications.
PEGDA hydrogel networks were copolymerized with poly(ethylene glycol) monoacrylate
(PEGMA), which could tether pendent cell-adhesive peptides to the network. The physical characteristics of a range of copolymer network compositions (5-20% w/w
PEGDA, 0-20% PEGMA) were determined and analyzed for the scaffold design. Mass swelling ratio (7.5±0.1 to 27.1±0.7) and shear modulus (4.0±0.6 to 104±4 kPa) data indicated that hydrogel properties were controlled quantitatively and independently of
PEGMA concentration. SMCs attached to PEGDA-co-GRGDSP-PEGMA (RGD-gels)
xiv in a ligand specific manner. RGD-gels also supported modulation toward a contractile
SMC phenotype that was indistinguishable from fibronectin control substrates. In these experiments, low serum medium with heparin induced rapid up-regulation of contractile phenotype marker mRNA (2.7- to 25-fold) and proteins, as well as intracellular organization of these markers. Based on these results, RGD-bearing PEGDA hydrogel scaffolds were engineered to modulate SMC phenotype by providing controlled release
of heparin. Using PEGDA molecular weight (1-6 kD) and concentration (10-30% w/w), a broad range of release profiles was achieved with durations from hours to weeks.
These cell-instructive scaffolds stimulated up-regulation of contractile phenotype markers
(~1.5-fold) that was driven by released heparin. These results suggest that the delivery of soluble signaling factors, such as heparin, from an RGD-gel scaffold is an effective approach to induce rapid changes in SMC phenotype. A cell-instructive scaffold system, such as described here, will facilitate the engineering of functional smooth muscle tissue for a variety of applications and may improve the long-term patency of TEBVs.
xv CHAPTER 1: Cardiovascular Disease, Bypass Grafting, and Intimal Hyperplasia
1.1. Summary of Cardiovascular Disease in the United States
Cardiovascular disease (CVD) is the leading cause of mortality in the United
States [1]. Cardiovascular disease primarily manifests itself as coronary heart disease
(CHD), stroke, heart failure, and/or hypertension, but also includes other diagnoses.
Among deaths due to CVD, 52% are caused by CHD, which includes myocardial infarctions (MI) [1]. This represents about 1 of every 5 deaths in the United States [1].
In addition to being the leading cause of mortality, CHD also is a substantial burden on the health care system. CHD accounted for nearly 1.8 million hospital discharges in
2006, and the projected cost of CHD in the United States for 2009 is $165 billion [1].
Clearly, management and treatment of cardiovascular disease is a critical health issue facing the United States.
1.2. Atherosclerosis
The underlying pathology leading to CHD, as well as peripheral artery disease, is atherosclerosis. The etiology and pathophysiology of atherosclerosis have been well reviewed elsewhere and will be summarized only briefly here [2, 3]. Atherosclerosis is a progressive, chronic disease. Lesions begin as low density lipoprotein (LDL) cholesterol and other lipids accumulate in the intima below areas of dysfunctional endothelium.
Monocytes are recruited to the area, where they accumulate lipid (resulting in macrophage “foam” cells). Factors released from these macrophages and from platelets adhered to the dysfunctional endothelium stimulate the accumulation of smooth muscle cells (SMCs) and additional inflammatory cells. SMCs proliferate and secrete extracellular matrix (ECM), forming a collagen I-rich fibrous cap above the cholesterol
1 filled core, resulting in a mature lesion. This process can weaken the surrounding vessel
wall, leading to aneurysm formation. If the fibrous cap is disrupted, catastrophic
thrombosis and/or the production of thromboemboli can result leading to vessel
occlusion. As the lesion grows, sometimes via the incorporation of non-occluding
thrombi, the lesion itself can result in critical stenosis of the vessel. Vessel occlusion
results in downstream ischemia and infarction.
1.3. Treatment Approaches for Cardiovascular Disease
1.3.1. Prevention
Preventative medical management of atherosclerotic disease is the least invasive
and most cost effective way to treat CHD. Detailed recommendations for preventative
care can be found elsewhere, but are summarized briefly below [4, 5]. For all
individuals, healthy lifestyle choices play an important role in reducing risk for CHD including low sodium intake, a balanced diet, minimal alcohol consumption, daily exercise, and cessation of smoking. Risk factors that are monitored in the clinic and
treated medically (in addition to lifestyle modifications) include the management of blood pressure, blood lipids, arrhythmias (such as atrial fibrillation), and diabetes. Low dose aspirin is also recommended for patients with elevated risk. For patients with known CHD (e.g. post-MI, positive catheterization, etc), more aggressive management of the above risks is recommended. Treatments for sequellae of previous disease also are included in a management plan, such as anti-thrombotic therapy after stent placement, beta-blockers post-MI, and renin-angiotensin-aldosterone axis management in patients with subsequent heart failure.
2 1.3.2. Angioplasty and Stent Placement
When medical management of CHD fails, more invasive procedures are often
required to restore adequate blood flow. The most common intervention is dilation of the
stenotic artery with a balloon catheter and placement of a stent. Such percutaneous coronary interventions (PCI) are less invasive than surgery and can be performed with
minimal additional effort during diagnostic catheterization procedures. In 2006,
approximately 1.3 million PCI were performed [1]. The average cost for each procedure
is approximately $48,000 [1]. The mortality for this procedure is 0.71% [1]. Greater than 90% of PCI now involve placement of a stent, and of these, greater than 70% were drug-eluting stents [1].
1.3.3. Bypass Grafting
Coronary artery bypass grafting (CABG) is one of the mainstays of treatment for
advanced cardiovascular disease. Compared with PCI employing drug eluting stents for
treatment of multi-vessel disease, the overall risk-adjusted outcomes over 18 months are
better for CABG (risk-adjusted hazard ratio for death: 0.80) [6]. However, CABG is a more extensive procedure and incurs greater risks and costs. The mortality associated with CABG is 1.94% [1]. The average cost of each procedure is just under $100,000 [1].
In 2006, approximately 448,000 of CABG procedures were performed on 253,000 patients, representing a slight decline over previous years [1]. The preferred bypass
conduits for these procedures are autologous vessels including the internal mammary
artery and the saphenous vein. Among these options, better outcomes have been
associated with the internal mammary artery [7, 8]. The patency of autologous vessels is
3 adequate to substantially improve and preserve life. However, the 10-year patency rates
for vein grafts is near 50%, which is sub-optimal [8, 9].
CABG is not an option for all patients, as 20-30% of patients do not have suitable autologous vessels [10, 11]. Unfortunately for these patients, the best available synthetic alternatives for small diameter (< 5 mm) reconstructions, expanded poly(tetrafluoroethylene) (ePTFE) grafts, have yielded disappointing results when used for CABG. In various clinical studies of ePTFE grafts with a 4 mm inner diameter used
for CABG, patency was only 59% at 12 months [12], 33% after 29 months [13], and 9%
after 45 months [14] (compared with roughly 50% for vein grafts after 10 years [9]).
Primary patency of small diameter ePTFE grafts used in peripheral reconstructions is also
poor varying from 31% at 24 months [15] to 41% at 36 months [16]. The development
of an artificial vascular prosthesis for small diameter applications, like CABG, that can
provide consistent long-term patency would greatly improve treatment outcomes for
these patients.
1.3.4. Failure Mechanisms of Bypass Grafts
Bypass graft failure can be divided into early and late regimes. The main causes
of early graft failure include thrombosis, infection, and surgical and/or material failures
of the graft [11]. Thrombosis is the most important cause of early graft failure. Between
3-12% of saphenous vein bypass grafts are occluded by thrombosis within 1 month of implantation [9]. Grafts made from ePTFE fare worse. In a study of 7 mm femoral- popliteal ePTFE bypass grafts, 5% were occluded by thrombosis after 1 month and 18% failed by 23 months [17], with rates for smaller diameter ePTFE grafts even higher.
Approximately 4% of grafts (including aortic replacements, bypass grafts, and
4 arteriovenous shunts) become infected [18]. Fibro-proliferative stenosis, known as
intimal hyperplasia (IH), is the major contributor to late failure, either by directly causing
stenosis or by providing a background for the development of atherosclerosis and
thrombosis.
1.4. Anastomotic Intimal Hyperplasia
1.4.1. Pathological Characterization
IH can occur in all types of vascular conduits, including artery, vein, and
prosthetic grafts and in arterio-venous shunts. The frequency of failure due to IH is prosthetic > vein > artery [8, 9, 19]. In one study using vein grafts for inguinal-to-below- the-knee popliteal bypass grafting, in which failure mechanisms were carefully documented, about 4% of grafts failed due to IH after 1-2 y of follow up, representing roughly 20% of all graft failures [20]. The contribution of IH to graft failure is thought to increase significantly after one year [8, 9]. IH also can provide a foundation for subsequent atherosclerosis [8, 9].
While this discussion focuses on IH in vascular grafts, IH can be induced in a number of situations, including endothelial denudation and wall injury during angioplasty
[9, 21, 22]. The pathogenesis of IH initiated by these causes share many features.
Normally, smooth muscle cells (SMCs) in the vessel wall adopt a quiescent, contractile phenotype. Following the inciting incident, SMCs, and perhaps adventitial fibroblasts
[23], change phenotype (see Chapter 2) [21]. Secretion of matrix metalloproteinases is up-regulated and SMCs penetrate the internal elastic lamina where they proliferate and elaborate extracellular matrix in the intima [2, 24]. Later in the development of the lesion synthetic SMCs re-differentiate toward a contractile phenotype [25].
5 The literature often does not distinguish IH caused by endothelial injury, angioplasty, or graft placement, although clearly, the stimulus for each is distinct.
Intimal hyperplasia found at the distal anastomosis of vascular grafts has been
characterized less extensively than the IH induced by endothelial damage and/or
angioplasty. The pathology of distal anastomotic IH (DAIH) is characterized by a typical
spatial distribution around the end-to-side anastomoses typically employed for these
procedures (Fig. 1.1). The lesions in DAIH occur at the suture line, primarily in the
“heel” and “toe” and on the “floor” of the outflow track in the native vessel (see Fig. 1.1)
[22, 26]. Like IH induced by other stimuli, the lesions are populated by SMCs that lack
myofilaments in their cytoplasm, possess extensive rough endoplasmic reticulum, and
secrete collagen [22]. One important distinction is that in DAIH with prosthetic grafts,
hyperplastic tissue grows over the surface of the prosthesis, to form a new layer, known
as the neointima [22]. These cells can be of anastomotic or transmural origin, depending
on the porosity of the prosthetic graft and distance from the anastomosis [27]. As the
pathology progresses, SMCs, particularly in the periphery of the lesion, begin to re-
express myofilaments and orient in the direction of blood flow [22]. The re-expression of
SMC marker proteins is consistent with lesions generated by angioplasty or endothelial
denudation [25].
The highly stereotyped location of lesion development suggests there are some
underlying causes of DAIH that may be distinct from IH in general. However, the
pathophysiology of this process is understood poorly and is related to a complex network
of mechanical and biological factors. These factors will be briefly reviewed in the next
several sections.
6
End-to-side Anastomosis
g ra ft
toe heel artery IH IH Suture line
IH Occlusion floor
Intimal hyperplasia (IH)
Figure 1.1-Typical distribution of intimal hyperplasia IH around a distal end-to-side anastomosis.
7 1.4.2. The Role of Injury
During graft placement, injury at the graft-artery anastomosis cannot be avoided, due to the necessity of incision and suturing. Direct trauma to smooth muscle tissue releases basic fibroblast growth factor (bFGF), which is normally stored in the cytoplasm of SMCs [21]. Endothelial damage stimulates platelet derived growth factor (PDGF) release from activated platelets [21]. PDGF and bFGF are potent mitogens for SMCs.
Coagulation factors such as α-thrombin and factor Xa can also stimulate SMC proliferation [28]. Injury also causes local inflammation. Inflammatory cells have been noted at the anastomoses and can play an important role in the development of IH [9, 24,
29]. For example, several days after endothelial denudation injury macrophages appear in the resulting lesion of proliferating SMCs [30, 31]. Disrupting the accumulation of these cells (using an anti-CD4 antibody) resulted in decreased SMC hyperplasia in vivo, suggesting these cells play an important role in the process [29]. Inflammatory cytokines including interleukin-1α [32], interleukin-8 [33], and tumor necrosis factor-α [34] can stimulate SMC proliferation. These stimuli can also induce the endogenous production of inflammatory cytokines by SMCs to generate a positive feedback cycle [30, 31, 35].
These factors, in concert with other cytokines, can initiate proliferation and migration of previously quiescent SMCs [21, 36].
One of the typical locations of DAIH is along the suture line, particularly at the heel and toe of the graft. Clearly, the insertion of sutures induces local injury to both endothelial cells and SMCs, which may contribute to IH development in these regions.
Furthermore, the relatively rigid ePTFE grafts and the even more rigid sutures, introduces a discontinuity in mechanical properties, known as compliance mismatch [37]. As
8 discussed below, compliance mismatch causes a variety of issues to arise. One such issue
is the concentration of stress at the graft-artery or suture-artery interface which, with the cyclic loading of the cardiac cycle, may induce chronic injury providing a continuous stimulus for inflammation and accompanying hyperplasia.
In an effort to better understand what other driving forces may be relevant to
DAIH progression, many researchers have investigated the correlation between the locations of DAIH and hemodynamic and mechanical conditions, using both in vivo and in vitro experiments.
1.4.3. The Role of Mechanical and Hemodynamic Factors: Animal Studies
The hypothesis that the driving force for IH is a mismatch in mechanical
properties has existed for nearly 30 years. The initial hypothesis was based largely on a
correlation between graft compliance and patency [19]. Walden and co-workers
measured the compliance of several graft types, ranging from glutaraldehyde treated
umbilical veins to ePTFE, and compared this data with clinical results [19]. Although a
clear correlation between compliance and 2-year patency was found, the biological and
physical properties of the grafts assessed varied widely. Although acknowledged by the
authors, these differences make interpretation of these data very difficult [11].
To overcome these problems, the same authors developed a model system using
gluteraldehyde to variably cross-link grafts formed from the explanted carotid artery [38].
“Compliant” and “stiff” grafts were prepared using this technique and implanted in dogs
with an end-to-end anastomosis for 12 weeks [38]. While the “stiff” grafts did result in
more failures (only 8/14 “stiff” failures versus 2/14 “compliant” grafts), failure occurred
exclusively by thrombosis and no evidence of IH was noted [38]. Again, significant
9 biological differences between lightly cross-linked and heavily cross-linked carotid
tissues were ignored completely. Inflammation, which certainly varied between these
two preparations, was not assessed. As a result, it is difficult to extrapolate from these
data the role of compliance mismatch per se on the development of graft failure.
Although the scientific evidence from these studies is weak, the results are of note
because they formed the foundation for the compliance mismatch hypothesis [39].
Dobrin and co-workers used various surgical set-ups with autologous saphenous
vein grafts to explore the role mechanics play in more detail, including compliance, longitudinal stress, and shear stress (adjusted by blood flow rate) [40]. They found the
greatest IH in grafts that had been placed in parallel with existing flow (reduced flow)
compared with an analogous graft where the parallel circuit had been ligated (high flow)
[40]. Furthermore, when reinforcing polymer meshes were placed around the veins to
prevent distension, IH and medial thickening was greatest in the non-reinforced regions,
suggesting wall strain may be one driving force for IH [40]. However, they did not
examine the transition regions between stiff and compliant materials in detail, so
compliance mismatch was not tested directly. On the basis of these results, the authors
argue that IH at the end-to-side anastomoses, which is typical for bypass grafting
procedures, is due to flow velocity because of the wide diameter at this point. However,
this overly simplistic argument completely ignores the complex mechanical and
hemodynamic environment in this complex geometry.
To further explore the role of shear in IH development, Kraiss and co-workers
utilized a serial-parallel implantation model where one 5 mm ePTFE graft was placed as
an aorto-aorto bypass (high shear) and two grafts were placed as aorto-iliac bypasses
10 (low shear) in baboons [41]. After 12 weeks, the low shear grafts developed greater
intimal thickening (5.9 vs. 3.4 mm2, p < 0.05) and had a greater rate of SMC proliferation
(0.24% vs. 0.14%, p < 0.05) [41]. However, the grafts in this study were completely
endothelialized at harvest and intimal thickening at the anastomoses was no greater than
in the mid-graft region [41]. Based on these limitations, it is difficult to extrapolate these
results to the complex flow environment of the anastomosis, where many factors may contribute to IH.
To more carefully assess the effect that hemodynamics have on the development
of anastomotic IH, Keynton et al. studied IH in dogs with end-to-side placed ePTFE
grafts with graft-to-artery diameter ratios of 1.0 and 1.5 [42]. They found that all
anastomoses developed IH in the floor region, but grafts with a diameter ratio of 1.5
developed greater IH in the toe region of the graft [42]. A strong inverse correlation was
observed between shear rate and IH thickness at any given location in the anastomosis (r
= 0.95) [42]. However, this correlation appears to be dominated by the floor IH and does not explain differences in toe IH, since shear in these regions was roughly the same for
both diameter ratios.
In a simpler study to characterize differences in IH between prosthetic and vein grafts, Bassiouny and co-workers characterized IH development in dogs with either ePTFE or saphenous vein grafts, all with the same fixed end-to-side anastomosis geometry [26]. They noted that while IH at the suture line was greater for the ePTFE
grafts, perhaps due to compliance mismatch, IH on the graft floor was independent of
graft type [26]. Based on observations that vein grafts have less suture line IH than
prosthetic grafts, surgeons have implemented the use of transitional cuffs of vein tissue
11 between the synthetic prosthesis and the native artery [43, 44]. Trubel and colleagues
found that the benefit of these patches is not that they reduce IH development, but rather
shift IH toward regions that can more effectively accommodate the hyperplastic tissue
(i.e. away from the relatively narrow floor-toe region) [45]. IH was still found in the
typical places: at the suture lines and at the floor of the blood vessel [45].
Despite limitations, in vivo experiments highlight several general principles
regarding IH formation. Low shear stress promotes IH, at least in systems where it can
be sensed by endothelial cells. Stiff mechanical properties of the graft are correlated with
IH formation, although a mechanism has not been clearly defined in vivo. IH at the floor
of the anastomosis appears to be unrelated to graft type, suggesting it is a consequence of the end-to-side anastomotic geometry and the hemodynamic conditions that it causes.
However, the role of non-injury related causes of suture line IH are still unclear. To overcome the experimental limitations of in vivo studies, many groups have also explored models of graft anastomoses to correlate DAIH formation with mechanical and hemodynamic factors which can be more easily measured and controlled in an in vitro setting.
1.4.4. The Role of Mechanical and Hemodynamic Factors: Modeling
Flow patterns at end-to-side anastomoses have been observed by several groups to
try to correlate aberrant flow profiles with regions of IH formation. Since floor IH was
independent of graft type in animal studies, flow patterns in this region were of particular
interest. A stagnation point has been observed in this region using a rigid end-to-side
anastomosis model with steady laminar flow, which varied in location with anastomotic
angle [46]. Although this model simplifies the complex flow conditions found in vivo,
12 these results may approximate flow toward the end of each pulse. Using silicone casts of
explanted anastomoses from their in vivo studies, Bassiouny and co-workers also observed a stagnation point with low and oscillating shear stress in the floor of the outflow tract that moved back and forth during the cardiac cycle [26]. Using a photochromic tracer, Ojha et al. studied both steady and pulsatile flow at the anastomosis
[47]. A stagnation point at the floor, surrounded by with divergent and helical flow patterns, was also observed in these studies [47]. Additionally, during pulsatile flow, an
oscillating shear pattern in the toe region was observed that correlates with IH
development in this region [47]. These studies clearly indicate the formation of a
stagnation point in the floor of the anastomosis. Coupled with clear in vivo evidence of
increased IH under low flow conditions [40, 41] and the reproducible development of
floor lesions, these results suggest aberrant flow is the likely cause of floor IH. These studies also suggest that other flow disturbances generated during pulsatile flow may be correlated with suture line IH.
Ballyk et al. performed a finite-element numerical simulation to better understand the mechanical environment at this complex suture line interface [48]. They found that compliance mismatch caused an increase in stress concentration near the suture line, but only in the toe region of end-to-side anastomoses, suggesting these increased suture stress concentration may be driving IH at the toe [48]. As noted above, these stress concentrations also may induce continuous injury and inflammation, contributing to IH.
Compliance mismatch may also directly affect the flow pattern near the suture line, particularly at the heel and toe of the anastomosis. Study of flow profiles at various simulated end-to-end graft anastomoses suggest a stiff-to-compliant transition or a small-
13 to-large diameter mismatch can result in reduced shear stress, which may contribute to IH
[49].
1.4.5. Conclusions
The results of modeling and in vivo investigations suggest there is a correlation between flow patterns, mechanical mismatch, and DAIH. The presence of a stagnation
point at the floor of the anastomosis provides compelling evidence that IH in this region
is flow derived. The relative roles of compliance mismatch, hemodynamics, and injury at
the suture line sites of IH have yet to be resolved convincingly, but it is likely that all
three factors play an important role and are interrelated. However, these results suggest
that the development of materials with well-matched mechanical and biological
properties, which also modulate the response to injury, have the potential to reduce suture
line IH. Such materials may provide enhanced long-term graft patency, improving
outcomes for patients with CHD and peripheral artery disease.
14 1.5. References
1. Lloyd-Jones D, Adams R, Carnethon M, De Simone G, Ferguson TB, Flegal K, Ford E, Furie K, Go A, Greenlund K, Haase N, Hailpern S, Ho M, Howard V, Kissela B, Kittner S, Lackland D, Lisabeth L, Marelli A, McDermott M, Meigs J, Mozaffarian D, Nichol G, O'Donnell C, Roger V, Rosamond W, Sacco R, Sorlie P, Stafford R, Steinberger J, Thom T, Wasserthiel-Smoller S, Wong N, Wylie-Rosett J, Hong Y, for the American Heart Association Statistics Committee and Stroke Statistics S. Heart Disease and Stroke Statistics--2009 Update: A Report From the American Heart Association Statistics Committee and Stroke Statistics Subcommittee. Circulation 2009;119(3):e21-181.
2. Kumar V, Abbas AK, Fausto N, Robbins SL, Cotran RS. Robbins and Cotran pathologic basis of disease. 7th ed. Philadelphia: Elsevier Saunders, 2005.
3. Ross R. Atherosclerosis -- An Inflammatory Disease. N Engl J Med 1999;340(2):115-126.
4. Pearson TA, Blair SN, Daniels SR, Eckel RH, Fair JM, Fortmann SP, Franklin BA, Goldstein LB, Greenland P, Grundy SM, Hong Y, Houston Miller N, Lauer RM, Ockene IS, Sacco RL, Sallis JF, Jr., Smith SC, Jr., Stone NJ, Taubert KA. AHA Guidelines for Primary Prevention of Cardiovascular Disease and Stroke: 2002 Update: Consensus Panel Guide to Comprehensive Risk Reduction for Adult Patients Without Coronary or Other Atherosclerotic Vascular Diseases. Circulation 2002;106(3):388-391.
5. Smith SC, Jr., Allen J, Blair SN, Bonow RO, Brass LM, Fonarow GC, Grundy SM, Hiratzka L, Jones D, Krumholz HM, Mosca L, Pasternak RC, Pearson T, Pfeffer MA, Taubert KA. AHA/ACC Guidelines for Secondary Prevention for Patients With Coronary and Other Atherosclerotic Vascular Disease: 2006 Update: Endorsed by the National Heart, Lung, and Blood Institute. Circulation 2006;113(19):2363-2372.
6. Hannan EL, Wu C, Walford G, Culliford AT, Gold JP, Smith CR, Higgins RS, Carlson RE, Jones RH. Drug-eluting stents vs. coronary-artery bypass grafting in multivessel coronary disease. N Engl J Med 2008;358(4):331-341.
7. Cameron A, Davis KB, Green G, Schaff HV. Coronary Bypass Surgery with Internal-Thoracic-Artery Grafts -- Effects on Survival over a 15-Year Period. N Engl J Med 1996;334(4):216-220.
8. Nwasokwa ON. Coronary-Artery Bypass Graft Disease. Ann Intern Med 1995;123(7):528-545.
9. Motwani JG, Topol EJ. Aortocoronary saphenous vein graft disease: pathogenesis, predisposition, and prevention. Circulation 1998;97(9):916-931.
15 10. Laube HR, Duwe J, Rutsch W, Konertz W. Clinical experience with autologous endothelial cell-seeded polytetrafluoroethylene coronary artery bypass grafts. J Thorac Cardiovasc Surg 2000;120(1):134-141.
11. Greenwald SE, Berry CL. Improving vascular grafts: the importance of mechanical and haemodynamic properties. J Pathol 2000;190(3):292-299.
12. Hehrlein FW, Schlepper M, Loskot F, Scheld HH, Walter P, Mulch J. The use of expanded polytetrafluoroethylene (PTFE) grafts for myocardial revascularization. J Cardiovasc Surg (Torino) 1984;25(6):549-553.
13. Sapsford RN, Oakley GD, Talbot S. Early and late patency of expanded polytetrafluoroethylene vascular grafts in aorta-coronary bypass. J Thorac Cardiovasc Surg 1981;81(6):860-864.
14. Chard RB, Johnson DC, Nunn GR, Cartmill TB. Aorta-coronary bypass grafting with polytetrafluoroethylene conduits. Early and late outcome in eight patients. J Thorac Cardiovasc Surg 1987;94(1):132-134.
15. Sayers RD, Raptis S, Berce M, Miller JH. Long-term results of femorotibial bypass with vein or polytetrafluoroethylene. Br J Surg 1998;85(7):934-938.
16. Faries PL, Logerfo FW, Arora S, Hook S, Pulling MC, Akbari CM, Campbell DR, Pomposelli FB, Jr. A comparative study of alternative conduits for lower extremity revascularization: all-autogenous conduit versus prosthetic grafts. J Vasc Surg 2000;32(6):1080-1090.
17. Veith FJ, Gupta S, Daly V. Management of early and late thrombosis of expanded polytetrafluoroethylene (PTFE) femoropopliteal bypass grafts: favorable prognosis with appropriate reoperation. Surgery 1980;87(5):581-587.
18. Darouiche RO. Current concepts - Treatment of infections associated with surgical implants. N Engl J Med 2004;350(14):1422-1429.
19. Walden R, L'Italien GJ, Megerman J, Abbott WM. Matched elastic properties and successful arterial grafting. Arch Surg 1980;115(10):1166-1169.
20. Donaldson MC, Mannick JA, Whittemore AD. Causes of primary graft failure after in situ saphenous vein bypass grafting. J Vasc Surg 1992;15(1):113-118; discussion 118-120.
21. Reidy MA. Regulation of Arterial Smooth Muscle Growth. In: Schwartz SM, Mecham RP, editors. The vascular smooth muscle cell : molecular and biological responses to the extracellular matrix. San Diego: Academic Press, 1995. p. xvi, 410 p.
16 22. Sottiurai VS, Yao JS, Batson RC, Sue SL, Jones R, Nakamura YA. Distal anastomotic intimal hyperplasia: histopathologic character and biogenesis. Ann Vasc Surg 1989;3(1):26-33.
23. Li G, Chen SJ, Oparil S, Chen YF, Thompson JA. Direct in vivo evidence demonstrating neointimal migration of adventitial fibroblasts after balloon injury of rat carotid arteries. Circulation 2000;101(12):1362-1365.
24. Raines EW, Koyama H, Carragher NO. The extracellular matrix dynamically regulates smooth muscle cell responsiveness to PDGF. Ann N Y Acad Sci 2000;902:39-51; discussion 51-32.
25. Christen T, Verin V, Bochaton-Piallat M, Popowski Y, Ramaekers F, Debruyne P, Camenzind E, van Eys G, Gabbiani G. Mechanisms of neointima formation and remodeling in the porcine coronary artery. Circulation 2001;103(6):882-888.
26. Bassiouny HS, White S, Glagov S, Choi E, Giddens DP, Zarins CK. Anastomotic intimal hyperplasia: mechanical injury or flow induced. J Vasc Surg 1992;15(4):708- 716; discussion 716-707.
27. Clowes AW, Kirkman TR, Reidy MA. Mechanisms of arterial graft healing. Rapid transmural capillary ingrowth provides a source of intimal endothelium and smooth muscle in porous PTFE prostheses. Am J Pathol 1986;123(2):220-230.
28. Rauch BH, Millette E, Kenagy RD, Daum G, Clowes AW. Thrombin- and factor Xa- induced DNA synthesis is mediated by transactivation of fibroblast growth factor receptor-1 in human vascular smooth muscle cells. Circ Res 2004;94(3):340-345.
29. Hancock WW, Adams DH, Wyner LR, Sayegh MH, Karnovsky MJ. CD4+ mononuclear cells induce cytokine expression, vascular smooth muscle cell proliferation, and arterial occlusion after endothelial injury. Am J Pathol 1994;145(5):1008-1014.
30. Tanaka H, Sukhova GK, Swanson SJ, Clinton SK, Ganz P, Cybulsky MI, Libby P. Sustained Activation of Vascular Cells and Leukocytes in the Rabbit Aorta after Balloon Injury. Circulation 1993;88(4):1788-1803.
31. Okamoto E, Couse T, De Leon H, Vinten-Johansen J, Goodman RB, Scott NA, Wilcox JN. Perivascular inflammation after balloon angioplasty of porcine coronary arteries. Circulation 2001;104(18):2228-2235.
32. Schultz K, Murthy V, Tatro JB, Beasley D. Endogenous interleukin-1 alpha promotes a proliferative and proinflammatory phenotype in human vascular smooth muscle cells. Am J Physiol Heart Circ Physiol 2007;292(6):H2927-2934.
33. Yue TL, Wang X, Sung CP, Olson B, McKenna PJ, Gu JL, Feuerstein GZ. Interleukin-8. A mitogen and chemoattractant for vascular smooth muscle cells. Circ Res 1994;75(1):1-7.
17 34. Heo SK, Yun HJ, Park WH, Park SD. Emodin inhibits TNF-alpha-induced human aortic smooth-muscle cell proliferation via caspase- and mitochondrial-dependent apoptosis. J Cell Biochem 2008;105(1):70-80.
35. Rose SL, Babensee JE. Complimentary endothelial cell/smooth muscle cell co- culture systems with alternate smooth muscle cell phenotypes. Ann Biomed Eng 2007;35(8):1382-1390.
36. Kohler TR. Intimal Hyperplasia. In: White RA, Hollier LH, editors. Vascular surgery : basic science and clinical correlations. 2nd ed. Elmsford, N.Y.: Futura, 2004. p. p.
37. Salacinski HJ, Goldner S, Giudiceandrea A, Hamilton G, Seifalian AM, Edwards A, Carson RJ. The mechanical behavior of vascular grafts: a review. J Biomater Appl 2001;15(3):241-278.
38. Abbott WM, Megerman J, Hasson JE, L'Italien G, Warnock DF. Effect of compliance mismatch on vascular graft patency. J Vasc Surg 1987;5(2):376-382.
39. Chan-Park MB, Shen JY, Cao Y, Xiong Y, Liu Y, Rayatpisheh S, Kang GC, Greisler HP. Biomimetic control of vascular smooth muscle cell morphology and phenotype for functional tissue-engineered small-diameter blood vessels. J Biomed Mater Res A 2009;88(4):1104-1121.
40. Dobrin PB, Littooy FN, Endean ED. Mechanical factors predisposing to intimal hyperplasia and medial thickening in autogenous vein grafts. Surgery 1989;105(3):393-400.
41. Kraiss LW, Kirkman TR, Kohler TR, Zierler B, Clowes AW. Shear stress regulates smooth muscle proliferation and neointimal thickening in porous polytetrafluoroethylene grafts. Arterioscler Thromb 1991;11(6):1844-1852.
42. Keynton RS, Evancho MM, Sims RL, Rodway NV, Gobin A, Rittgers SE. Intimal hyperplasia and wall shear in arterial bypass graft distal anastomoses: an in vivo model study. J Biomech Eng 2001;123(5):464-473.
43. Miller JH, Foreman RK, Ferguson L, Faris I. Interposition vein cuff for anastomosis of prosthesis to small artery. Aust N Z J Surg 1984;54(3):283-285.
44. Taylor RS, Loh A, McFarland RJ, Cox M, Chester JF. Improved technique for polytetrafluoroethylene bypass grafting: long-term results using anastomotic vein patches. Br J Surg 1992;79(4):348-354.
45. Trubel W, Schima H, Czerny M, Perktold K, Schimek MG, Polterauer P. Experimental comparison of four methods of end-to-side anastomosis with expanded polytetrafluoroethylene. Br J Surg 2004;91(2):159-167.
18 46. Keynton RS, Rittgers SE, Shu MC. The effect of angle and flow rate upon hemodynamics in distal vascular graft anastomoses: an in vitro model study. J Biomech Eng 1991;113(4):458-463.
47. Ojha M, Ethier CR, Johnston KW, Cobbold RS. Steady and pulsatile flow fields in an end-to-side arterial anastomosis model. J Vasc Surg 1990;12(6):747-753.
48. Ballyk PD, Walsh C, Butany J, Ojha M. Compliance mismatch may promote graft- artery intimal hyperplasia by altering suture-line stresses. J Biomech 1998;31(3):229- 237.
49. Weston MW, Rhee K, Tarbell JM. Compliance and diameter mismatch affect the wall shear rate distribution near an end-to-end anastomosis. J Biomech 1996;29(2):187-198.
19 CHAPTER 2: Regulation of SMC Phenotype: Implications for Vascular Tissue Engineering
2.1. Introduction
The goal of vascular tissue engineering is to generate functional vascular replacements that resist infection, thrombosis, and restenosis due to intimal hyperplasia
(IH). Although some progress has been made in minimizing early thrombosis [1, 2] and infection [3, 4], minimizing IH remains a key criterion for long-term graft patency. In this chapter, the molecular regulation of smooth muscle cell (SMC) behaviors, including those that result in IH, and implications for vascular tissue engineering will be reviewed.
Central to the development of IH and other vascular pathologies, such as restenosis and atherosclerosis, is a dysregulation of vascular SMC phenotype characterized by a loss of contractile proteins and a shift from cellular quiescence to increased production of extracellular matrix proteins, proliferation, and migration. While in vivo this phenotypic switch results in vascular stenosis and undesirable clinical outcomes, this remarkable phenotypic plasticity can be harnessed to form functional arteries ex vivo [5]. This approach can yield autologous, immunocompatible tissue without the need for elaborate and time consuming harvesting, purification, and expansion protocols necessary for progenitor cells, such as mesenchymal stem cells, and without the ethical issues associated with embryonic stem cells. Furthermore, this phenotypic plasticity also invites the possibility that a synthetic vascular conduit can be designed that guides tissue regeneration in vivo from adjacent native smooth muscle cells without the need for ex vivo culture. Aside from reducing production costs, this off-the- shelf capacity would greatly improve the technology’s utility for emergent revascularization procedures, such as coronary artery bypass grafting (CABG), where
20 there is a pressing clinical need for synthetic substitutes for patients without suitable autologous vessels.
In order to harness this potential, it is necessary to understand and control the molecular mechanisms underlying SMC phenotype switching. To recapitulate the function of vascular smooth muscle in a tissue engineered blood vessel (TEBV), SMC proliferation is required initially to populate the construct, and extracellular matrix
(ECM) deposition and remodeling are required to provide the appropriate mechanical strength and tissue architecture. Eventually these proliferative, synthetic SMCs must re- differentiate to a quiescent, contractile state. These contractile SMCs can regulate vascular tone and will be refractory to proliferative signals that might otherwise drive IH.
These processes require activation of diverse (and often opposing) cellular programs which must be appropriately controlled both spatially and temporally. Recent work on the cell and molecular biology of SMCs has elucidated many intra- and extracellular factors that affect SMC phenotype [6]. Application of this information to the field of vascular tissue engineering is critical for the development of complex, bioactive scaffold systems that can control SMC behavior.
2.2. The Continuum of SMC Phenotypes
In vivo, smooth muscle tissues play an important role in a wide range of systems from the vasculature to reproduction. In order to perform a diversity of functions, SMC phenotype spans a continuum from quiescent and contractile to proliferative and synthetic [7, 8]. At the contractile extreme are SMCs with a fully functional contractile apparatus that responds to small molecule signals of physiological significance such as acetylcholine and norepinephrine. In early studies, these cells were characterized by an
21 ultrastructure (observed by transmission electron microscopy) composed of tightly bundled myofilaments and minimal rough endoplasmic reticulum (ER), golgi, or free
ribosomes [9-11]. Contractile smooth muscle tissues also generally contained little
connective tissue that would necessitate extensive SMC synthetic capacity [9]. In
culture, these cells possess a dense fusiform morphology [9, 12]. At the synthetic
phenotype extreme of this continuum are fibroblast-like SMCs, which contain minimal
contractile proteins and secrete ECM. The ultrastructure of these cells shows a cytoplasm
devoid of contractile bundles with extensive rough ER, golgi and ribosomes [9-11]. In
culture, these cells initially adopt a broad, spread shape, then begin to grow over one
another in a “hill-and-valley” morphology [9, 12]. A synthetic phenotype is also
correlated with SMC proliferation, with the number of S-phase cultured SMCs increasing
from 3-5% to 40-60% during primary culture and pathologies such as IH [9]. Most
SMCs, even SMCs in contractile tissues, lie somewhere along the continuum. For
example, SMCs in small muscular arteries typically have 80-90% of their cytoplasm
filled by myofilaments, whereas SMCs in the aorta typically contain only 60-70%,
indicating that aortic SMCs have both contractile and synthetic functions [9].
Between these extremes there are many markers that indicate a cell’s position
along this continuum. Contractile SMCs, which predominate in normal vessels, exhibit a
mature contractile apparatus including smooth muscle α-actin (SMαA), smooth muscle
myosin heavy chains SM1 and SM2, and calponin. The relative expression of these
proteins can be used to localize SMCs on the contractile-synthetic continuum (discussed
extensively below) [7, 13]. An overview of the characteristics of each phenotype can be
found in Fig. 2.1.
22
Synthetic SMC Contractile SMC
A As c carbachol carbachol
B s Bc
C. Expression of Selected Phenotypic Markers Synthetic Contractile Med.-Low α-actin High Low SM-MHC High Low calponin High Low SM22-α High Low h-caldesmin High High l-caldesmin Low
Figure 2.1-Summary of characteristics of SMC phenotype, which varies along a continuum from synthetic and proliferative to contractile and quiescent. As,c. Ligand induced contractility is the most rigorous definition of contractile SMC phenotype (Images: [14]). Bs,c. The cellular ultrastructure when viewed by TEM shows the presence of abundant rough endoplasmic reticulum (arrows) in the synthetic phenotype while myofilaments are the predominant feature in contractile cells (Images: [10, 15]). C. Additional marker proteins commonly used to study SMC phenotype [7].
23 2.3. Markers of Contractile SMC Phenotype
The goal of generating contractile smooth muscle tissue requires a functional
understanding of many cell systems including receptors, signal transduction machinery, and a functional contractile apparatus consisting of many diverse proteins. Since SMCs lose their contractility early during the contractile-to-synthetic transition that takes place in culture, it is convenient to monitor the expression of marker proteins that more precisely indicate where on the synthetic-contractile continuum the SMC lies. Indeed many of the proteins used as markers are part of the SMC’s contractile apparatus.
Moreover, the contractility of an SMC is not the only phenotypic characteristic that describes the “contractile” SMCs that populate healthy vasculature. Marker proteins can also be correlated with the SMC’s likelihood to respond to mitogens [16, 17] or can characterize the composition of secreted ECM proteins which differ between contractile and synthetic SMCs [18]. This section will briefly review some of the commonly utilized markers of contractile phenotype, but is by no means a comprehensive listing of all relevant markers. For a more detailed review, the reader is referred to Owens [7].
2.3.1. Smooth Muscle α-Actin
Smooth muscle α-actin (SMαA) is a 43 kD protein that is one of 6 actin isoforms
(3-α, 1-β, 2-γ: ACTA1, ACTA2, ACTC1, ACTB, ACTG1, ACTG2) individually
encoded in the human genome [19, 20]. SMCs express the smooth muscle forms of α-
and γ-actin as well as β-actin and non-muscle γ-actin [21]. The fraction of SMαA
expression in vascular SMCs increases until 30 days after birth in rats [21]. Functionally,
SMαA, as the predominant actin isoform, forms the thin filament of the classical
contractile apparatus. Given the diverse transcriptional regulation of the various actin
24 isoforms, evolutionarily it would be expected that, despite their high degree of homology,
each plays a specialized role in SMC contraction. Although the functional significance of
each SMC actin isoform is not completely understood, there is some evidence that they
play differential roles in cytoskeletal remodeling or mediating contraction from various stimuli [22].
SMαA is the most ubiquitous marker of SMC lineage and differentiated
phenotype used for research purposes. It is, therefore, unfortunate that it is also the least
SMC specific. Although SMαA is one of the earliest markers expressed in SMCs during
their differentiation, it is also expressed temporarily in non-smooth muscle tissues such as
skeletal muscle [23]. In the mature organism, SMαA can also be expressed in
myofibroblasts during the wound healing process, but its expression appears to depend, at
least in part, on separate transcriptional activation pathways than vascular SMCs [24].
There is also evidence that transforming growth factor (TGF)-β1 can induce the
expression of SMαA in endothelial cells [25, 26]. As a result, SMαA should only be used
in conjunction with other markers of contractile phenotype, especially when used to
assess the differentiation status of cultured SMCs.
2.3.2. Smooth Muscle Myosin Heavy Chain (SM-MHC)
Smooth muscle myosin heavy chains (SM-MHC) are a set of four 200-204 kD
proteins that are translated from four alternately spliced transcripts from the same gene
(MYH11) [27]. The four splice variants represent the combination of two unique c-
terminal modules, delineating SM-1 and SM-2, and the presence or absence of a 21 nucleotide exon in near the 5’ region, delineating the A (- exon) and B (+ exon) variants
[27]. The mechanisms regulating SM-MHC splicing have yet to be elucidated, but there
25 is clear evidence of spatial and temporal regulation of expression of all four isoforms [28,
29] suggesting a control system exists. In addition to SM-MHC, SMCs express two
additional gene products, non-muscle myosin heavy chains (NMHC) A (MYH9) and B
(MYH10) [7]. In particular, NMHC-B (also known as SMemb) is expressed in
embryonic aortic SMCs as well as SMCs in intimal lesions and in culture [28, 30, 31].
Functionally, SM-MHC is part of a six-member complex (2 heavy chains, two
non-regulatory myosin light chains and 2 regulatory light chains), that functions as an
ATP-ase to generate force along actin filaments. There is evidence that sequence
differences between the four splice variants are of functional significance. The 21
nucleotide insert, which delineates the B isoform from the A isoform, codes for a change in the amino acid sequence in the ATP binding pocket of SM-MHC and results in dramatic changes in the kinetics of myosin cycling in vitro [32] resulting in increased contractility in vivo [29]. It has been proposed that differential expression of the A and B
isoforms may contribute to functional (tonic versus phasic) differences in smooth muscle
contractility [32]. The c-terminal isoforms also play a critical role in vascular SMC
function. Transgenic mice with a deletion of the SM-2 specific exon died within 30 d
after birth and had hypercontractile smooth muscle tissues, suggesting that SM-2 may
play an important role in modulating contraction [33].
SM-MHC, in particular the SM-2 isoforms, is one of the latest markers of SMC
differentiation with expression in the vasculature increasing during late embryogenesis
and into the post-natal period [28]. The SM-2 isoform, in particular, is lost rapidly from cultured SMCs [34] and also is down-regulated in intimal lesions [31]. For these reasons,
26 SM-MHC, especially the SM-2 isoform, has been proposed as the most definitive marker
of differentiated vascular SMCs [7, 35].
2.3.3. Calponin
Smooth muscle basic (h1) calponin is an approximately 34 kD actin, tropomyosin,
and calmodulin binding protein [36, 37]. Basic h1 calponin (gene: CNN1) is one of three
human calponin genes, but is the only gene product with selective expression in smooth
muscle [37, 38]. Calponin is thought to play a regulatory role in SMC contraction.
Calponin contains a number of actin binding domains that facilitate its co-localization
with actin filaments [39, 40]. Furthermore, calponin co-localizes only in the central
regions of the actin filaments, where actin-myosin interactions predominantly take place,
and is excluded from the cell periphery near cell-matrix adhesions sites [41]. Calponin
inhibits the activity of the myosin ATP-ase in vitro in a calcium dependent fashion [42],
suggesting a direct role in modulating SMC contraction. However, calponin also
contains binding sites for extracellular signal regulated kinase (ERK) and protein kinase
C (PKC) [43, 44], and as a result it has been suggested that calponin serves as an adapter
facilitating localized signaling of these kinases [38]. However, the physiological significance of these in vitro observations is unclear. The most striking phenotype of calponin knockout mice is dysregulation of bone formation [45]. However, these mice also display reduced sensitivity to α-adrenergic vasoconstriction, impaired regulation of blood flow [46, 47], and increased unloaded shortening velocity of smooth muscle preparations ex vivo [48], consistent with multiple modes of regulation. Calponin is an intermediate marker of cell lineage and is expressed after SMαA but before the SM-2
27 isoform of SM-MHC [23]. Although calponin is expressed in cardiac muscle during development, its expression appears to be restricted to smooth muscle in the adult [49].
2.3.4. SM-22α
SM-22α, also known as transgelin, is a 22 kD actin binding protein with substantial homology to calponin [50]. Like calponin, SM-22α likely plays a role in modulating the contractile response. SM-22α interacts with actin and co-localizes with actin filaments in SMCs [51]. SM-22α knockout mice develop normally and do not display any overt deficits in physiologic smooth muscle functions [52]. Explanted smooth muscle tissue from these mice does display reduced calcium-independent response to phenylephrine or phorbal esters (which activate PKC), suggesting SM-22α plays a role transducing contraction signals that do not depend on myosin phosphorylation [53]. During development, SM-22α is expressed both in cardiac and smooth muscle tissue, but its expression appears to be restricted to vascular and visceral smooth muscle in the adult [52, 54, 55], although expression in myofibroblast also has been reported [56]. Expression of SM-22α is an early marker of SMC lineage, occurring after SMαA, but before calponin in the chick embryo [23].
2.3.5. Smoothelin
The smoothelins are a family of recently identified, actin binding, SMC specific marker proteins [57, 58]. Two smoothelin proteins, designated A (59 kD) and B (110 kD), are expressed from one gene (SMTN) by separate promoters, only one of which
(Smoothelin A) contains a CArG box, which is common in the promoters of many other marker genes (see below) [57, 59]. Smoothelin A begins in the middle of the tenth exon of smoothelin B [59]. Each gene product also has three splice variants which determine
28 the c-terminus of the protein [59]. The function of smoothelin has not been well characterized, but it is thought to involve regulation of the contractile apparatus.
Smoothelin binds with actin and co-localizes with actin filaments at the subcellular level
[60, 61]. Smoothelin-A/B knockout mice have severe abnormalities in intestinal smooth muscle development, significantly decreased contractility, and expire at a young age [62].
Mice deficient in only smoothelin-A develop normally, but their vascular smooth muscle displays decreased contractility and increased arterial distension, and the mice develop hypertension [63]. These data suggest a clear role for smoothelin in smooth muscle contraction. Based on studies in the chick embryo, Smoothelin-A is predominantly expressed in visceral smooth muscle of the adult, although it is transiently expressed in the heart and vascular tissues during development [64]. Smoothelin-B is predominantly expressed in the vasculature of the adult, although it is transiently expressed in visceral smooth muscle [64]. Smoothelin is down-regulated in cultured SMCs and in neo-intimal lesions after balloon angioplasty, although after healing, smoothelin expression is re- established in the neo-intima [60, 65]. Culture in heparin appears to help retain smoothelin expression in cultured cells [60], but there is no evidence that it will induce re-expression in cultured cells that have lost smoothelin expression. After it is lost in vitro, smoothelin is not thought to be re-inducable [57], although there have been isolated reports of smoothelin re-expression in cultured SMCs [66]. Furthermore, there is no evidence of smoothelin expression in myofibroblasts [58] nor is smoothelin up-regulated by TGF-β1 [60], which typically up-regulates SMC markers such as SMαA in myofibroblasts [67]. As a result, smoothelin, along with SM-MHC, is an excellent specific marker for differentiated SMCs.
29 2.3.6. Others
A wide array of other marker proteins also have been identified and used for
monitoring SMC phenotype. In particular, the heavy isoform of caldesmon (h-
caldesmon) has SMC selective expression. H-caldesmon is differentiated from light (l)- caldesmon which is present in non-smooth muscle tissue via the product of alternative
splicing of one gene (CALD1) [68]. The l-caldesmon splice variant rapidly becomes the
predominant form in cultured SMCs [69]. Functionally caldesmon also plays a role
regulating SMC contraction [38]. Other markers used include aortic carboxypeptidase-
like protein (ACLP) [70-73], smooth muscle myosin light chain kinase (SM-MLCK) [74,
75], telokin (a c-terminal fragment of SM-MLCK) [73], cysteine and glycine rich protein
1 [74], desmin [72], and focal adhesion kinase related nonkinase (FRNK) [73]. A partial
list of the SMC transcriptome can be found in an excellent review by Miano [76].
2.4. Mediators of SMC Phenotype
Because of the role SMC proliferation plays in vascular pathology,
disproportionate effort has been allocated to studying the mechanisms that promote SMC
proliferation, migration, and other markers of synthetic phenotype. Here, these factors
will be briefly reviewed. However, the focus of this section will be weighted toward
factors that promote contractile SMC phenotype, since this poses the greatest challenge to
vascular tissue engineering, especially in the context of re-differentiating synthetic cells
toward a contractile phenotype.
2.4.1. The Role of Soluble Signaling Factors
Extracellular signaling molecules play a major role in determining the phenotypic fate of vascular SMCs. A wide variety of signaling factors have been implicated in the
30 transition of SMCs into the proliferative, synthetic phenotype including platelet-derived
growth factor (PDGF), basic fibroblast growth factor (bFGF), insulin-like growth factors
(IGFs), epidermal growth factor (EGF), α-thrombin, factor Xa, angiotensin II, endothelin-
1, and unsaturated lysophosphatidic acids [69, 77-81]. In vitro fetal bovine serum (FBS)
is also commonly used to stimulate SMC proliferation and de-differentiation. An
overview of these signaling pathways is shown in Fig. 2.2.
The array of extracellular signaling factors that can prevent SMC de-
differentiation and proliferation and/or promote contractile phenotype are fewer in
number and include: IGF-1 (limited to primary SMC isolates) [82], angiotensin II,
soluble heparin, and TGF-β1.
2.4.1.1. Heparin
The ability of heparin, one of many glycosaminoglycans with a semi-repeating
disaccharide structure (Fig. 2.3), to inhibit SMC proliferation has been well described in
vivo and in vitro [79, 80, 83, 84]. Although the effect of heparin has been known for
some time, the mechanism of this effect still is incompletely understood and appears to
be multi-factorial in nature.
The relationship between heparin structure and its anti-proliferative activity has
been studied extensively [85]. To better understand the structural determinants of
heparin function, it is useful to briefly review heparin biosynthesis and structure.
Heparin is synthesized from a repeating disaccharide of D-glucuronic acid (GlcA) and D- glucosamine (GlcN) [85]. Subsequently, the amino group is either sulfated or acetylated,
C-5 in the GlcA is epimerized (swapped about its chiral center) to generate L-iduronic
acid (IdoA), and the hydroxyl groups on the C-6 (or rarely C-3) of the GlcN derivatives
31
α-thrombin factor Xa Heparin Endothelin-1 (Unknown Mechanism) LPA PDGF Angiotensin II EGF IGF Elastin Fibronectin Laminin bFGF TGF-β1
GPCR GPCR α5β1 M TK any R Integrin Re cep tors TK TG R Fβ SPG R +H SYNTHETIC ? CONTRACTILE Figure 2.2-Brief overview of mechanisms involved in the modulation of SMC phenotype. The mechanism of action for heparin is unclear and may act by inhibiting binding of extracellular growth factors or secondary autocrine signaling factors, inhibiting intracellular signal transduction by these stimuli, and/or directly promote contractile phenotype. Angiotensin II action can induce both synthetic and contractile characteristics. Abbreviations: bFGF-basic fibroblast growth factor, PDGF-platelet derived growth factors, EGF-epidermal growth factors, IGF-insulin like growth factors, LPA-lysophosphatidic acid, TGF-β1-transforming growth factor beta 1, RTK-receptor tyrosine kinase, HSPG-heparan sulfate proteoglycan, GPCR-G-protein coupled receptor, TGFβR-TGF-β receptor. (Many references contributed to this diagram, see text for details).
32 and the on C-2 of the IdoA (or GlcA) acid are sulfated [85]. Heparin has a larger fraction of IdoA and higher degree of sulfation than its cousin, heparan sulfate [85]. An overview of heparin structure can be found in Fig. 2.3.
The precise structural determinants of antiproliferative activity have yet to be fully explained. Non-anticoagulant heparin effectively inhibits SMC proliferation in vitro [86] and in vivo [87] and it has been well established that heparin’s antiproliferative activity is unrelated to its anticoagulant activity. Heparin chain lengths greater than 10-
12 repeats (roughly 7 kD) seem to exhibit roughly similar degrees of anti-proliferative activity (as a function of mass concentration) while tetrasaccharide heparin derivatives typically have minimal activity [86, 88, 89]. Heparin chains with 5-10 repeats give inconsistent results, with some studies showing antiproliferative activity [86, 89, 90] while others do not [88]. Significant differences in activity between suppliers [89, 91] and cell types [86, 89] may partially explain these differences.
Sulfation patterns at the 4 sulfation sites in heparin also influence antiproliferative activity. Heparin’s antiproliferative activity does not correlate with the presence of the anticoagulant pentasaccharide motif containing a N-sulfo-D-glucosamine-3-sulfate sugar
(which is rare in full length heparin) [91], though studies only examining the effects of
the pentasaccharide found 3-O-sulfation to increase activity slightly [89]. Substitution of
N-acetylation for N-sulfation resulted in a slight [92] to significant [86] decrease in
antiproliferative activity. Likewise, selective removal of 6-O-sulfate [92] or 2-O-sulfate
[90] groups resulted in a slight decrease in antiproliferative activity. Over-sulfation
(resulting typically in 3,6-GlcN-sulfate and 2,3-IdoA-sulfate), can restore function to less-active (due to small size, N-acetylation, or other chemical modification) heparins,
33
B. General Structure A. Constituent Sugars (Heparin and Heparan sulfate)
- O O O COO H2C OR HC HC HC O α/β O O HC OH HO CH HC NH2 OH OR α O HO CH HC OH HO CH OR NHR'
HC OH HO CH HC OH L-iduronic acid D-glucosamine CH OH HC OH HC OH (D-glucuronic acid)
C C H2C OH HO O HO O C. Most Common Repeat COO- H2C OH O O O (Heparin) COO- OH OH OH - HO OH HO OH HO OH H2C OSO3 O α O OH OH NH COO- 2 O OH OH α D-glucuronic acid L-iduronic acid D-glucosamine O - - OSO3 NHSO3
L-iduronate-2-sulfate N-sulfo-D-glucosamine-6-sulfate
Figure 2.3-Chemical composition and structure of heparin showing A) the Fischer projections of the most common constituent sugar derivatives, B) the general structure of heparin molecules, and C) the most common heparin repeat structure. Commonly sulfated groups are shown in blue. Rarely sulfated groups are shown in red. The orientation of the glycosidic bond shown as α/β in the general structure depends on the stereochemistry of C-5 in the acidic sugar derivatives. “R” groups in general structure: R=H or SO3-, R’ =SO3- or COCH3.
34 while it has limited impact on the activity of full-size native heparin [86, 92]. Over-
sulfation can confer antiproliferative activity to non-heparin glycosaminoglycans such as
hyaluronic acid, dermatan sulfate, and chondroitin sulfate [93]. These results suggest that
the overall level of sulfation (beyond some critical level, as found in unmodified heparin preparations) is the most important determinant of antiproliferative activity, and that there is not one critical sulfation site or structural motif that mediates the antiproliferative effect.
One of the chief inducers of SMC growth is bFGF. This growth factor can stimulate cell growth directly or can stimulate growth in an autocrine fashion after release secondary to stimulation by another factor such as PDGF, thrombin, or factor Xa [81, 94,
95]. It is well known that heparan sulfate proteoglycans (HSPGs) on the cell surface act as low affinity receptors for bFGF and are involved with proper presentation to and full activation of the high affinity FGF receptor [96-98]. Furthermore, bFGF autocrine signaling following thrombin stimulation of SMCs requires syndecan-4, one of the proteoglycan cell surface receptors [95], and bFGF released after PDGF, thrombin, or factor Xa stimulation remains on the SMC’s surface and can be detached by a soluble heparin wash [81, 94] These results suggest that HSPGs play a pivotal role capturing and presenting released bFGF to its receptor to enhance stimulation. A similar mechanism is at employed in rat SMCs when PDGF stimulates the matrix metalloproteinase (MMP) mediated release of heparin binding EGF which augments the proliferative signal of PDGF [99]. This process too can be inhibited by heparin.
However, the relatively obvious hypothesis that heparin disrupts bFGF signaling cannot
explain the range of observations concerning the role of heparin in inhibiting many
35 stimuli. For example, when soluble bFGF is presented to the cells, heparin can both
potentiate [80, 100, 101] and inhibit [84, 96] its signaling. It is known that small amounts
of heparin can facilitate bFGF binding while larger amounts inhibit it [102]. However, it
is unclear whether this effect explains these results since heparin in these studies was
used in relatively high concentration. Furthermore, the response of PDGF to heparin has
not been consistent. In some studies heparin inhibits PDGF stimulated proliferation [78,
94, 103] while in others it has no effect [80, 101] which could be related to species differences (human and rat inhibited, baboon not inhibited) or the presence of small amounts of serum during stimulation (which seems to be correlated with the ability of
heparin to inhibit PDGF-stimulated proliferation). However, it should be noted that in many studies heparin consistently inhibits serum stimulated SMC proliferation [80, 100,
101, 104, 105].
The inconsistencies in these experiments have prompted the exploration of other mechanisms of heparin regulation of SMC growth. Heparin can be internalized [106] via cell-surface heparin sulfate proteoglycans [107] and can activate the double stranded
RNA protein kinase, PKR, which blocks the G1-S transition [104]. Other possible
mechanisms have been proposed including direct signaling through an unspecified
surface receptor, activation of protein phosphatases, and modulation of cell cycle
progression machinery [103, 105, 108]. It is likely that heparin utilizes more than one of
these proposed pathways to modulate cell phenotype.
The well established antiproliferative effect of heparin does not necessarily imply
that heparin can promote a contractile SMC phenotype. The ability of heparin to promote
the expression of contractile phenotype markers, in addition to inhibiting SMC
36 proliferation, has been studied much less extensively. A limited number of studies have
shown that heparin can induce expression of SMαA [109-111] and other smooth muscle
contractile markers [109]. Heparin has also been shown to delay the loss of smoothelin
expression in cultured SMCs [60]. However, the role that heparin’s antiproliferative
signal transduction pathways play in contractile gene expression remains unclear.
Likewise, the role that heparin structure and sulfation patterns play in mediating marker
expression has not been studied. Further exploration of these areas will better establish
the utility of heparin as a mediator of contractile re-differentiation from cultured or
synthetic SMCs.
2.4.1.2. Transforming Growth Factor β1
In contrast to heparin, TGF-β1 has well described ability to both inhibit
proliferation and induce the expression of contractile SMC marker genes in the absence of
stimuli. Active TGF-β1 is a 25 kD homodimer of two 112 amino acid polypeptide chains
which are cleaved from longer propeptides [112]. In cultured vascular SMCs, TGF-β1 inhibits growth induced by serum, PDGF and EGF [113-115], although there is some evidence (from studies done with chick SMCs) that the specific response to TGF-β1 may depend upon the origin of the SMCs, with cells from the aortic arch proliferating in response to TGF-β1 while cells from the abdominal aorta may be growth inhibited [116].
TGF-β1 has also been shown to enhance the expression and organization of SMαA and other more rigorous differentiation markers such as SM-MHC and SM22α in SMC lines as well as primary rat and human SMC cultures [113, 115, 117-119]. Furthermore, in rat and mouse animal models, the level of TGF-β1 in the neointima and damaged media of injured
37 vessels is decreased and correlated with a decrease in SMαA , type IV collagen, and SM-
MHC [120].
Classically, TGF-β1 signals via the Smad family of signaling molecules [121].
Smad-2 nuclear translocation has been correlated with the growth inhibition and SMαA
expression in ocular microvascular pericytes [119] and Smad-3 has been associated with increased contractile marker gene expression via interaction with δEF-1[122]. Once in the
nucleus Smad-2 likely initiates transcription of other transcription factors that ultimately
regulate gene expression. This observation is consistent with the ability of cycloheximide
(a protein synthesis inhibitor) to inhibit transcription of SMC markers, although it does not
rule out the possibility that other signaling pathways also contribute to transcription factor
synthesis [118]. Indeed, other signaling pathways have been implicated in TGF-β1
mediated stimulation of contractile SMC phenotype involving the intracellular Src tyrosine
kinases or RhoA tyrosine kinases and PKN [115, 118]. The role of MAP kinases in this
pathway is not clear since one of these studies demonstrates a role for p38 MAPK while the
other suggests it is not involved. However, the MAP kinase ERK, which is stimulated by
many of the growth stimulating extracellular signals, can play a role in blocking the effects
of TGF-β1 [118]. It should also be noted that while TGF-β1 has been shown to reduce
proliferation and induce contractile SMC marker gene expression, it is unclear whether
TGF-β1 stimulation is sufficient to restore ligand induced contractility to cultured SMCs.
2.4.1.3. Angiotensin II
Angiotensin II induces expression of SMαA in cultured rat aortic SMC via the
angiotensin II type 2 receptor at least in part via increased expression of myocardin (see below) [123]. This result suggests that angiotensin II would also up-regulate the
38 expression of other myocardin dependent marker genes such as SM-MHC, which
empirically had been observed before the discovery of myocardin [124]. Angiotensin II
can also stimulate SMC proliferation [125, 126], although this has not been a consistent
finding [127].
2.4.2. The Role of the Extracellular Matrix
The effects of the ECM on vascular SMCs and their closely related visceral and
pulmonary cousins have been well studied. It has been known for some time that SMCs
rapidly loose their contractile apparatus and adopt a synthetic phenotype in culture [10].
Early reports demonstrated fibronectin (FN), derived from the serum usually used to coat
the substrates for these cultures, most potently supported a loss of contractile phenotype
[11]. Normally SMCs are surrounded by a basal lamina composed predominantly of type
IV collagen and laminin (LN). It appears that this basal lamina is critical for the
maintenance of contractile smooth muscle phenotype, perhaps in part because it forms the interface between the SMC’s contractile apparatus and the ECM [128]. It was later discovered that, in contrast to FN, the basement membrane proteins LN and/or type IV collagen could delay but not eliminate the transition to the synthetic phenotype, even when cells were cultured under serum-free conditions [15, 69, 129-131]. SMCs seeded on these substrates rapidly began to produce their own provisional FN matrix, which becomes the dominant cell-ECM interaction and is correlated with an eventual phenotypic shift [15, 132, 133].
RGD-peptide dependent interactions are critical for this transition. Soluble RGD peptide can delay the transition to the synthetic phenotype on FN [130, 131, 133] or enhance LN’s mitigating effects on SMC response to mitogens like PDGF [11].
39 Furthermore, a substrate of RGD peptide alone was sufficient to induce SMC
dedifferentiation [133].
Some evidence also exists suggesting that LN can promote the expression of
contractile markers in cultured vascular SMCs, as well as mitigate their response to mitogens [132, 134, 135]. LN can also attenuate the response to PDGF and thrombin in
airway SMCs [136]. Study of airway SMCs also has suggested that re-differentiation of
cultured SMCs may involve the production of endogenous basement membrane LN, although the predominant laminin produced, α2β1γ1, is of a different form from LN-1,
which is typically been used to promote contractile phenotype in other reports [137].
Interestingly, the non-basement membrane ECM protein elastin also can promote
contractile phenotype [131, 138]. Studies in tropoelastin null mice suggest this
modulation may occur via a G-protein coupled receptor (GPCR) and can be mimicked by
the short elastin peptide VGVAPG. However, in this study contractile phenotype was
defined by the presence of focal adhesions and stress fibers, which, as evidenced by their
prominent formation on FN, are not robust indicators of phenotype [138, 139]. The
structure of the matrix itself may also play a role. Fibrillar but not degraded collagen can
attenuate vascular SMC response to PDGF [13].
The signaling pathways involved with ECM-dependent modulation of phenotype
have been explored to a limited degree. Inhibition of tyrosine kinases by genistein
resulted in decreased cell spreading and an attenuated progression to the synthetic
phenotype of primary rat SMCs, with the implication that decreased activity of focal
adhesion kinase, which was attenuated by genistein, was involved in this effect [140].
However, this study did not explore alternate mechanisms for this effect, which is
40 problematic since many signaling pathways that promote synthetic SMC phenotype
involve tyrosine kinases. Furthermore, the loss of α7β1 integrin, which is a LN receptor
and part of the linkage between the contractile apparatus and the basement membrane
[128], resulted in decreased expression of contractile SMC markers and increased
proliferation via a Ras-MAPK mediated signaling pathway [141], suggesting that ECM-
α7β1 interactions may normally check this proliferation inducing signaling pathway [142].
Other work has investigated the role of insulin-like growth factor 1 (IGF-1). In primary culture of gizzard SMCs plated on LN, the maintenance of ligand-induced contractility was maintained by endogenous expression of IGF-1 which signaled to the cells in an autocrine/paracrine fashion via a signal cascade involving PI3 kinase and PKB [69]. In
subsequent work, a similar effect was seen in rat SMCs which was consistent with the
signal transduction paradigm developed for gizzard SMCs, although the release and
autocrine signaling of IGF was not specifically tested with these cells [14]. Furthermore,
it appears that this signaling pathway is available only to freshly isolated cells, since sub-
cultured rat SMCs are stimulated to proliferate by IGF-1 [135] at least partly due to
altered activity of SHP-2, a phosphatase which helps to inhibit Ras mediated ERK and
p38 MAPK stimulated cell proliferation in freshly isolated cells by removing a key
phosphotyrosine docking site that adaptor proteins require for Ras activation [82].
2.4.3. The Role of Mechanical Stimulation
The vascular media is constantly subjected to cyclic mechanical loading in vivo,
both in the embryo and in the adult. As a result, it has been widely assumed that the
mechanical environment plays an important role in determining SMC phenotype.
Although the effects of cyclic mechanical strain have been studied extensively, the
41 precise mechanisms that dictate the effects of cyclic strain on SMC phenotype still is
understood poorly with many conflicting reports [143]. There are several important SMC
behaviors that appear to be regulated, at least in part, by cyclic mechanical loading: cell
morphology and alignment, proliferation, ECM elaboration, and expression of contractile
marker proteins.
Most of the studies of cyclic mechanical strain have utilized the Flexcell
apparatus which uses a flexible silicone membrane on the bottom of a 6-well culture plate
that is distended by vacuum from below (for example, see ref. [144]) yielding a non
uniform strain field that is generally oriented radially with larger magnitudes near the
edges and limited strain in the center. A number of studies have employed alternative
geometries, such as tubes [145-147] or uniaxially strained sheets [148, 149], or an
improved version of the Flexcell apparatus that provides equibiaxial strain [150].
Cellular organization obviously is required for contractile smooth muscle tissue to
produce a unified mechanical response. In vitro, SMCs tend to align perpendicular to the direction of strain [148, 151-156]. After the cessation of strain, SMCs tend to lose this
orientation [153, 155]. It is unclear if this alignment is of functional significance or the result of positive selection of cells with an orientation that results in the smallest change in cytoplasmic dimensions. This orientation would also suggest that the contractile apparatus of the cells is disengaged from sensing or modulating this strain since stress fibers typically orient along the long axis of cultured SMCs (Chapter 5, [157]). SMCs
typically align in a circumferential manner in muscular arteries [158] and in tubular
vascular constructs [146] such that cells would be aligned essentially parallel to
circumferential strain.
42 The literature is conflicted regarding the SMC proliferative response to cyclic strain. Early studies with embryonic rat aortic SMCs demonstrated increased cell proliferation in response to cyclic strain that was in part due to release of paracrine/autocrine PDGF [144, 159]. Subsequent studies by other groups have confirmed this response in neonatal rat cells [150, 160] but have shown increased [161], decreased [150, 162], or unchanged [154, 160] proliferation with adult rat aortic cells or
cell lines. SMCs derived from other species also present conflicting results. Cyclic
mechanical strain increased proliferation in rabbit and bovine SMCs [152, 163] but had
no effect on the growth of human or canine SMCs [164, 165].
Some studies have suggested that a laminin or elastin ECM attenuates the ability
of SMCs to proliferate in response to cyclic strain [159, 165], but this was not a
consistent finding [150, 160]. It has been suggested that FN-cell interactions are
important for transducing strain into proliferation, since the RGD peptide or soluble
fibronectin can inhibit neonatal rat SMC proliferation in response to strain [159]. It is
interesting that the effects of the ECM modulated response to cyclic strain follow a
similar pattern to the effects of ECM on SMC phenotype in static culture, that is laminin
and elastin prevent de-differentiation while FN promotes the proliferative, synthetic
phenotype [15, 69, 129-131].
SMCs, typically of rat or rabbit origin, cultured in three dimensional scaffold
systems seem to proliferate slightly in response to mechanical stimulation [146-148].
However, the culture duration in these systems is typically longer than Flexcell-based
experiments and unstrained samples tend to lose cell population, suggesting the enhanced
growth might not be due to strain per se but due to enhanced transport of nutrients into
43 the scaffolds which simply enhances cell viability. The effect of cyclic strain on
intrascaffold transport has not been well characterized for smooth muscle tissue
engineered constructs and may be an important element underlying increased cell
proliferation in these systems.
Cyclic mechanical strain also increased the production of extracellular matrix
components such as collagen and elastin in tissue culture models [164, 166]. There is some evidence to suggest that this response is mediated by paracrine release of TGF-β1,
which is known to directly stimulate collagen production [167]. This effect has led to the
implementation of cyclic loading to improve the mechanical properties of engineered
vascular tissues. Cyclic mechanical strain increased the collagen and elastin content,
organization, and overall strength of 3-dimensional smooth muscle tissues [146-148].
Many studies have used these results as a rationale for mechanical stimulation of tissue
engineered blood vessels [5, 147, 168, 169]
In apparent conflict with the synthetic phenotypic response outlined above, cyclic
strain can also increase the expression of markers of contractile phenotype including:
SMαA [154, 160], calponin [154], SM-22α [154], h-caldesmon [163], and SM-MHC
isoforms [30, 154]. This change in marker expression appeared to be related to
intracellular signaling or short-lived paracrine signaling, since conditioned medium did
not induce contractile marker expression [154]. However, these results were not
consistent in all studies. Some reports indicate that strain has no effect on the expression
of marker proteins [146, 170]. It has also been reported that cyclic strain applied to a 3-D
collagen sponge scaffold, prevented the shifting of SMCs toward an osteogenic
44 phenotype [149]. These studies were performed for long culture durations and the effects
of medium transport due to strain were not discussed.
Given the inconsistencies in the phenotypic response of SMCs to cyclic strain, it
is not surprising that the signaling mechanisms underlying these responses are not well
understood. Immediately following the initiation of cyclic strain all three classical MAP
kinase systems (ERK1/2, JNK, and p38) are activated in a transient fashion with a peak response about 10-15 min after initiation and a return to baseline after 30-60 min [151,
152, 154, 170]. In particular, the p38 pathway is an important component of the system.
Inhibition of p38 activation prevents strain induced alignment [151, 152, 154] and SMαA promoter activity [160]. While p38 may be necessary for these responses, it is not clear that strain directly signals through this pathway or if this is simply a more globally required element of the system, especially since blocking p38 tends to reduce marker gene transcript even in the absence of strain [154]. Putative roles for calcium channels and tyrosine kinases have also been proposed [161, 166]. It is also clear that paracrine release of soluble mediators including angiotensin II, PDGF, TGF-β1, and IGF-1 play an important role and may antagonize each other’s effects [144, 153, 161, 162, 166, 171]. It has been suggested that phenotypic outcome in response to cyclic strain may depend on the phenotype of the cells before strain [145] or the magnitude and duration of the strain
[143]. Future studies will provide a more complete understanding of the signaling processes involved in mechanical stimulation of vascular SMCs.
Clearly, the SMC response to cyclic mechanical strain depends on the state of the cells (both origin and phenotype) [145], and additional work is needed to better understand the conditions that regulate this response. While it is clear that mechanical
45 input plays an important role in the phenotypic modulation of SMCs, a lack of knowledge
regarding the mechanism by which cyclic strain exerts its effects on cells limits its utility
for tissue engineering. Pathways that regulate conflicting phenotypic outcomes such as proliferation and ECM production (synthetic properties) and expression of contractile markers must be more clearly defined so the tissue engineer can specifically target the appropriate cell behavior.
2.4.4. The Role of Endothelium
It has long been recognized that endothelial cells (ECs) play an important role
guiding SMC behavior [172]. Small molecules released from ECs in vivo such as nitric
oxide [173] and endothelin-1 [174] have been shown in vitro to inhibit or stimulate SMC
growth, respectively. SMCs are also known to send projections toward the endothelium
[175]. A variety of co-culture systems have been utilized to explore these interactions
including direct co-culture [176], transmembrane culture [177-179], and bioreactor systems [169, 180]. ECs in these studies tend to increase SMC proliferation [175, 179,
180], suggesting that EC presence promotes synthetic vascular SMC phenotype.
However, many of these studies are performed under static culture conditions, which likely alters the response of the ECs compared with ECs under shear, although increased
SMC proliferation has also been observed in a 3D tissue construct with monolayer ECs cultured under shear [180]. It is also important to note that hyperplastic smooth muscle
lesions have been noted in the “floor” region of the distal anastomoses of vascular
reconstructions, a region that contains native endothelial and smooth muscle tissue but
where there is abnormal (zero) shear stress [181]. SMCs in the presence of ECs also tend
to up-regulate expression of the mitogen PDGF [176] and the inflammatory cytokines
46 interleukin-8 (IL-8) and monocyte chemotactic protein-1 (MCP-1) [177], even as other
proteins associated with the synthetic SMC phenotype are down-regulated, such as
collagen [180] and bFGF [176]. Furthermore, EC-SMC interactions may depend on the
SMC phenotype. The synthetic SMC phenotypic state during co-culture has been shown
to increase the expression of inflammatory signals in both SMCs and ECs [177, 178].
Early studies indicated that non-sheared ECs can delay the contractile-to-synthetic
transition of primary SMCs [9]. While ECs clearly play a critical role in modulating
SMC behavior in vivo, this regulation process is complex. Neither the simple presence or
absence of ECs will result in appropriate SMC behavior, but the appropriate environment
must be provided to both cell types to achieve control of SMC phenotype.
2.4.5. The Role of Inflammation
Inflammation also contributes to a loss of contractile SMC phenotype. Cytokines released from inflammatory cells can directly stimulate SMC growth and play an important role in the development of IH [182, 183]. Several days after endothelial denudation injury macrophages appear in the resulting lesion of proliferating, synthetic
SMCs [184, 185]. Disrupting the accumulation of these cells (using an anti-CD4 antibody) resulted in decreased SMC hyperplasia in vivo, suggesting these cells play an important role in the process [183]. Furthermore, many of the factors these cells generate have been linked directly with SMC proliferation including interleukin-1α [186], IL-8
[187], C-reactive protein [188], and tumor necrosis factor-α [189]. Various stimuli from
activated endothelium and inflammatory cells can also induce the endogenous production
of inflammatory cytokines and markers in SMCs, such as MCP-1 [177], IL-8 [177],
vascular cell adhesion molecule-1 (VCAM-1) [185], intercellular adhesion molecule-1
47 (ICAM-1) [184], class II major histocompatibility complex (MHC-II) [184]. Expression
of these molecules by de-differentiated SMCs provides a mechanism of positive
feedback, accelerating SMC proliferation. These inflammatory processes likely also result in down-regulation of contractile marker proteins, which generally has been correlated SMC proliferation, although this effect has not been studied extensively.
2.5. Molecular Regulation of SMC Gene Expression
In order to affect long-term changes in SMC phenotype, the above mediators ultimately alter gene expression profiles. The current paradigm of phenotypic switching at the genetic level is broadly akin to a switch that toggles between expression of contractile SMC markers and a more generic de-differentiated transcriptional profile without contractile marker expression [6]. In the context of this “switch” the expression of individual marker genes is further modulated by a variety of co-factors at the level of transcription. This section will briefly review these mechanisms of phenotypic regulation in SMCs. A more complete discussion can be found in excellent reviews by Miano et al.
[76] and Kawai-Kowase and Owens [6].
2.5.1. Serum Response Factor
Underlying this broad molecular switch is the 62-67 kD transcription factor, serum response factor (SRF) [190]. SRF was initially identified as a transcription factor that acts as a promoter for c-fos, a gene involved in the early stages of cell proliferation
[191, 192]. SRF is activated by transcription following serum stimulation and does not require additional protein translation to exert its effects [193]. However, it was quickly realized that SRF also was active in the promoter of sarcomeric muscle specific α-actin
(not to be confused with smooth muscle specific α-actin discussed throughout the rest of
48 this work) [194]. This began over two decades of work to describe how SRF
differentially regulates a wide range of genes for seemingly disparate cellular programs.
SRF binds as a homodimer to a consensus sequence in DNA of CC(A or T)6GG,
called a CArG box [76, 193]. Putative CArG elements have been identified in the
promoter/enhancer regions of nearly 200 genes [195], and many of these genes are
involved in formation and regulation of the cytoskeleton or contractile apparatus [76].
Most, but not all, markers of contractile SMC phenotype contain a least one CArG box
including SMαA, calponin, SM-22α, and SM-MHC [76]. The ability for SRF to activate
specific transcriptional programs within the wide range of genes containing CArG boxes
depends upon the presence of program specific co-activators and repressors. Specific
expression of many smooth muscle specific genes is substantially enhanced by the co-
activator myocardin.
2.5.2. Myocardin
Myocardin is a central component of the SMC phenotype master switch and can
drive expression for most, but not all, contractile marker genes [72]. Myocardin is a 96
kD transcription factor that directly interacts with SRF dimers via a domain in its N-
terminus rather than binding directly to DNA [196]. Myocardin appears to be regulated,
at least in part, at the transcriptional level and is generally restricted to the nucleus [72,
197]. Myocardin also contains a leucine zipper domain, which allows myocardin dimers
to bridge adjacent CArG boxes in the promoter region of many SMC marker genes including SMαA, calponin, SM-22α, and SM-MHC [74, 190]. This six member
myocardin2-SRF4 dimer complex seems to enhance activation of these genes [74]. It has
also been suggested that myocardin’s interaction with SRF enhances SRF’s binding to
49 degenerate CArG boxes in the promoters of some SMC marker genes [198] such as SM-
MHC, SMαA, and calponin, which typically contain one guanine substitution in the A/T rich part of the CArG sequence [190].
Myocardin co-activation alone, however does not fully explain the transcriptional control of SMC marker genes for several reasons. Over-expression of myocardin in rat aortic SMCs, mesenchymal stem cells, and fibroblast results in inappropriate activation of skeletal and cardiac muscle genes and fails to activate non-CArG containing markers such as smoothelin-B [72, 199]. Dominant negative myocardin fails to interrupt marker expression in a SMC differentiation model cell line A404 [72, 200]. Furthermore, ACLP, which has SMC restricted expression, does not contain a CArG box and does not depend on SRF or myocardin for transcription [71]. Even among genes with known CArG containing promoters, some possess only a single CArG box such as h-caldesmon and telokin suggesting that the myocardin dimerization hypothesis [74] has limitations.
Furthermore, SMC marker genes are not expressed uniformly during development nor lost uniformly during pathogenesis [7] implying, not surprisingly, that additional layers of control are involved in marker gene expression. Most strikingly, although myocardin null mice die in utero [201], myocardin-null mouse embryonic stem cells can express some SMC markers in vitro and the vasculature of embryos, formed from a chimera of wild type and myocardin-null cells, contain myocardin-null SMCs expressing a normal complement of SMC markers [202].
2.5.3. Other Co-activation Systems
Other co-activation schemes can function in conjunction with the myocardin-SRF system to drive marker expression. The SMαA promoter has been studied most
50 thoroughly and, in addition to three CArG boxes, contains two E-boxes (CANNTG
motifs), two MCAT elements (AGGAATG motifs) and a TGF-β control element (Fig.
2.4) [203]. The E-boxes are bound by basic helix-loop-helix transcription factor dimers
that enhance SRF-dependent transcription via the protein inhibitor of activated STAT
(PIAS)-1 [204]. The two MCAT boxes may be involved in parallel SMαA regulatory
pathways in myofibroblasts which are not critical for SMCs [24]. The homeobox binding protein Prx-1 also has been shown to be important in SMαA transcription [123].
Furthermore, myocardin related transcription factors (MRTFs) may also play a role in modulating gene expression in a fashion similar to myocardin via sensing changes in the
actin cytoskeleton [76, 205].
2.5.4. Repressor Systems
Several repressor pathways are known to affect the regulation of many of the
CArG-box-regulated SMC marker genes. Two examples include Elk-1 and Kruppel-like
factor 4 (KLF-4). Elk-1 is a transcriptional cofactor that, like myocardin, interacts with
SRF to modulate transcription. Elk-1 is activated by phosphorylation by ERK1/2 and
JNK MAP kinases, depending on the stimulus, and increases SRF dependent
transcription of c-fos [206]. Thus Elk-1 promotes cell proliferation [206]. Elk-1 can
interact with SRF and DNA in the promoter region of several SMC marker genes to
inhibit myocardin binding and marker gene activation [75, 207, 208], and this activity
seems to vary between genes [207]. KLF-4 inhibits marker transcription, perhaps via
multiple pathways. KLF-4 binds in the promoter region of several SMC marker genes
[209, 210], which may allow it to block SRF binding to SMC promoters [211] and recruit
histone deacetylases that alter chromatin structure to limit transcription factor access to
51 the promoter regions [208]. KLF4 can also suppress myocardin [211]. In addition to
enhancing SMαA transcription via interactions with E-box binding proteins, PIAS1 may
promote gene expression by inhibiting the action of KLF-4 [212]. In addition to these
modifiers, additional inhibitory pathways have been identified. HERP1, which is up-
regulated in cultured SMCs, binds to SRF to inhibit its binding to CArG boxes [213].
Ets-1, which is related to Elk-1, is up-regulated in vascular injury and suppresses marker gene expression [214]. An overview of both transcriptional activation and repression mechanisms, using SMαA transcription as an example, can be found in Fig. 2.4.
2.5.5. Discussion
Substantial progress has been made in recent years in our understanding of the
molecular regulation of SMC phenotype. However, it is clear that additional work is
needed to more fully understand the convergence and significance of each regulatory
pathway in the overall differentiation process. For example, how are non-CArG containing genes regulated? Which pathways are most important for re-differentiation of synthetic SMCs? How does one engage the multitude of regulatory systems employed for each gene to re-establish a functional contractile apparatus?
In the context of tissue engineering, understanding the molecular regulation of
SMC phenotype is critical to effectively generate appropriately differentiated smooth
muscle tissues. Such information allows the engineer to peer within the black box of the
cell to better understand specifically how a bioactive scaffold system is affecting cell
behavior. For example, the role of MCAT elements in biomaterial-delivered, TGF-β1-
stimulated modulation of SMC phenotype [146, 167] may help to elucidate if these cells
52
RNA PIAS1 Myocardin Myocardin Myocardin A Pol II Complex SRF SRF SRF SRF SRF SRF bHLH bHLH bHLH bHLH MCAT E-box E-box MCAT CArG CArG TCE TATA α-Actin CArG -320 -252-214 -184 -120 -70 -56 +1098
P HERP1 SRF SRF Elk-1 SRF B SRF SRF SRF KLF-4 MCAT E-box E-box MCAT CArG CArG TCE TATA α-Actin CArG -320 -252-214 -184 -120 -70 -56 +1098
Histone Deacetylases SRF SRF SRF Myocardin Myocardin
Myocardin SRF SRF Myocardin SRF
Figure 2.4-Molecular regulation of smooth muscle α-actin transcription, illustrating example mechanisms of transcription activation in differentiated smooth muscle cells (A) and mechanisms of down-regulation (B). A. Transcription is activated by SRF binding to CArG box up- and down-stream of the TATA box, enhanced by the co-activator, myocardin. Additional elements further enhance transcription, such as bHLH transcription factors via PIAS-1. B. Transcription is down-regulated by phospho-Elk-1 blocking myocardin interactions with SRF. KLF-4 and HERP-1 block SRF binding to CArG boxes via sequestration. KLF-4 also activates histone deacetylases which close chromatin structure, limiting transcription factor access to the promoter region. Abbreviations: bHLH: basic helix-loop-helix transcription factor, PIAS-1: protein inhibitor of activated STAT-1, SRF: serum response factor, KLF4: Kruppel-like factor 4 (KLF4), TCE: TGF-β control element.
53 are differentiating toward a mature SMC phenotype or following a tangential pathway
toward a more myofibroblast-like state. If the ultimate goal is the regeneration of functional contractile smooth muscle, clearly the former would be preferable. In addition, as the molecular regulation of SMC phenotype becomes better characterized, it may provide useful targets for gene therapy that could potentially initiate SMC differentiation. Gene delivery of myocardin may be a good initial target to investigate whether an appropriate scaffold micro-environmental context could increase selective expression of SMC marker genes over skeletal or cardiac muscle genes, the expression of which has been observed in myocardin over-expression models [72]. Furthermore, this information provides a variety of targets for in situ gene knock-down delivery strategies, such as siRNA, that may mitigate SMC hyperplasia in vivo. One obvious target would be
Elk-1, since it plays a role both in stimulating SMC proliferation and inhibiting SMC
marker gene expression. Consideration of the molecular control of SMC gene expression
in the rational design of bioactive scaffold materials may lead to significant
improvements in the ability of these materials to regulate SMC phenotype.
2.6. Engineered Biomaterial Approaches to Regulate SMC Phenotype
Biomaterial and tissue engineers have begun to investigate the effects of scaffold
chemistry and structure on SMC phenotype. These studies have used engineered
materials as model systems to explore factors that affect SMC behavior. Novel materials
and fabrication strategies also have been developed to modulate SMC phenotype. This
section will briefly review this work.
54 2.6.1. Effects of Cell-Scaffold Interactions on Phenotype
Early work in smooth muscle tissue engineering explored the effect simple scaffold materials such as poly(glycolic acid) (PGA), poly(lactic-co-glycolic acid), and collagen gels had on SMC behavior [18]. SMCs in polyester scaffolds showed a higher ratio of elastin:collagen production than collagen gels, suggesting a shift toward a contractile-like phenotype [18]. Although the 3D structure of these matrices was dissimilar, the observed differences likely were due, at least in part, to differences in the scaffold chemistry either by affecting SMCs directly or by inducing differences in the adsorbed protein content on the scaffolds [18]. Despite these promising results, SMCs in
PGA vascular grafts [5] showed incomplete differentiation and contractility in vivo, and later study suggested that breakdown products from PGA may promote synthetic SMC phenotype [215].
In an effort to more precisely understand the role specific cell-ECM interactions play in SMC phenotype modulation, cell-adhesive peptides and sugars have been immobilized on various surfaces and their effect on cell behavior studied. For SMCs seeded on modified glass, collagen production decreased with ligand concentration and collagen per cell was highest on weakly adhesive substrates (such as RGE modified glass), though these results were confounded by high non-specific cell binding to the substrates [216]. However, similar results were obtained for SMCs seeded on poly(ethylene glycol) (PEG)-based hydrogels [217], which resist non-specific attachment
(Chapter 5, [157]). The use of tethered TGF-β1 also increased the synthesis of extracellular matrix components on RGD-containing PEG-based gel systems [167].
Short hyaluronic acid fragments tethered to glass induced increased elastin production
55 and crosslinking [218]. However, the biological characterization of SMC phenotype in these studies was limited to indirect markers. SMCs, attached to RGD-bearing PEG- based hydrogels, have been shown to qualitatively express markers of contractile phenotype [219]. This expression may be related to the mechanical characteristics of the underlying substrate [219]. Recently, it has been shown that RGD-bearing hydrogels with highly specific cell-matrix interactions, can support a robust, quantitative re- expression of contractile marker mRNA and proteins, given appropriate culture conditions (Chapter 5, [157]).
Many groups have noted that SMC behavior differs between 2D and 3D systems.
Culture in 3D collagen gels resulted in decreased proliferation and increased collagen synthesis compared with 2D collagen cultures [220]. Furthermore, inhibition of ERK activation using PD98059 induced cell proliferation for SMCS in 3D collagen matrices whereas it inhibited growth in 2D cultures on the same substrate, suggesting culture geometry plays an important role modulating this signal [221]. Synthetic scaffolds must be biodegradable and/or highly porous to permit 3D SMC culture. To allow for degradation in synthetic gel scaffold systems, the gel network must incorporate a degradable component. SMCs cultured in 3D PEG-based gel systems with an elastase degradable sequence showed higher collagen production than SMCs in non-degradable controls [222]. Culture in 3D, PEG-based hydrogel scaffolds resulted in small but significant increases in the smooth muscle markers SMαA and SM-MHC compared with control SMCs cultured on tissue culture plastic, although it was difficult to discern from this study the contributions of the scaffold geometry from substrate chemistry or mechanical properties (gel versus tissue culture polystyrene) [223]. Increased
56 mechanical modulus in 3D fibrinogen/PEG-based hydrogel scaffolds was also correlated with increased expression of vinculin, a marker of focal adhesions, and SMC differentiation markers, but only for SMCs overexpressing RhoA, which is a signaling protein that plays an important role in focal adhesion formation [224].
The interplay between the scaffold chemistry, mechanical stimulation, and external biochemical stimulation has also been explored. SMCs on fibronectin coated
PGA-based scaffolds increase elastin production in response to mechanical stimulation more than SMCs in collagen gels [148]. Mechanical stimulation of rat aortic SMCs when seeded in a collagen-based tubular graft-like construct resulted in increased compaction of the collagen which could be further enhanced by stimulation with TGF-β1, which also improved histological organization and increased SMαA expression [146]. Interestingly,
SMCs transfected with cyclic GMP dependent protein kinase, which promotes contractile
SMC phenotype [16, 225], did not affect histological organization in collagen gels although SMαA expression was dramatically increased, especially with exogenous TGF-
β1 [226]. These results suggest that forced expression of contractile SMC markers per se may not substantially facilitate the organization of vascular tissues in vitro. Recently,
RGD-bearing PEG diacrylate (PEGDA) hydrogels were utilized as a 3D scaffold material to examine the effects of both SMC co-culture with ECs and cyclic mechanical loading
[169]. When subjected to both conditions, SMCs showed a modest up-regulation of elastin, calponin, and myocardin as well as a slight decrease in collagen production [169].
However, this study was limited by the use of non-degradable PEGDA, which does not permit normal SMC morphology post-encapsulation, and the non-physiologic 3D encapsulation of the ECs (as opposed to monolayer culture) [169].
57 2.6.2. Micropatterning
Rat A7r5 cells organized into aligned patterns when seeded in 160 μm wide
channels of photopolymerized poly(caprolactone-lactide-glycolide) diacrylate
microchannels with some evidence of increased SMαA production in these cells [227].
We have observed similar banding patterns in our cultures (Chapter 5, [157]). However, the mechanism by which the channels induce alignment is unclear. Since cultured SMCs have a propensity toward alignment, the uniform cell orientation in this system may be due simply to preclusion of alignment perpendicular to the channels (because they are narrower than the length of an SMC). It is also unclear if this alignment directly affects
SMC phenotype, although there some limited data to suggest SMCs in these microchannels may up-regulate SMαA [227].
2.6.3. Conducting polymers
Cyclic electrical stimulation of vascular SMCs using an ECM coated, conducting
polymeric (polypyrrole/hyaluronic acid) scaffold stimulated SMC proliferation and
concomitantly stimulated up-regulation of SMαA and SM-MHC [228]. Blockade of L-
type calcium channels abrogated this effect [228], consistent with others that have shown
that calcium influx through L-type calcium channels (induced by depolarization with
KCl) can result in increased expression of CArG dependent marker genes [229]
2.7. Conclusions
The phenotypic plasticity of SMCs confers these cells with great regenerative potential. However, to engineer functional smooth muscle tissues while minimizing development of hyperplastic pathologies such as IH, SMC phenotype must be well
controlled. In the last several decades, a better understanding of the myriad factors that
58 regulate SMC phenotype has emerged. The intracellular signaling machinery that relays
these signals and the transcriptional machinery that ultimately results in phenotype changes are also becoming clearer. A panel of markers that can be utilized to quantitatively characterize SMC phenotype has been well established. However, the best approaches to modulate synthetic, cultured SMCs toward a functional contractile phenotype remain largely unknown, though the fact that SMCs seem to re-differentiate in vivo [65] suggests that given the right conditions, this goal is attainable.
Despite these advances, the tissue engineering literature for smooth muscle tissues remains largely observational in nature. Few studies (for example, [146, 169]) exploit the well established basic science literature on the topic to generate scaffold systems specifically designed to modulate SMCs toward a contractile phenotype. Mechanical stimulation, which has yielded inconsistent results in the basic science literature, has been
the most commonly used strategy, although this approach does tend to yield tissues that
are more mechanically robust. Critical parameters such as cell origin and cell phenotype during stimulation generally have been ignored. Study of scaffolds that promote contractile phenotype using stimulation with exogenous signaling factors such as heparin and TGF-β1 has been surprisingly limited, given the clear evidence of the efficacy of
these approaches.
Further confounding the interpretation of these studies is a lack of clear,
consistent, and quantitative readouts of SMC phenotype. Most studies have employed
indirect measures such as extracellular matrix synthesis and cell proliferation. Given the
diversity factors that can affect these non-specific cell behaviors, it is challenging to
compare true SMC phenotypes between studies. Studies that have examined SMC
59 phenotype markers directly have used qualitative techniques such as immunostaining,
which, without quantification, also yields results that are difficult to compare between
studies. Expanded use of appropriate (semi)quantitative techniques for assessing markers of contractile SMC phenotype including qPCR, western blot, ELISA, and flow cytometry will aide in the comparison of tissue engineering strategies to promote the formation of contractile smooth muscle tissue. Expanded analysis of the transcriptional regulation of
marker gene expression [169] will lend further insight into mechanisms by which novel
scaffold designs are capable of modulating SMC phenotype. Future work that combines
scaffold materials with well-defined cell-material interactions, soluble signals, and
mechanical stimulation, utilizing quantitative methods will likely succeed in devising
systems that are capable of inducing re-expression of true contractile SMC phenotype.
60 2.8. References
1. Meinhart JG, Deutsch M, Fischlein T, Howanietz N, Froschl A, Zilla P. Clinical autologous in vitro endothelialization of 153 infrainguinal ePTFE grafts. Ann Thorac Surg 2001;71(5 Suppl):S327-331.
2. Laube HR, Duwe J, Rutsch W, Konertz W. Clinical experience with autologous endothelial cell-seeded polytetrafluoroethylene coronary artery bypass grafts. J Thorac Cardiovasc Surg 2000;120(1):134-141.
3. Schneider F, O'Connor S, Becquemin JP. Efficacy of Collagen Silver-Coated Polyester and Rifampin-Soaked Vascular Grafts to Resist Infection from MRSA and Escherichia coli in a Dog Model. Annals of Vascular Surgery 2008;22(6):815- 821.
4. Darouiche RO, Mansouri MD. In vitro activity and in vivo efficacy of antimicrobial-coated vascular grafts. Annals of Vascular Surgery 2004;18(4):497- 501.
5. Niklason LE, Gao J, Abbott WM, Hirschi KK, Houser S, Marini R, Langer R. Functional arteries grown in vitro. Science 1999;284(5413):489-493.
6. Kawai-Kowase K, Owens GK. Multiple repressor pathways contribute to phenotypic switching of vascular smooth muscle cells. Am J Physiol Cell Physiol 2007;292(1):C59-69.
7. Owens GK. Regulation of differentiation of vascular smooth muscle cells. Physiol Rev 1995;75(3):487-517.
8. Stegemann JP, Hong H, Nerem RM. Mechanical, biochemical, and extracellular matrix effects on vascular smooth muscle cell phenotype. J Appl Physiol 2005;98(6):2321-2327.
9. Campbell JH, Campbell GR. Vascular smooth muscle in culture. Boca Raton, Fla.: CRC Press, 1987.
10. Chamley-Campbell J, Campbell GR, Ross R. The smooth muscle cell in culture. Physiol Rev 1979;59(1):1-61.
11. Hedin U, Thyberg J. Plasma fibronectin promotes modulation of arterial smooth- muscle cells from contractile to synthetic phenotype. Differentiation 1987;33(3):239-246.
61 12. Rzucidlo EM, Martin KA, Powell RJ. Regulation of vascular smooth muscle cell differentiation. J Vasc Surg 2007;45 Suppl A:A25-32.
13. Raines EW, Koyama H, Carragher NO. The extracellular matrix dynamically regulates smooth muscle cell responsiveness to PDGF. Ann N Y Acad Sci 2000;902:39-51; discussion 51-32.
14. Hayashi K, Takahashi M, Kimura K, Nishida W, Saga H, Sobue K. Changes in the balance of phosphoinositide 3-kinase/protein kinase B (Akt) and the mitogen- activated protein kinases (ERK/p38MAPK) determine a phenotype of visceral and vascular smooth muscle cells. J Cell Biol 1999;145(4):727-740.
15. Hedin U, Bottger BA, Forsberg E, Johansson S, Thyberg J. Diverse effects of fibronectin and laminin on phenotypic properties of cultured arterial smooth muscle cells. J Cell Biol 1988;107(1):307-319.
16. Boerth NJ, Dey NB, Cornwell TL, Lincoln TM. Cyclic GMP-dependent protein kinase regulates vascular smooth muscle cell phenotype. J Vasc Res 1997;34(4):245-259.
17. Chamley-Campbell JH, Campbell GR, Ross R. Phenotype-dependent response of cultured aortic smooth muscle to serum mitogens. J Cell Biol 1981;89(2):379-383.
18. Kim BS, Nikolovski J, Bonadio J, Smiley E, Mooney DJ. Engineered smooth muscle tissues: regulating cell phenotype with the scaffold. Exp Cell Res 1999;251(2):318-328.
19. Herman IM. Actin isoforms. Curr Opin Cell Biol 1993;5(1):48-55.
20. Khaitlina SY. Functional specificity of actin isoforms. Int Rev Cytol 2001;202:35- 98.
21. Owens GK, Loeb A, Gordon D, Thompson MM. Expression of smooth muscle- specific alpha-isoactin in cultured vascular smooth muscle cells: relationship between growth and cytodifferentiation. J Cell Biol 1986;102(2):343-352.
22. Kim HR, Gallant C, Leavis PC, Gunst SJ, Morgan KG. Cytoskeletal remodeling in differentiated vascular smooth muscle is actin isoform dependent and stimulus dependent. Am J Physiol Cell Physiol 2008;295(3):C768-778.
23. Duband JL, Gimona M, Scatena M, Sartore S, Small JV. Calponin and SM 22 as differentiation markers of smooth muscle: spatiotemporal distribution during avian embryonic development. Differentiation 1993;55(1):1-11.
62 24. Gan Q, Yoshida T, Li J, Owens GK. Smooth muscle cells and myofibroblasts use distinct transcriptional mechanisms for smooth muscle alpha-actin expression. Circ Res 2007;101(9):883-892.
25. Deissler H, Deissler H, Lang GK, Lang GE. TGFbeta induces transdifferentiation of iBREC to alphaSMA-expressing cells. Int J Mol Med 2006;18(4):577-582.
26. Arciniegas E, Sutton AB, Allen TD, Schor AM. Transforming growth factor beta 1 promotes the differentiation of endothelial cells into smooth muscle-like cells in vitro. J Cell Sci 1992;103 ( Pt 2):521-529.
27. Babu GJ, Warshaw DM, Periasamy M. Smooth muscle myosin heavy chain isoforms and their role in muscle physiology. Microsc Res Tech 2000;50(6):532- 540.
28. Frid MG, Printesva OY, Chiavegato A, Faggin E, Scatena M, Koteliansky VE, Pauletto P, Glukhova MA, Sartore S. Myosin heavy-chain isoform composition and distribution in developing and adult human aortic smooth muscle. J Vasc Res 1993;30(5):279-292.
29. Babu GJ, Pyne GJ, Zhou Y, Okwuchukuasanya C, Brayden JE, Osol G, Paul RJ, Low RB, Periasamy M. Isoform switching from SM-B to SM-A myosin results in decreased contractility and altered expression of thin filament regulatory proteins. Am J Physiol Cell Physiol 2004;287(3):C723-729.
30. Reusch P, Wagdy H, Reusch R, Wilson E, Ives HE. Mechanical strain increases smooth muscle and decreases nonmuscle myosin expression in rat vascular smooth muscle cells. Circ Res 1996;79(5):1046-1053.
31. Kuro-o M, Nagai R, Nakahara K, Katoh H, Tsai RC, Tsuchimochi H, Yazaki Y, Ohkubo A, Takaku F. cDNA cloning of a myosin heavy chain isoform in embryonic smooth muscle and its expression during vascular development and in arteriosclerosis. J Biol Chem 1991;266(6):3768-3773.
32. Lauzon AM, Tyska MJ, Rovner AS, Freyzon Y, Warshaw DM, Trybus KM. A 7- amino-acid insert in the heavy chain nucleotide binding loop alters the kinetics of smooth muscle myosin in the laser trap. J Muscle Res Cell Motil 1998;19(8):825- 837.
33. Chi M, Zhou Y, Vedamoorthyrao S, Babu GJ, Periasamy M. Ablation of smooth muscle myosin heavy chain SM2 increases smooth muscle contraction and results in postnatal death in mice. Proc Natl Acad Sci U S A 2008;105(47):18614-18618.
63 34. Babij P, Kawamoto S, White S, Adelstein RS, Periasamy M. Differential expression of SM1 and SM2 myosin isoforms in cultured vascular smooth muscle. Am J Physiol 1992;262(3 Pt 1):C607-613.
35. Miano JM, Cserjesi P, Ligon KL, Periasamy M, Olson EN. Smooth muscle myosin heavy chain exclusively marks the smooth muscle lineage during mouse embryogenesis. Circ Res 1994;75(5):803-812.
36. Winder SJ, Walsh MP. Calponin. Curr Top Cell Regul 1996;34:33-61.
37. Rozenblum GT, Gimona M. Calponins: adaptable modular regulators of the actin cytoskeleton. Int J Biochem Cell Biol 2008;40(10):1990-1995.
38. Morgan KG, Gangopadhyay SS. Invited review: cross-bridge regulation by thin filament-associated proteins. J Appl Physiol 2001;91(2):953-962.
39. Mezgueldi M, Mendre C, Calas B, Kassab R, Fattoum A. Characterization of the regulatory domain of gizzard calponin. Interactions of the 145-163 region with F- actin, calcium-binding proteins, and tropomyosin. J Biol Chem 1995;270(15):8867- 8876.
40. Mino T, Yuasa U, Nakamura F, Naka M, Tanaka T. Two distinct actin-binding sites of smooth muscle calponin. Eur J Biochem 1998;251(1-2):262-268.
41. Burgstaller G, Kranewitter WJ, Gimona M. The molecular basis for the autoregulation of calponin by isoform-specific C-terminal tail sequences. J Cell Sci 2002;115(Pt 10):2021-2029.
42. Winder SJ, Walsh MP. Smooth muscle calponin. Inhibition of actomyosin MgATPase and regulation by phosphorylation. J Biol Chem 1990;265(17):10148- 10155.
43. Leinweber B, Parissenti AM, Gallant C, Gangopadhyay SS, Kirwan-Rhude A, Leavis PC, Morgan KG. Regulation of protein kinase C by the cytoskeletal protein calponin. J Biol Chem 2000;275(51):40329-40336.
44. Leinweber BD, Leavis PC, Grabarek Z, Wang CL, Morgan KG. Extracellular regulated kinase (ERK) interaction with actin and the calponin homology (CH) domain of actin-binding proteins. Biochem J 1999;344 Pt 1:117-123.
45. Yoshikawa H, Taniguchi SI, Yamamura H, Mori S, Sugimoto M, Miyado K, Nakamura K, Nakao K, Katsuki M, Shibata N, Takahashi K. Mice lacking smooth muscle calponin display increased bone formation that is associated with
64 enhancement of bone morphogenetic protein responses. Genes Cells 1998;3(10):685-695.
46. Masuki S, Takeoka M, Taniguchi S, Nose H. Enhanced baroreflex sensitivity in free-moving calponin knockout mice. Am J Physiol Heart Circ Physiol 2003;284(3):H939-946.
47. Masuki S, Takeoka M, Taniguchi S, Yokoyama M, Nose H. Impaired arterial pressure regulation during exercise due to enhanced muscular vasodilatation in calponin knockout mice. J Physiol 2003;553(Pt 1):203-212.
48. Matthew JD, Khromov AS, McDuffie MJ, Somlyo AV, Somlyo AP, Taniguchi S, Takahashi K. Contractile properties and proteins of smooth muscles of a calponin knockout mouse. J Physiol 2000;529 Pt 3:811-824.
49. Miano JM, Olson EN. Expression of the smooth muscle cell calponin gene marks the early cardiac and smooth muscle cell lineages during mouse embryogenesis. J Biol Chem 1996;271(12):7095-7103.
50. Assinder SJ, Stanton JA, Prasad PD. Transgelin: an actin-binding protein and tumour suppressor. Int J Biochem Cell Biol 2009;41(3):482-486.
51. Fu Y, Liu HW, Forsythe SM, Kogut P, McConville JF, Halayko AJ, Camoretti- Mercado B, Solway J. Mutagenesis analysis of human SM22: characterization of actin binding. J Appl Physiol 2000;89(5):1985-1990.
52. Zhang JC, Kim S, Helmke BP, Yu WW, Du KL, Lu MM, Strobeck M, Yu Q, Parmacek MS. Analysis of SM22alpha-deficient mice reveals unanticipated insights into smooth muscle cell differentiation and function. Mol Cell Biol 2001;21(4):1336-1344.
53. Je HD, Sohn UD. SM22alpha is required for agonist-induced regulation of contractility: evidence from SM22alpha knockout mice. Mol Cells 2007;23(2):175- 181.
54. Lepore JJ, Cheng L, Min Lu M, Mericko PA, Morrisey EE, Parmacek MS. High- efficiency somatic mutagenesis in smooth muscle cells and cardiac myocytes in SM22alpha-Cre transgenic mice. Genesis 2005;41(4):179-184.
55. Li L, Miano JM, Cserjesi P, Olson EN. SM22 alpha, a marker of adult smooth muscle, is expressed in multiple myogenic lineages during embryogenesis. Circ Res 1996;78(2):188-195.
65 56. Faggin E, Puato M, Zardo L, Franch R, Millino C, Sarinella F, Pauletto P, Sartore S, Chiavegato A. Smooth muscle-specific SM22 protein is expressed in the adventitial cells of balloon-injured rabbit carotid artery. Arterioscler Thromb Vasc Biol 1999;19(6):1393-1404.
57. van Eys GJ, Niessen PM, Rensen SS. Smoothelin in vascular smooth muscle cells. Trends Cardiovasc Med 2007;17(1):26-30.
58. van der Loop FT, Schaart G, Timmer ED, Ramaekers FC, van Eys GJ. Smoothelin, a novel cytoskeletal protein specific for smooth muscle cells. J Cell Biol 1996;134(2):401-411.
59. Rensen SS, Thijssen VL, De Vries CJ, Doevendans PA, Detera-Wadleigh SD, Van Eys GJ. Expression of the smoothelin gene is mediated by alternative promoters. Cardiovasc Res 2002;55(4):850-863.
60. Christen T, Bochaton-Piallat ML, Neuville P, Rensen S, Redard M, van Eys G, Gabbiani G. Cultured porcine coronary artery smooth muscle cells. A new model with advanced differentiation. Circ Res 1999;85(1):99-107.
61. Niessen P, Clement S, Fontao L, Chaponnier C, Teunissen B, Rensen S, van Eys G, Gabbiani G. Biochemical evidence for interaction between smoothelin and filamentous actin. Exp Cell Res 2004;292(1):170-178.
62. Niessen P, Rensen S, van Deursen J, De Man J, De Laet A, Vanderwinden JM, Wedel T, Baker D, Doevendans P, Hofker M, Gijbels M, van Eys G. Smoothelin-a is essential for functional intestinal smooth muscle contractility in mice. Gastroenterology 2005;129(5):1592-1601.
63. Rensen SS, Niessen PM, van Deursen JM, Janssen BJ, Heijman E, Hermeling E, Meens M, Lie N, Gijbels MJ, Strijkers GJ, Doevendans PA, Hofker MH, De Mey JG, van Eys GJ. Smoothelin-B deficiency results in reduced arterial contractility, hypertension, and cardiac hypertrophy in mice. Circulation 2008;118(8):828-836.
64. Deruiter MC, Rensen SS, Coolen GP, Hierck BP, Bergwerff M, Debie WM, Gittenberger-De Groot AC, Van Eys GJ. Smoothelin expression during chicken embryogenesis: detection of an embryonic isoform. Dev Dyn 2001;221(4):460-463.
65. Christen T, Verin V, Bochaton-Piallat M, Popowski Y, Ramaekers F, Debruyne P, Camenzind E, van Eys G, Gabbiani G. Mechanisms of neointima formation and remodeling in the porcine coronary artery. Circulation 2001;103(6):882-888.
66 66. Baker SC, Southgate J. Towards control of smooth muscle cell differentiation in synthetic 3D scaffolds. Biomaterials 2008;29(23):3357-3366.
67. Desmouliere A, Geinoz A, Gabbiani F, Gabbiani G. Transforming growth factor- beta 1 induces alpha-smooth muscle actin expression in granulation tissue myofibroblasts and in quiescent and growing cultured fibroblasts. J Cell Biol 1993;122(1):103-111.
68. Sobue K, Sellers JR. Caldesmon, a novel regulatory protein in smooth muscle and nonmuscle actomyosin systems. J Biol Chem 1991;266(19):12115-12118.
69. Hayashi K, Saga H, Chimori Y, Kimura K, Yamanaka Y, Sobue K. Differentiated phenotype of smooth muscle cells depends on signaling pathways through insulin- like growth factors and phosphatidylinositol 3-kinase. J Biol Chem 1998;273(44):28860-28867.
70. Layne MD, Endege WO, Jain MK, Yet SF, Hsieh CM, Chin MT, Perrella MA, Blanar MA, Haber E, Lee ME. Aortic carboxypeptidase-like protein, a novel protein with discoidin and carboxypeptidase-like domains, is up-regulated during vascular smooth muscle cell differentiation. J Biol Chem 1998;273(25):15654- 15660.
71. Layne MD, Yet SF, Maemura K, Hsieh CM, Liu X, Ith B, Lee ME, Perrella MA. Characterization of the mouse aortic carboxypeptidase-like protein promoter reveals activity in differentiated and dedifferentiated vascular smooth muscle cells. Circ Res 2002;90(6):728-736.
72. Yoshida T, Kawai-Kowase K, Owens GK. Forced expression of myocardin is not sufficient for induction of smooth muscle differentiation in multipotential embryonic cells. Arterioscler Thromb Vasc Biol 2004;24(9):1596-1601.
73. Yoshida T, Gan Q, Shang Y, Owens GK. Platelet-Derived Growth Factor-BB Represses Smooth Muscle Cell Marker Genes via Changes in Binding of MKL Factors and Histone Deacetylases to their Promoters. Am J Physiol Cell Physiol 2006.
74. Wang Z, Wang DZ, Pipes GC, Olson EN. Myocardin is a master regulator of smooth muscle gene expression. Proc Natl Acad Sci U S A 2003;100(12):7129- 7134.
75. Wang Z, Wang DZ, Hockemeyer D, McAnally J, Nordheim A, Olson EN. Myocardin and ternary complex factors compete for SRF to control smooth muscle gene expression. Nature 2004;428(6979):185-189.
67 76. Miano JM, Long X, Fujiwara K. Serum response factor: master regulator of the actin cytoskeleton and contractile apparatus. Am J Physiol Cell Physiol 2007;292(1):C70-81.
77. Hedin U, Daum G, Clowes AW. Heparin inhibits thrombin-induced mitogen- activated protein kinase signaling in arterial smooth muscle cells. J Vasc Surg 1998;27(3):512-520.
78. Pukac L, Huangpu J, Karnovsky MJ. Platelet-derived growth factor-BB, insulin- like growth factor-I, and phorbol ester activate different signaling pathways for stimulation of vascular smooth muscle cell migration. Exp Cell Res 1998;242(2):548-560.
79. Pukac LA, Carter JE, Ottlinger ME, Karnovsky MJ. Mechanisms of inhibition by heparin of PDGF stimulated MAP kinase activation in vascular smooth muscle cells. J Cell Physiol 1997;172(1):69-78.
80. Daum G, Hedin U, Wang Y, Wang T, Clowes AW. Diverse effects of heparin on mitogen-activated protein kinase-dependent signal transduction in vascular smooth muscle cells. Circ Res 1997;81(1):17-23.
81. Rauch BH, Millette E, Kenagy RD, Daum G, Clowes AW. Thrombin- and factor Xa-induced DNA synthesis is mediated by transactivation of fibroblast growth factor receptor-1 in human vascular smooth muscle cells. Circ Res 2004;94(3):340- 345.
82. Hayashi K, Shibata K, Morita T, Iwasaki K, Watanabe M, Sobue K. Insulin receptor substrate-1/SHP-2 interaction, a phenotype-dependent switching machinery of insulin-like growth factor-I signaling in vascular smooth muscle cells. J Biol Chem 2004;279(39):40807-40818.
83. Bingley JA, Hayward IP, Campbell JH, Campbell GR. Arterial heparan sulfate proteoglycans inhibit vascular smooth muscle cell proliferation and phenotype change in vitro and neointimal formation in vivo. J Vasc Surg 1998;28(2):308-318.
84. Lindner V, Olson NE, Clowes AW, Reidy MA. Inhibition of smooth muscle cell proliferation in injured rat arteries. Interaction of heparin with basic fibroblast growth factor. J Clin Invest 1992;90(5):2044-2049.
85. Garg HG, Thompson BT, Hales CA. Structural determinants of antiproliferative activity of heparin on pulmonary artery smooth muscle cells. Am J Physiol Lung Cell Mol Physiol 2000;279(5):L779-789.
68 86. Wright TC, Castellot JJ, Petitou M, Lormeau JC, Choay J, Karnovsky MJ. Structural Determinants of Heparins Growth Inhibitory Activity - Interdependence of Oligosaccharide Size and Charge. Journal of Biological Chemistry 1989;264(3):1534-1542.
87. Guyton JR, Rosenberg RD, Clowes AW, Karnovsky MJ. Inhibition of Rat Arterial Smooth-Muscle Cell-Proliferation by Heparin - Invivo Studies with Anti-Coagulant and Non-Anticoagulant Heparin. CircRes 1980;46(5):625-634.
88. Garg HG, Cindhuchao N, Quinn DA, Hales CA, Thanawiroon C, Capila I, Linhardt RJ. Heparin oligosaccharide sequence and size essential for inhibition of pulmonary artery smooth muscle cell proliferation. Carbohydr Res 2002;337(21-23):2359- 2364.
89. Castellot JJ, Choay J, Lormeau JC, Petitou M, Sache E, Karnovsky MJ. Structural Determinants of the Capacity of Heparin to Inhibit the Proliferation of Vascular Smooth-Muscle Cells .2. Evidence for a Pentasaccharide Sequence That Contains a 3-O-Sulfate Group. J Cell Biol 1986;102(5):1979-1984.
90. Garg HG, Mrabat H, Yu L, Freeman C, Li B, Zhang F, Linhardt RJ, Hales CA. Significance of the 2-O-sulfo group of L-iduronic acid residues in heparin on the growth inhibition of bovine pulmonary artery smooth muscle cells. Carbohydr Res 2008;343(14):2406-2410.
91. Garg HG, Joseph PAM, Yoshida K, Thompson BT, Hales CA. Antiproliferative role of 3-O-sulfate glucosamine in heparin on cultured pulmonary artery smooth muscle cells. Biochem Biophys Res Commun 1996;224(2):468-473.
92. Garg HG, Yu L, Hales CA, Toida T, Islam T, Linhardt RJ. Sulfation patterns in heparin and heparan sulfate: effects on the proliferation of bovine pulmonary artery smooth muscle cells. Biochim Biophys Acta 2003;1639(3):225-231.
93. Garg HG, Joseph PAM, Thompson BT, Hales CA, Toida T, Imanari T, Capila I, Linhardt RJ. Effect of fully sulfated glycosaminoglycans on pulmonary artery smooth muscle cell proliferation. Arch Biochem Biophys 1999;371(2):228-233.
94. Millette E, Rauch BH, Defawe O, Kenagy RD, Daum G, Clowes AW. Platelet- derived growth factor-BB-induced human smooth muscle cell proliferation depends on basic FGF release and FGFR-1 activation. Circ Res 2005;96(2):172-179.
95. Rauch BH, Millette E, Kenagy RD, Daum G, Fischer JW, Clowes AW. Syndecan-4 is required for thrombin-induced migration and proliferation in human vascular smooth muscle cells. J Biol Chem 2005;280(17):17507-17511.
69 96. Bono F, Rigon P, Lamarche I, Savi P, Salel V, Herbert JM. Heparin inhibits the binding of basic fibroblast growth factor to cultured human aortic smooth-muscle cells. Biochem J 1997;326 ( Pt 3):661-668.
97. Rapraeger AC, Krufka A, Olwin BB. Requirement of heparan sulfate for bFGF- mediated fibroblast growth and myoblast differentiation. Science 1991;252(5013):1705-1708.
98. Yayon A, Klagsbrun M, Esko JD, Leder P, Ornitz DM. Cell surface, heparin-like molecules are required for binding of basic fibroblast growth factor to its high affinity receptor. Cell 1991;64(4):841-848.
99. Kalmes A, Vesti BR, Daum G, Abraham JA, Clowes AW. Heparin blockade of thrombin-induced smooth muscle cell migration involves inhibition of epidermal growth factor (EGF) receptor transactivation by heparin-binding EGF-like growth factor. Circ Res 2000;87(2):92-98.
100. Kazi M, Lundmark K, Religa P, Gouda I, Larm O, Ray A, Swedenborg J, Hedin U. Inhibition of rat smooth muscle cell adhesion and proliferation by non- anticoagulant heparins. J Cell Physiol 2002;193(3):365-372.
101. Kenagy RD, Clowes AW. Regulation of baboon arterial smooth muscle cell plasminogen activators by heparin and growth factors. Thromb Res 1995;77(1):55- 61.
102. Fannon M, Forsten KE, Nugent MA. Potentiation and inhibition of bFGF binding by heparin: a model for regulation of cellular response. Biochemistry 2000;39(6):1434-1445.
103. Savage JM, Gilotti AC, Granzow CA, Molina F, Lowe-Krentz LJ. Antibodies against a putative heparin receptor slow cell proliferation and decrease MAPK activation in vascular smooth muscle cells. J Cell Physiol 2001;187(3):283-293.
104. Patel RC, Handy I, Patel CV. Contribution of double-stranded RNA-activated protein kinase toward antiproliferative actions of heparin on vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 2002;22(9):1439-1444.
105. Zhao Y, Xiao W, Templeton DM. Suppression of mitogen-activated protein kinase phosphatase-1 (MKP-1) by heparin in vascular smooth muscle cells. Biochem Pharmacol 2003;66(5):769-776.
70 106. Castellot JJ, Wong K, Herman B, Hoover RL, Albertini DF, Wright TC, Caleb BL, Karnovsky MJ. Binding and Internalization of Heparin by Vascular Smooth-Muscle Cells. J Cell Physiol 1985;124(1):13-20.
107. Quarto N, Amalric F. Heparan sulfate proteoglycans as transducers of FGF-2 signalling. J Cell Sci 1994;107 ( Pt 11):3201-3212.
108. Fasciano S, Patel RC, Handy I, Patel CV. Regulation of vascular smooth muscle proliferation by heparin: inhibition of cyclin-dependent kinase 2 activity by p27(kip1). J Biol Chem 2005;280(16):15682-15689.
109. Hashimoto T, Kihara M, Sato K, Imai N, Tanaka Y, Sakai M, Tamura K, Hirawa N, Toya Y, Kitamura H, Umemura S. Heparin recovers AT1 receptor and its intracellular signal transduction in cultured vascular smooth muscle cells. FEBS Lett 2005;579(1):281-284.
110. Orlandi A, Ropraz P, Gabbiani G. Proliferative Activity and Alpha-Smooth Muscle Actin Expression in Cultured Rat Aortic Smooth-Muscle Cells Are Differently Modulated by Transforming Growth-Factor-Beta-1 and Heparin. Exp Cell Res 1994;214(2):528-536.
111. Stegemann JP, Nerem RM. Altered response of vascular smooth muscle cells to exogenous biochemical stimulation in two- and three-dimensional culture. Exp Cell Res 2003;283(2):146-155.
112. Hinck AP, Archer SJ, Qian SW, Roberts AB, Sporn MB, Weatherbee JA, Tsang ML, Lucas R, Zhang BL, Wenker J, Torchia DA. Transforming growth factor beta 1: three-dimensional structure in solution and comparison with the X-ray structure of transforming growth factor beta 2. Biochemistry 1996;35(26):8517-8534.
113. Bjorkerud S. Effects of transforming growth factor-beta 1 on human arterial smooth muscle cells in vitro. Arterioscler Thromb 1991;11(4):892-902.
114. Owens GK, Geisterfer AA, Yang YW, Komoriya A. Transforming growth factor- beta-induced growth inhibition and cellular hypertrophy in cultured vascular smooth muscle cells. J Cell Biol 1988;107(2):771-780.
115. Deaton RA, Su C, Valencia TG, Grant SR. Transforming growth factor-beta 1- induced expression of smooth muscle marker genes involves activation of PKN and p38 MAPK. Journal of Biological Chemistry 2005;280(35):31172-31181.
116. Topouzis S, Majesky MW. Smooth muscle lineage diversity in the chick embryo. Two types of aortic smooth muscle cell differ in growth and receptor-mediated
71 transcriptional responses to transforming growth factor-beta. Dev Biol 1996;178(2):430-445.
117. Hautmann MB, Madsen CS, Owens GK. A transforming growth factor beta (TGFbeta) control element drives TGFbeta-induced stimulation of smooth muscle alpha-actin gene expression in concert with two CArG elements. J Biol Chem 1997;272(16):10948-10956.
118. Kawai-Kowase K, Sato H, Oyama Y, Kanai H, Sato M, Doi H, Kurabayashi M. Basic fibroblast growth factor antagonizes transforming growth factor-beta1- induced smooth muscle gene expression through extracellular signal-regulated kinase 1/2 signaling pathway activation. Arterioscler Thromb Vasc Biol 2004;24(8):1384-1390.
119. Papetti M, Shujath J, Riley KN, Herman IM. FGF-2 antagonizes the TGF-beta1- mediated induction of pericyte alpha-smooth muscle actin expression: a role for myf-5 and Smad-mediated signaling pathways. Invest Ophthalmol Vis Sci 2003;44(11):4994-5005.
120. Grainger DJ, Metcalfe JC, Grace AA, Mosedale DE. Transforming growth factor- beta dynamically regulates vascular smooth muscle differentiation in vivo. J Cell Sci 1998;111 ( Pt 19):2977-2988.
121. Alberts B. Molecular biology of the cell. 4th ed. New York: Garland Science, 2002.
122. Nishimura G, Manabe I, Tsushima K, Fujiu K, Oishi Y, Imai Y, Maemura K, Miyagishi M, Higashi Y, Kondoh H, Nagai R. DeltaEF1 mediates TGF-beta signaling in vascular smooth muscle cell differentiation. Dev Cell 2006;11(1):93- 104.
123. Yoshida T, Hoofnagle MH, Owens GK. Myocardin and Prx1 contribute to angiotensin II-induced expression of smooth muscle alpha-actin. Circ Res 2004;94(8):1075-1082.
124. Turla MB, Thompson MM, Corjay MH, Owens GK. Mechanisms of angiotensin II- and arginine vasopressin-induced increases in protein synthesis and content in cultured rat aortic smooth muscle cells. Evidence for selective increases in smooth muscle isoactin expression. Circ Res 1991;68(1):288-299.
125. Zhang F, Sun AS, Yu LM, Wu Q, Gong QH. Effects of isorhynchophylline on angiotensin II-induced proliferation in rat vascular smooth muscle cells. J Pharm Pharmacol 2008;60(12):1673-1678.
72 126. Watanabe T, Pakala R, Katagiri T, Benedict CR. Serotonin potentiates angiotensin II - induced vascular smooth muscle cell proliferation. Atherosclerosis 2001;159(2):269-279.
127. Geisterfer AAT, Peach MJ, Owens GK. Angiotensin-Ii Induces Hypertrophy, Not Hyperplasia, of Cultured Rat Aortic Smooth-Muscle Cells. CircRes 1988;62(4):749-756.
128. Moiseeva EP. Adhesion receptors of vascular smooth muscle cells and their functions. Cardiovasc Res 2001;52(3):372-386.
129. Hayward IP, Bridle KR, Campbell GR, Underwood PA, Campbell JH. Effect of extracellular matrix proteins on vascular smooth muscle cell phenotype. Cell Biol Int 1995;19(10):839-846.
130. Thyberg J, Hultgardh-Nilsson A. Fibronectin and the basement membrane components laminin and collagen type IV influence the phenotypic properties of subcultured rat aortic smooth muscle cells differently. Cell Tissue Res 1994;276(2):263-271.
131. Yamamoto M, Yamamoto K, Noumura T. Type I collagen promotes modulation of cultured rabbit arterial smooth muscle cells from a contractile to a synthetic phenotype. Exp Cell Res 1993;204(1):121-129.
132. Hedin UL, Daum G, Clowes AW. Disruption of integrin alpha 5 beta 1 signaling does not impair PDGF-BB-mediated stimulation of the extracellular signal- regulated kinase pathway in smooth muscle cells. J Cell Physiol 1997;172(1):109- 116.
133. Hedin U, Bottger BA, Luthman J, Johansson S, Thyberg J. A substrate of the cell- attachment sequence of fibronectin (Arg-Gly-Asp-Ser) is sufficient to promote transition of arterial smooth muscle cells from a contractile to a synthetic phenotype. Dev Biol 1989;133(2):489-501.
134. Qin H, Ishiwata T, Wang R, Kudo M, Yokoyama M, Naito Z, Asano G. Effects of extracellular matrix on phenotype modulation and MAPK transduction of rat aortic smooth muscle cells in vitro. Exp Mol Pathol 2000;69(2):79-90.
135. Zheng B, Duan C, Clemmons DR. The effect of extracellular matrix proteins on porcine smooth muscle cell insulin-like growth factor (IGF) binding protein-5 synthesis and responsiveness to IGF-I. J Biol Chem 1998;273(15):8994-9000.
73 136. Hirst SJ, Twort CH, Lee TH. Differential effects of extracellular matrix proteins on human airway smooth muscle cell proliferation and phenotype. Am J Respir Cell Mol Biol 2000;23(3):335-344.
137. Tran T, McNeill KD, Gerthoffer WT, Unruh H, Halayko AJ. Endogenous laminin is required for human airway smooth muscle cell maturation. Respir Res 2006;7:117.
138. Karnik SK, Brooke BS, Bayes-Genis A, Sorensen L, Wythe JD, Schwartz RS, Keating MT, Li DY. A critical role for elastin signaling in vascular morphogenesis and disease. Development 2003;130(2):411-423.
139. Karnik SK, Wythe JD, Sorensen L, Brooke BS, Urness LD, Li DY. Elastin induces myofibrillogenesis via a specific domain, VGVAPG. Matrix Biol 2003;22(5):409- 425.
140. Hedin U, Thyberg J, Roy J, Dumitrescu A, Tran PK. Role of tyrosine kinases in extracellular matrix-mediated modulation of arterial smooth muscle cell phenotype. Arterioscler Thromb Vasc Biol 1997;17(10):1977-1984.
141. Welser JV, Lange N, Singer CA, Elorza M, Scowen P, Keef KD, Gerthoffer WT, Burkin DJ. Loss of the alpha7 integrin promotes extracellular signal-regulated kinase activation and altered vascular remodeling. Circ Res 2007;101(7):672-681.
142. Wilson E. Alpha 7 beta 1 integrin: putting the brakes on smooth muscle cell proliferation. Circ Res 2007;101(7):651-653.
143. Williams B. Mechanical influences on vascular smooth muscle cell function. J Hypertens 1998;16(12 Pt 2):1921-1929.
144. Wilson E, Mai Q, Sudhir K, Weiss RH, Ives HE. Mechanical strain induces growth of vascular smooth muscle cells via autocrine action of PDGF. J Cell Biol 1993;123(3):741-747.
145. Cappadona C, Redmond EM, Theodorakis NG, McKillop IH, Hendrickson R, Chhabra A, Sitzmann JV, Cahill PA. Phenotype dictates the growth response of vascular smooth muscle cells to pulse pressure in vitro. Exp Cell Res 1999;250(1):174-186.
146. Stegemann JP, Nerem RM. Phenotype modulation in vascular tissue engineering using biochemical and mechanical stimulation. Ann Biomed Eng 2003;31(4):391- 402.
74 147. Jeong SI, Kwon JH, Lim JI, Cho SW, Jung Y, Sung WJ, Kim SH, Kim YH, Lee YM, Kim BS, Choi CY, Kim SJ. Mechano-active tissue engineering of vascular smooth muscle using pulsatile perfusion bioreactors and elastic PLCL scaffolds. Biomaterials 2005;26(12):1405-1411.
148. Kim BS, Nikolovski J, Bonadio J, Mooney DJ. Cyclic mechanical strain regulates the development of engineered smooth muscle tissue. Nat Biotechnol 1999;17(10):979-983.
149. Nikolovski J, Kim BS, Mooney DJ. Cyclic strain inhibits switching of smooth muscle cells to an osteoblast-like phenotype. Faseb J 2003;17(3):455-457.
150. Hipper A, Isenberg G. Cyclic mechanical strain decreases the DNA synthesis of vascular smooth muscle cells. Pflugers Arch 2000;440(1):19-27.
151. Chen Q, Li W, Quan Z, Sumpio BE. Modulation of vascular smooth muscle cell alignment by cyclic strain is dependent on reactive oxygen species and P38 mitogen-activated protein kinase. J Vasc Surg 2003;37(3):660-668.
152. Li W, Chen Q, Mills I, Sumpio BE. Involvement of S6 kinase and p38 mitogen activated protein kinase pathways in strain-induced alignment and proliferation of bovine aortic smooth muscle cells. J Cell Physiol 2003;195(2):202-209.
153. Standley PR, Cammarata A, Nolan BP, Purgason CT, Stanley MA. Cyclic stretch induces vascular smooth muscle cell alignment via NO signaling. Am J Physiol Heart Circ Physiol 2002;283(5):H1907-1914.
154. Qu MJ, Liu B, Wang HQ, Yan ZQ, Shen BR, Jiang ZL. Frequency-dependent phenotype modulation of vascular smooth muscle cells under cyclic mechanical strain. J Vasc Res 2007;44(5):345-353.
155. Dartsch PC, Hammerle H, Betz E. Orientation of cultured arterial smooth muscle cells growing on cyclically stretched substrates. Acta Anat (Basel) 1986;125(2):108-113.
156. Liu B, Qu MJ, Qin KR, Li H, Li ZK, Shen BR, Jiang ZL. Role of cyclic strain frequency in regulating the alignment of vascular smooth muscle cells in vitro. Biophys J 2008;94(4):1497-1507.
157. Beamish JA, Fu A, Choi A, Haq N, Kottke-Marchant K, Marchant RE. The influence of RGD-bearing hydrogels on the re-expression of contractile vascular smooth muscle cell phenotype. Biomaterials 2009;in press.
75 158. Gartner LP, Hiatt JL. Color atlas of histology. 4th ed. Philadelphia: Lippincott Williams & Wilkins, 2006.
159. Wilson E, Sudhir K, Ives HE. Mechanical strain of rat vascular smooth muscle cells is sensed by specific extracellular matrix/integrin interactions. J Clin Invest 1995;96(5):2364-2372.
160. Tock J, Van Putten V, Stenmark KR, Nemenoff RA. Induction of SM-alpha-actin expression by mechanical strain in adult vascular smooth muscle cells is mediated through activation of JNK and p38 MAP kinase. Biochem Biophys Res Commun 2003;301(4):1116-1121.
161. Standley PR, Obards TJ, Martina CL. Cyclic stretch regulates autocrine IGF-I in vascular smooth muscle cells: implications in vascular hyperplasia. Am J Physiol 1999;276(4 Pt 1):E697-705.
162. Chapman GB, Durante W, Hellums JD, Schafer AI. Physiological cyclic stretch causes cell cycle arrest in cultured vascular smooth muscle cells. Am J Physiol Heart Circ Physiol 2000;278(3):H748-754.
163. Birukov KG, Shirinsky VP, Stepanova OV, Tkachuk VA, Hahn AW, Resink TJ, Smirnov VN. Stretch affects phenotype and proliferation of vascular smooth muscle cells. Mol Cell Biochem 1995;144(2):131-139.
164. O'Callaghan CJ, Williams B. Mechanical strain-induced extracellular matrix production by human vascular smooth muscle cells: role of TGF-beta(1). Hypertension 2000;36(3):319-324.
165. Spofford CM, Chilian WM. The elastin-laminin receptor functions as a mechanotransducer in vascular smooth muscle. Am J Physiol Heart Circ Physiol 2001;280(3):H1354-1360.
166. Joki N, Kaname S, Hirakata M, Hori Y, Yamaguchi T, Fujita T, Katoh T, Kurokawa K. Tyrosine-kinase dependent TGF-beta and extracellular matrix expression by mechanical stretch in vascular smooth muscle cells. Hypertens Res 2000;23(2):91-99.
167. Mann BK, Schmedlen RH, West JL. Tethered-TGF-beta increases extracellular matrix production of vascular smooth muscle cells. Biomaterials 2001;22(5):439- 444.
76 168. Hahn MS, McHale MK, Wang E, Schmedlen RH, West JL. Physiologic pulsatile flow bioreactor conditioning of poly(ethylene glycol)-based tissue engineered vascular grafts. Ann Biomed Eng 2007;35(2):190-200.
169. Bulick AS, Munoz-Pinto DJ, Qu X, Mani M, Cristancho D, Urban M, Hahn MS. Impact of endothelial cells and mechanical conditioning on smooth muscle cell extracellular matrix production and differentiation. Tissue Eng Part A 2009;15(4):815-825.
170. Reusch HP, Chan G, Ives HE, Nemenoff RA. Activation of JNK/SAPK and ERK by mechanical strain in vascular smooth muscle cells depends on extracellular matrix composition. Biochem Biophys Res Commun 1997;237(2):239-244.
171. Li Q, Muragaki Y, Hatamura I, Ueno H, Ooshima A. Stretch-induced collagen synthesis in cultured smooth muscle cells from rabbit aortic media and a possible involvement of angiotensin II and transforming growth factor-beta. J Vasc Res 1998;35(2):93-103.
172. Davies PF, Olesen SP, Clapham DE, Morrel EM, Schoen FJ. Endothelial Communication - State of the Art Lecture. Hypertension 1988;11(6):563-572.
173. Thomae KR, Nakayama DK, Billiar TR, Simmons RL, Pitt BR, Davies P. The effect of nitric oxide on fetal pulmonary artery smooth muscle growth. J Surg Res 1995;59(3):337-343.
174. Janakidevi K, Fisher MA, Del Vecchio PJ, Tiruppathi C, Figge J, Malik AB. Endothelin-1 stimulates DNA synthesis and proliferation of pulmonary artery smooth muscle cells. Am J Physiol 1992;263(6 Pt 1):C1295-1301.
175. Fillinger MF, Sampson LN, Cronenwett JL, Powell RJ, Wagner RJ. Coculture of endothelial cells and smooth muscle cells in bilayer and conditioned media models. J Surg Res 1997;67(2):169-178.
176. Heydarkhan-Hagvall S, Helenius G, Johansson BR, Li JY, Mattsson E, Risberg B. Co-culture of endothelial cells and smooth muscle cells affects gene expression of angiogenic factors. J Cell Biochem 2003;89(6):1250-1259.
177. Rose SL, Babensee JE. Complimentary endothelial cell/smooth muscle cell co- culture systems with alternate smooth muscle cell phenotypes. Ann Biomed Eng 2007;35(8):1382-1390.
77 178. Rose SL, Babensee JE. Smooth muscle cell phenotype alters cocultured endothelial cell response to biomaterial-pretreated leukocytes. J Biomed Mater Res A 2008;84(3):661-671.
179. Yoshida H, Nakamura M, Makita S, Hiramori K. Paracrine effect of human vascular endothelial cells on human vascular smooth muscle cell proliferation: transmembrane co-culture method. Heart Vessels 1996;11(5):229-233.
180. Williams C, Wick TM. Endothelial cell-smooth muscle cell co-culture in a perfusion bioreactor system. Ann Biomed Eng 2005;33(7):920-928.
181. Haruguchi H, Teraoka S. Intimal hyperplasia and hemodynamic factors in arterial bypass and arteriovenous grafts: a review. J Artif Organs 2003;6(4):227-235.
182. Motwani JG, Topol EJ. Aortocoronary saphenous vein graft disease: pathogenesis, predisposition, and prevention. Circulation 1998;97(9):916-931.
183. Hancock WW, Adams DH, Wyner LR, Sayegh MH, Karnovsky MJ. CD4+ mononuclear cells induce cytokine expression, vascular smooth muscle cell proliferation, and arterial occlusion after endothelial injury. Am J Pathol 1994;145(5):1008-1014.
184. Tanaka H, Sukhova GK, Swanson SJ, Clinton SK, Ganz P, Cybulsky MI, Libby P. Sustained Activation of Vascular Cells and Leukocytes in the Rabbit Aorta after Balloon Injury. Circulation 1993;88(4):1788-1803.
185. Okamoto E, Couse T, De Leon H, Vinten-Johansen J, Goodman RB, Scott NA, Wilcox JN. Perivascular inflammation after balloon angioplasty of porcine coronary arteries. Circulation 2001;104(18):2228-2235.
186. Schultz K, Murthy V, Tatro JB, Beasley D. Endogenous interleukin-1 alpha promotes a proliferative and proinflammatory phenotype in human vascular smooth muscle cells. Am J Physiol Heart Circ Physiol 2007;292(6):H2927-2934.
187. Yue TL, Wang X, Sung CP, Olson B, McKenna PJ, Gu JL, Feuerstein GZ. Interleukin-8. A mitogen and chemoattractant for vascular smooth muscle cells. Circ Res 1994;75(1):1-7.
188. Cirillo P, Golino P, Calabro P, Cali G, Ragni M, De Rosa S, Cimmino G, Pacileo M, De Palma R, Forte L, Gargiulo A, Corigliano FG, Angri V, Spagnuolo R, Nitsch L, Chiariello M. C-reactive protein induces tissue factor expression and promotes smooth muscle and endothelial cell proliferation. Cardiovasc Res 2005;68(1):47-55.
78 189. Heo SK, Yun HJ, Park WH, Park SD. Emodin inhibits TNF-alpha-induced human aortic smooth-muscle cell proliferation via caspase- and mitochondrial-dependent apoptosis. J Cell Biochem 2008;105(1):70-80.
190. Miano JM. Serum response factor: toggling between disparate programs of gene expression. J Mol Cell Cardiol 2003;35(6):577-593.
191. Prywes R, Roeder RG. Purification of the c-fos enhancer-binding protein. Mol Cell Biol 1987;7(10):3482-3489.
192. Treisman R. Identification of a protein-binding site that mediates transcriptional response of the c-fos gene to serum factors. Cell 1986;46(4):567-574.
193. Norman C, Runswick M, Pollock R, Treisman R. Isolation and properties of cDNA clones encoding SRF, a transcription factor that binds to the c-fos serum response element. Cell 1988;55(6):989-1003.
194. Boxer LM, Prywes R, Roeder RG, Kedes L. The sarcomeric actin CArG-binding factor is indistinguishable from the c-fos serum response factor. Mol Cell Biol 1989;9(2):515-522.
195. Sun Q, Chen G, Streb JW, Long X, Yang Y, Stoeckert CJ, Jr., Miano JM. Defining the mammalian CArGome. Genome Res 2006;16(2):197-207.
196. Wang D, Chang PS, Wang Z, Sutherland L, Richardson JA, Small E, Krieg PA, Olson EN. Activation of cardiac gene expression by myocardin, a transcriptional cofactor for serum response factor. Cell 2001;105(7):851-862.
197. Mack CP, Hinson JS. Regulation of smooth muscle differentiation by the myocardin family of serum response factor co-factors. J Thromb Haemost 2005;3(9):1976-1984.
198. Hendrix JA, Wamhoff BR, McDonald OG, Sinha S, Yoshida T, Owens GK. 5' CArG degeneracy in smooth muscle alpha-actin is required for injury-induced gene suppression in vivo. J Clin Invest 2005;115(2):418-427.
199. van Tuyn J, Knaan-Shanzer S, van de Watering MJ, de Graaf M, van der Laarse A, Schalij MJ, van der Wall EE, de Vries AA, Atsma DE. Activation of cardiac and smooth muscle-specific genes in primary human cells after forced expression of human myocardin. Cardiovasc Res 2005;67(2):245-255.
200. Manabe I, Owens GK. Recruitment of serum response factor and hyperacetylation of histones at smooth muscle-specific regulatory regions during differentiation of a
79 novel P19-derived in vitro smooth muscle differentiation system. Circ Res 2001;88(11):1127-1134.
201. Li S, Wang DZ, Wang Z, Richardson JA, Olson EN. The serum response factor coactivator myocardin is required for vascular smooth muscle development. Proc Natl Acad Sci U S A 2003;100(16):9366-9370.
202. Pipes GC, Sinha S, Qi X, Zhu CH, Gallardo TD, Shelton J, Creemers EE, Sutherland L, Richardson JA, Garry DJ, Wright WE, Owens GK, Olson EN. Stem cells and their derivatives can bypass the requirement of myocardin for smooth muscle gene expression. Dev Biol 2005;288(2):502-513.
203. Owens GK, Kumar MS, Wamhoff BR. Molecular regulation of vascular smooth muscle cell differentiation in development and disease. Physiol Rev 2004;84(3):767-801.
204. Kawai-Kowase K, Kumar MS, Hoofnagle MH, Yoshida T, Owens GK. PIAS1 activates the expression of smooth muscle cell differentiation marker genes by interacting with serum response factor and class I basic helix-loop-helix proteins. Mol Cell Biol 2005;25(18):8009-8023.
205. Miralles F, Posern G, Zaromytidou AI, Treisman R. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 2003;113(3):329-342.
206. Cavigelli M, Dolfi F, Claret FX, Karin M. Induction of c-fos expression through JNK-mediated TCF/Elk-1 phosphorylation. Embo J 1995;14(23):5957-5964.
207. Zhou J, Hu G, Herring BP. Smooth muscle-specific genes are differentially sensitive to inhibition by Elk-1. Mol Cell Biol 2005;25(22):9874-9885.
208. Yoshida T, Gan Q, Owens GK. Kruppel-like factor 4, Elk-1, and histone deacetylases cooperatively suppress smooth muscle cell differentiation markers in response to oxidized phospholipids. Am J Physiol Cell Physiol 2008;295(5):C1175- 1182.
209. Liu Y, Sinha S, Owens G. A transforming growth factor-beta control element required for SM alpha-actin expression in vivo also partially mediates GKLF- dependent transcriptional repression. J Biol Chem 2003;278(48):48004-48011.
210. Adam PJ, Regan CP, Hautmann MB, Owens GK. Positive- and negative-acting Kruppel-like transcription factors bind a transforming growth factor beta control element required for expression of the smooth muscle cell differentiation marker SM22alpha in vivo. J Biol Chem 2000;275(48):37798-37806.
80 211. Liu Y, Sinha S, McDonald OG, Shang Y, Hoofnagle MH, Owens GK. Kruppel-like factor 4 abrogates myocardin-induced activation of smooth muscle gene expression. J Biol Chem 2005;280(10):9719-9727.
212. Kawai-Kowase K, Ohshima T, Matsui H, Tanaka T, Shimizu T, Iso T, Arai M, Owens GK, Kurabayashi M. PIAS1 mediates TGFbeta-induced SM alpha-actin gene expression through inhibition of KLF4 function-expression by protein sumoylation. Arterioscler Thromb Vasc Biol 2009;29(1):99-106.
213. Doi H, Iso T, Yamazaki M, Akiyama H, Kanai H, Sato H, Kawai-Kowase K, Tanaka T, Maeno T, Okamoto E, Arai M, Kedes L, Kurabayashi M. HERP1 inhibits myocardin-induced vascular smooth muscle cell differentiation by interfering with SRF binding to CArG box. Arterioscler Thromb Vasc Biol 2005;25(11):2328-2334.
214. Dandre F, Owens GK. Platelet-derived growth factor-BB and Ets-1 transcription factor negatively regulate transcription of multiple smooth muscle cell differentiation marker genes. Am J Physiol Heart Circ Physiol 2004;286(6):H2042- 2051.
215. Higgins SP, Solan AK, Niklason LE. Effects of polyglycolic acid on porcine smooth muscle cell growth and differentiation. J Biomed Mater Res A 2003;67(1):295-302.
216. Mann BK, Tsai AT, Scott-Burden T, West JL. Modification of surfaces with cell adhesion peptides alters extracellular matrix deposition. Biomaterials 1999;20(23- 24):2281-2286.
217. Mann BK, West JL. Cell adhesion peptides alter smooth muscle cell adhesion, proliferation, migration, and matrix protein synthesis on modified surfaces and in polymer scaffolds. J Biomed Mater Res 2002;60(1):86-93.
218. Joddar B, Ibrahim S, Ramamurthi A. Impact of delivery mode of hyaluronan oligomers on elastogenic responses of adult vascular smooth muscle cells. Biomaterials 2007;28(27):3918-3927.
219. Peyton SR, Raub CB, Keschrumrus VP, Putnam AJ. The use of poly(ethylene glycol) hydrogels to investigate the impact of ECM chemistry and mechanics on smooth muscle cells. Biomaterials 2006;27(28):4881-4893.
220. Li S, Lao J, Chen BP, Li YS, Zhao Y, Chu J, Chen KD, Tsou TC, Peck K, Chien S. Genomic analysis of smooth muscle cells in 3-dimensional collagen matrix. Faseb J 2003;17(1):97-99.
81 221. Hong H, McCullough CM, Stegemann JP. The role of ERK signaling in protein hydrogel remodeling by vascular smooth muscle cells. Biomaterials 2007;28(26):3824-3833.
222. Mann BK, Gobin AS, Tsai AT, Schmedlen RH, West JL. Smooth muscle cell growth in photopolymerized hydrogels with cell adhesive and proteolytically degradable domains: synthetic ECM analogs for tissue engineering. Biomaterials 2001;22(22):3045-3051.
223. Adelow C, Segura T, Hubbell JA, Frey P. The effect of enzymatically degradable poly(ethylene glycol) hydrogels on smooth muscle cell phenotype. Biomaterials 2008;29(3):314-326.
224. Peyton SR, Kim PD, Ghajar CM, Seliktar D, Putnam AJ. The effects of matrix stiffness and RhoA on the phenotypic plasticity of smooth muscle cells in a 3-D biosynthetic hydrogel system. Biomaterials 2008;29(17):2597-2607.
225. Brophy CM, Woodrum DA, Pollock J, Dickinson M, Komalavilas P, Cornwell TL, Lincoln TM. cGMP-dependent protein kinase expression restores contractile function in cultured vascular smooth muscle cells. J Vasc Res 2002;39(2):95-103.
226. Stegemann JP, Dey NB, Lincoln TM, Nerem RM. Genetic modification of smooth muscle cells to control phenotype and function in vascular tissue engineering. Tissue Eng 2004;10(1-2):189-199.
227. Shen JY, Chan-Park MB, He B, Zhu AP, Zhu X, Beuerman RW, Yang EB, Chen W, Chan V. Three-dimensional microchannels in biodegradable polymeric films for control orientation and phenotype of vascular smooth muscle cells. Tissue Eng 2006;12(8):2229-2240.
228. Rowlands AS, Cooper-White JJ. Directing phenotype of vascular smooth muscle cells using electrically stimulated conducting polymer. Biomaterials 2008;29(34):4510-4520.
229. Wamhoff BR, Bowles DK, McDonald OG, Sinha S, Somlyo AP, Somlyo AV, Owens GK. L-type voltage-gated Ca2+ channels modulate expression of smooth muscle differentiation marker genes via a rho kinase/myocardin/SRF-dependent mechanism. Circ Res 2004;95(4):406-414.
82 CHAPTER 3: Vascular Tissue Engineering
3.1. Introduction
Tissue engineering is the application of biology, materials science, and
engineering to restore, maintain, or improve tissue function [1]. Central to this approach
is the use of living cells to provide the complex set of responses and behaviors necessary
to recapitulate tissue function. Scaffold systems and/or culture conditions are used to
guide cell behavior. This chapter briefly reviews recent progress toward the development
of functional tissue engineered blood vessels (TEBVs) using these principles.
3.2. Normal Blood Vessel Histology
Blood vessels range in size from the aorta (1 cm scale) to capillaries (10 μm
scale). The most pressing clinical need is for small diameter (< 5 mm) muscular artery
substitutes. The histology of these blood vessels is classically divided into three layers
(from lumen outward): the tunica intima, the tunica media, and the tunica adventitia (Fig.
3.1) [2]. The tunica intima normally consists of endothelial cells (ECs) and their
basement membrane. ECs normally prevent blood coagulation and thrombosis, but if
injured or stimulated, can promote thrombosis, coagulation, and recruitment of
inflammatory cells [3]. ECs also sense hemodynamic conditions [3]. The intima is
separated from the tunica media by the internal elastic lamina (IEL). In pathological conditions (see Chapter 1), smooth muscle cells (SMCs) penetrate the IEL and proliferate in the intima, hence this process is called “intimal” hyperplasia. The tunica media consists of circumferentially aligned SMCs and interwoven collagen fibers and elastic laminae [2]. The media is the primary load bearing portion of the vessels, imparts its elasticity, and regulates vessel diameter via contraction/relaxation of the SMCs [3]. The
83 tunica adventitia consists of connective tissue and the vascular supply for the blood
vessel itself, known as vaso vasorum, which send capillaries into the tunica media. The
goal of vascular tissue engineering is to regenerate functional forms of all three tissue
layers.
3.3. Design Criteria
Design criteria for TEBVs have been considered widely over the last several
decades and have remained largely unchanged [4, 5]. These criteria are summarized in
Table 3.1. Briefly, the graft should be biocompatible (non-toxic and
immunocompatible), resistant to infection, thrombosis, and intimal hyperplasia, easily handled and sutured, readily available (off-the-shelf), and reasonably economical to produce and store.
3.4. Tissue Engineered Modification of Existing Graft Materials
Expanded poly(tetrafluoroethylene) (ePTFE) has been utilized for decades as a graft material for large inner diameter (ID) prostheses (> 6 mm) because it is strong, stable, and biocompatible. Several groups have undertaken work to improve the function of ePTFE using tissue engineering approaches. Endothelialization has been the major thrust of this work, given the importance of this cell layer in regulation of hemostasis. In two clinical studies in humans, the lumen of ePTFE grafts was coated with fibrin glue and seeded with autologous ECs expanded in vitro [6, 7]. These grafts were used for coronary artery bypass in 14 patients [6] or infrainguinal bypass in 136 patients [7]. In the coronary position, grafts (4 mm ID), 19 of 21 grafts (placed in 14 patients) remained patent after 7.5-48 months of follow-up [6]. While still inferior to artery grafts, these results were superior to unmodified ePTFE grafts, which have ~60% patency after 1 year
84
MUSCULAR ARTERY STRUCTURE
LAYERS Intima (blue)
Media (orange)
Endothelium Adventitia (green)
Blood
Internal elastic lamina
Smooth muscle cell Vaso vasorum Elastin fibers Fibroblast Collagen fibers NOT TO SCALE
Figure 3.1-Structure of a muscular artery (not to scale).
85
Table 3.1-Design criteria for tissue engineered blood vessels (TEBV)
Design Criteria
Biocompatible: Non-toxic No cell death No induction of malignancies No toxic degradation products No immune activation No chronic inflammation Non-allergic No specific immune response (i.e. graft rejection)
Maintain long-term patency: Non-thrombogenic Resist intimal hyperplasia
Resist infection/easily sterilized (excluding cellular component)
Adequate material properties: Burst pressure greater than 1700 mmHg Reasonable compliance matching Available in many lengths and diameters Good suture retention Flexible, non-kinking Easily handled
Readily available for emergent procedures
Economical to produce
86 [8], and compared favorably with vein grafts (~90% 1-year patency) [9]. The peripheral
bypass grafts (6-7 mm ID) had an estimated 63% patency after 7 years [7]. These rates
were similar to vein grafts (~70% at 2 years) and superior to unmodified ePTFE grafts
(31% at 2 years) [10]. While this tissue engineering approach is promising because it utilizes materials that are already in clinical use, it also requires ex vivo EC culture several weeks in advance, limiting its use for emergent procedures. Approaches to enhance in vivo endothelialization are currently being explored, including stimulation of transmural endothelialization and promoting attachment of ECs or endothelial progenitor cells (EPCs) via EC/EPC-selective ligands on the graft surface [11, 12].
3.5. Fully Tissue Engineered Blood Vessels (TEBVs)
While the approaches described in section 3.4 may be promising, they are limited by the use of non-resorbable scaffold materials, such as ePTFE. Because ePTFE cannot be remodeled, it may chronically induce intimal hyperplasia due to compliance mismatch
(see Chapter 1) and is poorly suited for pediatric vascular surgery where vessels must grow along with the patient. For these reasons, development of a fully TEBV has attracted much research interest over the last several decades. Many approaches have been explored and will be reviewed below.
3.5.1. Decellularized Scaffolds
The use of decellularized arteries has attracted much interest because these materials already have the appropriate vessel architecture and, at least theoretically, can
be remodeled by invading cells. These scaffolds are typically explanted, decellularized
with a combination of enzymes, surfactants, and base solutions, and sometimes
chemically modified to improve mechanical or biological properties. For example,
87 Conklin et al. tested decellularized porcine common carotid arteries that were modified
on the lumen with heparin to prevent thrombosis [13]. Limited in vivo testing in a canine
carotid model showed patency for 67 d and showed SMC infiltration and
endothelialization, although there were no controls implanted with which to compare in
this study [13].
In contrast, most studies have attempted to recellularize the grafts, at least partially, prior to implantation. Borschel et al. tested decellularized rat iliac arteries seeded with or without in vitro expanded ECs in a femoral interposition model in rats
[14]. After 4 weeks, 89% of the EC-seeded grafts remained patent compared with 29% of EC-unseeded grafts [14]. Grafts showed infiltration of smooth muscle α-actin (SMαA)
positive cells, a well formed endothelium, and mechanical properties similar to native
arteries [14]. Kaushal et al. used decellularized porcine iliac arteries seeded with
autologous endothelial progenitor cells (EPCs), which were shear preconditioned ex vivo
and tested in an ovine carotid interposition model for 130 d [15]. Grafts without EPC
seeding failed due to thrombosis within 5 d, while EPC-seeded grafts remained patent for
30 d [15]. The decellularized scaffold showed evidence of remodeling, SMC infiltration,
and vasoactivity [15]. Cho and colleagues explored the use of decellularized vessels
seeded with bone marrow derived cells (BMCs) to provide both ECs and SMCs [16, 17].
Decellularized canine carotid artery scaffolds were seeded with autologous BMCs and
implanted in a canine carotid interposition model [16]. After 8 weeks 40% of the seeded
grafts were patent compared with none of the unseeded grafts [16]. Subsequently, the
same approach was used with an abdominal aortic interposition model in juvenile pigs
[17]. Owing to the larger size of the graft (7 mm vs. 3 mm for the canine experiments),
88 all of these grafts remained patent, showed remodeling and endothelialization, and even
showed limited growth as the juvenile animals matured [17].
Other tubular tissues also have been explored as decellularized scaffolds. Schaner
et al. generated decellularized saphenous vein allografts with mechanical properties that
were similar to native vein and implanted them in a canine carotid interposition model for
2 weeks [18]. Despite the lack of endothelium, no thrombosis was noted in these short-
term studies. Daniel et al. developed an ex vivo processing method to prepare
decellularized human umbilical vein grafts with reproducible properties by freezing
samples on a mandrel and using a lathe to uniformly remove adventitial tissue [19]. The
process reduced variability between samples compared with manual dissection and
vessels retained adequate mechanical properties [19]. Although these grafts permitted
some cellular in-growth in vitro, they were not tested in vivo [19]. Huynh et al. formed
tubes from flaps of cleaned, decellularized intestinal submucosa that was wrapped
around a 4 mm mandrel, cross-linked, and coated with a thin luminal layer of collagen
[20]. These grafts remained patent for 90 d in a rabbit carotid interposition model and
showed infiltration of SMαA positive cells, re-endothelialization, and vasoactivity [20].
3.5.2. Scaffoldless Approaches
Another approach, developed by L’Heureux and co-workers, has been the use of
blood vessels engineered from sheets of cultured human cells [21-23]. In early studies, a procedure was developed to recapitulate the endothelium-IEL-media-adventitia structure of native blood vessels. To form these constructs acellular collagen sheets and then sheets of SMCs were wrapped around a 3 mm mandrel, matured in culture, surrounded by fibroblast sheets, matured in culture again, removed from the mandrel, and, in some in
89 vitro studies, seeded with ECs [21]. These constructs were placed in a canine femoral interposition model without ECs [21]. The resulting constructs developed robust mechanical properties in vitro and were able to be implanted in vivo surgically, although the patency rate (50% at 7 d) was poor, despite immunosuppressive and anticoagulant therapy, owing to a lack of endothelial pre-seeding and immunological rejection of the human cells used for graft preparation [21]. Since it was determined in later studies that the mechanical properties of these constructs were conferred mainly by the fibroblast layer, the SMCs were omitted. Composite grafts formed from human cells, which also contained an endothelial layer in all cases, were utilized in a variety of animal models
(dog, athymic rat, and macaque) to demonstrate surgical feasibility and long-term patency [22]. Overall patency in athymic rat aortic interposition studies was ~85% after
225 d and all of the grafts (N = 4) in the macaque model remained patent for 8 weeks
[22]. Recently, autologous graft materials formed using these techniques have been used as shunts for hemodialysis in human trials with moderate success (60% patency at 6 months) [23].
3.5.3. Synthetic Scaffolds Made from Natural Materials
Decellularized scaffolds have limited potential for modification and engineering.
A variety of natural materials have been explored to increase the engineering control of scaffold properties, while retaining the remodeling potential observed with decellularized
TEBVs. In particular, type I collagen has been used widely as a scaffold material for
TEBVs because it is one of the major structural extracellular matrix (ECM) proteins in the vasculature. Collagen can be readily formed into gels in vitro, allowing collagen-
90 based scaffold systems to be formed into a variety of geometries, including tubes for
vascular tissue engineering.
Weinberg and Bell are widely credited with the development of the first truly
tissue engineered blood vessel scaffold, based on a collagen gel [24]. SMCs, collagen,
and medium were poured into an annular mold, the resulting gel was cultured for 1 week
(during which it contracted considerably), a Dacron mesh was slipped around the outside, additional layers were formed with SMCs or fibroblasts in a similar fashion, and the lumen was seeded with ECs [24]. The resulting constructs had the gross appearance of muscular arteries, but, even with the Dacron reinforcement, had poor mechanical strength
and lacked cellular organization and alignment, making them inadequate for any in vivo
testing [24].
Tranquillo et al. attempted to overcome this problem by using a strong magnetic
field to orient growing collagen fibrils circumferentially during gel formation, while following a procedure that was otherwise similar to Weinberg and Bell [25]. The resulting constructs were stronger than those prepared without magnetic orientation, with the overall mechanical properties dependent on the cell density and culture duration [25].
However, the constructs were still significantly weaker than native vessels [25-27].
Tranquillo and co-workers also found that glycation (i.e., incubation of the collagen matrices with glucose or ribose) increased the mechanical modulus of the materials several fold [28]. However, the biological impact of this strategy is not clear.
Mechanical stimulation to enhance the mechanical properties of collagen-based
TEBVs has been explored extensively by Nerem and co-workers. After forming a collagen gel-rat aortic SMC construct on a hollow silicone mandrel, pulsatile mechanical
91 stimulation was applied at 1 Hz and ~10% radial strain for up to 8 d [29]. This process
resulted in a 2-3 fold increase in the mechanical modulus of the constructs, as well as increased compaction, increased yield stress and yield strain, and cellular and collagen circumferential alignment [29]. The final modulus obtained (~120 kPa) approached the required mechanical properties, although the compliance of these structures was greater than native arteries and the burst strength was not measured [29]. However, it was later observed that these results appeared to be cell-type dependent, with human SMCs and fibroblast showing attenuated strengthening in response to mechanical stimulation, perhaps because of altered matrix metalloproteinase (MMP) expression [30]. The supplementation of exogenous factors, particularly transforming growth factor (TGF)-β1, have also been shown to augment mechanical stimulation to increase gel compaction and cell alignment [31].
Fibrin gels have also attracted interest as natural materials for TEBVs. This material can be formed easily into a weak hydrogel and can be remodeled by encapsulated cells. Grassl et al. used fibrin-based gels to form media equivalents with comparable mechanical properties to rat abdominal aorta [32]. These constructs were formed in a similar fashion to collagen constructs: cells and matrix (fibrinogen and thrombin) were poured into an annular mold and cultured for 3-6 weeks to allow the gels to compact and remodel [32]. Constructs that were seeded with neonatal rat SMCs
(which were found to synthesize 2-3 fold greater ECM than adult SMCs) and stimulated with TGF-β1 and insulin developed an ultimate tensile strength (circumferential) that was similar to rat aorta (1.4 MPa vs. 2.1 MPa, respectively) [32]. These grafts were nearly an
92 order of magnitude stronger than collagen grafts prepared using methods similar to
Weinberg and Bell (without the Dacron mesh) [24].
Andreadis and co-workers utilized fibrin scaffolds with fetal lamb SMCs and ECs to construct tissue engineered grafts suitable for low pressure in vivo experiments [33].
ECs were seeded on the abluminal surface of the graft then positioned in the lumen by turning the construct inside-out [33]. Grafts were placed in a low pressure external jugular vein interposition model and remained patent for up to 15 weeks (N=2) [33].
Grafts showed remodeling and the reformation of elastic laminae [33]. As an extension to this work, the same group utilized the SMαA promoter to select bone marrow-derived smooth muscle-like progenitor cells in place of fetal SMCs [34]. These cells displayed high proliferative capacity, yet retained high expression of several SMC marker genes
[34]. When these cells were seeded in fibrin-based tubular reactors and cultured for 2 weeks ex vivo, the resulting constructs displayed contraction in response to KCl and norepinephrine, as well as higher modulus and ultimate strength than control grafts formed with cultured SMCs [34]. Graft histology after 5 weeks in vivo in an ovine jugular vein model, showed a mature vessel wall with contractile marker expressing
SMCs and well-developed elastic laminae and interstitial collagen [34].
Scaffolds formed from modified hyaluronic acid (HYAFF-11, a benzyl ester derivative of hyaluronic acid) have also been tested in a porcine carotid interposition model [35]. HYAFF-11 tubes (4 mm ID) were implanted without any cell pre-seeding and were monitored for 5 months, with 70% patency [35]. Failures were due to intimal hyperplasia and/or thrombosis [35]. The lumen of these vessels was endothelialized and
93 the media was populated with oriented cells and had remodeled with prominent formation
of elastic lamina [35].
3.5.4. Synthetic Scaffolds Made from Biodegradable Polymers
Biodegradable polymers also have attracted interest as scaffold materials for
TEBVs because their properties can be controlled to a much greater extent than natural
scaffold materials. In particular, biodegradable polyesters have been explored
extensively. Niklason et al. developed a poly(glycolic) acid (PGA)-based scaffold
system for engineering small diameter blood vessels [36]. PGA mesh scaffolds were
sewn into tubes, surface treated with sodium hydroxide to improve cell adhesion, seeded
with bovine aortic SMCs, and cultured for up to 8 weeks. The addition of ascorbic acid
and culture under pulsatile conditions (2.8 Hz, 5% radial strain) greatly improved the
burst strength of the vessels [36]. Limited in vivo experiments were performed in miniature swine that demonstrated patency for up to 4 weeks [36]. While an impressive step forward, the luminal portion of these scaffolds were poorly populated and remodeled during culture, and the bulk of the graft’s mechanical properties were conferred by a dense cellular layer on the adventitial side of the graft. The long term feasibility of this approach was not demonstrated in vivo beyond these proof of principle studies (N=1 pre- pulsed graft with autologous cells) [36].
Another approach to improve the mechanical properties of degradable polymeric scaffolds is to employ a two polymer hybrid construct. Shum-Tim et al. used a PGA mesh as a scaffold for vascular cells, but wrapped the construct with an impervious poly(hydroxyalkanoate) (PHA) sheet with slow degradation properties and high
mechanical strength [37]. Hybrid grafts, seeded in vitro with autologous mixed vascular
94 cells, were implanted in an ovine aortic interposition model with patency demonstrated after 5 months [37]. Well-developed histological organization was observed that was similar to native arteries with elastic laminae in the media and ECs lining the lumen [37].
However, after this period the PHA sheet remained intact, making it unclear how the graft would fair as this material degrades over the course of years.
Woven poly(dioxanone) also has been tested as a graft material in a porcine carotid interposition model [38]. The results were poor with greater than 50% of grafts
occluded after 1 week, with the remaining grafts exhibiting significant (> 20 %) stenosis
[38]. These grafts were surrounded by large volumes of connective tissue [38]. These
results strongly suggested this material was not suitable as a scaffold for vascular tissue
engineering, although the grafts were not pre-endothelialized, which may have improved
patency.
Biodegradable polymeric vascular materials have progressed to limited use in humans [39]. Matsumura et al. utilized poly(caprolactone-co-lactide) polymer sponges
reinforced with either PGA or poly(L-lactic acid) (PLLA) fibers [39]. These constructs
were seeded with autologous vascular cells, bone marrow, or purified bone marrow constituents and implanted in low pressure positions in pediatric patients during reconstructive surgery for congenital cardiac malformations [39]. These scaffold systems have been used in 42 patients with no complications reported (follow-up 1 month to 3 years) [40].
3.5.5. Electrospun Scaffolds
Biodegradable polymers also have been used to generate electrospun nanofibrous
scaffolds for vascular tissue engineering. Electrospun fibers are formed from a polymer
95 solution ejected from a high voltage needle toward a grounded target. The electric force overcomes surface tension, to produce a constant stream of liquid instead of droplets
[41]. The target can take on many shapes and can be mechanized to allow the formation of aligned fibers in a variety of geometries.
Atala, Yoo, and co-workers have developed a number of biodegradable polymer- based electrospun vascular graft materials [42-44]. Using a poly(ε-caprolactone) (PCL) and collagen blend (50/50 wt/wt), 4.7 mm ID scaffolds with ~500 nm randomly oriented fiber tubes were formed [42]. Before cell seeding, these tubes displayed similar compliance (0.07 %/mmHg) to native muscular arteries and had a burst pressure of greater than 4000 mmHg, although these measurements were performed with internal tubing [42]. It is unclear how these properties would change if this porous material were loaded directly with fluid. These scaffolds were seeded lumenally with ECs and externally with SMCs and were preconditioned for up to 9 d of culture in vitro [42, 43].
Histology after in vitro experiments showed poor SMC migration into the interstitium of the construct [43]. When placed in vivo as an aorto-iliac bypass in rabbits, 7/8 grafts were patent after 1 month and showed poor tissue in-growth and remodeling by histology
[43]. Atala, Yoo, and coworkers have also developed more complicated fiber compositions containing collagen, elastin and poly(lactic-co-glycolic acid) (PLGA) for use in TEBVs [44].
Hashi et al. utilized electrospun PLLA fiber sheets to generate small diameter (0.7 mm) grafts [45]. In contrast to Atala, Yoo, and colleagues, uniaxial strain was used to align the fibers of the sheet, which was then rolled into a tubular graft [45].
Mesenchymal stem cells (MSCs) were seeded on the sheets before rolling to encapsulate
96 them in the graft [45]. When implanted in a carotid interposition model in athymic rats for 60 d, both MSC-seeded and acellular grafts developed a highly cellularized region near the lumen with moderate cell penetration into the bulk of the scaffold [45].
Acellular graft preparations developed initial thrombus formation and more intimal hyperplasia (IH) [45]. It appeared that the MSCs conferred anti-thrombotic properties to the grafts (studied in vitro) [45]. It is likely that the IH was directly related to early thrombus deposits. Though better than reported by Atala, Yoo, and coworkers [44], cell penetration into the scaffold region was sub-optimal and the remodeling of the graft was less extensive than grafts made from natural materials (discussed above).
Pektok et al. compared 2 mm electrospun PCL scaffolds (random orientation, 1.9
μm fibers) with ePTFE grafts in a rat aortic interposition study for up to 24 weeks [46].
All grafts remained patent and significant stenosis was noted in only 2/15 ePTFE grafts and in none of the PCL grafts [46]. PCL grafts were endothelialized and showed moderate cellular infiltration (cells not identified), but aligned cells and collagen deposition were only prominent near the neointima [46].
3.5.6. Hydrogel Scaffolds
West and co-workers have been developing poly(ethylene glycol) diacrylate
(PEGDA) hydrogels as scaffold materials for TEBVs over the last decade [47-50]. These materials have limited interactions with cells unless specific cell-adhesive ligands are included and have easily controlled physical properties (Chapters 4 and 5, [51, 52]).
Using this material, Hahn et al. formed tubular hydrogels (3 mm ID, 1.7 mm wall thickness) with encapsulated smooth muscle cells [53]. These non-endothelialized tubes were cultured under quasi-physiologic pulsatile conditions (120/80 mmHg, 1-2 Hz) for 8
97 weeks [53]. Although collagen production and cell population were slightly greater in grafts subjected to mechanical stimulation, the constructs remained weak (several times greater compliance than a native artery, despite a significantly thicker vessel wall) and histological analysis showed a lack of cellular remodeling or smooth muscle tissue architecture [53]. In an effort to improve SMC behavior in the constructs, Bulick et al. co-cultured similar constructs with an layer of entrapped endothelial cells [54]. They found that in the presence of mechanical stimulation, ECs may have slightly increased elastin production and calponin expression by the SMCs, but microscopic analysis still showed poor development of tissue architecture [54]. It also is unlikely that encapsulated
ECs behaved normally, as these cells exist in monolayers in vivo.
Though these studies suggest that PEG-based materials are poorly suited for vascular tissue engineering, this work has several key weaknesses. Highly cross-linked hydrogels were formed from non-degradable starting components to enhance the mechanical integrity of the grafts (note that mixing of degradable with non-degradable macromers [53] still yields a non-degradable network). Substantial cellular remodeling of such matrices in an 8-week time span is not probable, as is required for cell spreading in 3D. Even these highly cross-linked, thick hydrogel tubes were weaker than native vessels, suggesting hydrogels are ill-suited to provide mechanical support for grafts. A better approach would be to utilize a lightly cross-linked, degradable, and cell-instructive hydrogel material to control cellular regeneration and a more robust polymer network to bear the mechanical load of the construct.
98 3.6. Discussion
Vascular tissue engineering has attracted much research interest over the past several decades, due to the poor performance of synthetic vascular prostheses in small diameter applications and a pressing clinical need for such grafts. Despite an immense research effort, a reliable TEBV has yet to be developed.
One difficulty with tissue engineered approaches is finding an autologous cell source with sufficient regenerative potential. Patients that need these prostheses are generally elderly and possess cells with limited growth potential, unlike the young cells used in most animal models. For example, a sufficient number of ECs for pre-seeding could not be obtained in about 5% of patients in one clinical trial [7]. Significant differences have also been noted for SMCs depending on source species and age [30, 32].
When SMCs harvested from elderly patients were cultured in PGA scaffolds similar to
Niklason et al. [36], very poor results were obtained (burst pressure ~100 mmHg) [55].
Retrovirally expressing telomerase in these cells improved growth potential, but these older cells still displayed reduced biosynthesis of collagen, resulting in mechanically weak vessels [55]. Promising results have been obtained using bone marrow derived cells, which empirically show enhanced regenerative potential in TEBVs [34, 56].
However, the biology of these cells is not well understood and the mechanism of this effect is not known [34, 56]. These results suggest that finding cell sources and/or culture techniques with sufficient, well-characterized regenerative potential is an important future direction for TEBV research.
Despite cell-source limitations, much progress has been made improving TEBV function. One of the key improvements has been the recognition that a confluent
99 endothelium is critical to prevent early failure due to thrombosis [14, 15]. The utility of
this strategy for improving the performance of ePTFE grafts has been demonstrated in
human trials [6, 7]. However, this approach generally is limited by the necessity of an
extra surgical procedure for EC harvest and extensive ex vivo culture [6, 7]. Thus, this approach is not feasible unless bypass surgery is planned 6-8 weeks in advance, which is not practical in most cases. Unlike humans, grafts implanted in animal models (e.g. pig or rabbit) without endothelium are generally found to be endothelialized at explantation,
after just weeks in vivo [20, 35]. Short graft lengths, which have less graft area to cover,
and the use of young animals facilitate rapid EC coverage [57]. These results have been
difficult to extrapolate to humans because grafts used in bypass procedures are typically
longer (> 10 cm) and human ECs (in patients needing bypass) tend to have less
regenerative potential [57]. Development of strategies to promote in vivo
endothelialization in humans [11, 12] is an important future direction for this work.
Regeneration of a functional, non-hyperplastic medial layer also is critical to
prevent long-term failure due to anastomotic intimal hyperplasia. To date, the results of
the in vivo and in vitro studies reviewed here generally show an inverse relationship
between medial regeneration and the engineering control of the scaffold system (Fig
3.2.). The use of decellularized grafts has resulted in regeneration of smooth muscle
tissue that is remarkably similar to native blood vessels [16, 20]. While encouraging,
these approaches have some fundamental limitations that suggest these successes may not
be translated easily to humans. These scaffolds may retain up to 6% of cellular debris
and, like all natural materials, have the potential to induce an immune response,
especially if derived from xenogeneic sources [18]. The size, shape, mechanical
100
Scaffolds for Vascular Tissue Engineering
↓ Engineering Control ↑ Engineering Control ↑ Success in Animals ↓ Success in Animals
Decellularized Cell Sheets Natural ECM Biodegradable polymers Artery Fibroblasts Collagen Porous solids Vein SMCs Fibrin Woven meshes Intestinal mucosa Hyaluronan Electrospinning Hydrogels
Figure 3.2-Overview of scaffolds used for tissue engineered blood vessels (TEBVs) and animal model results. In studies to date, generally there is an inverse relationship between the remodeling capacity of the scaffold in animal models and the engineering control of the scaffold parameters. Scaffold materials have been placed along the continuum in approximate locations based on the results of in vivo experiments and the relative degree of control each system affords.
101 characteristics, and biological response are largely determined by the explanted tissue
from which the scaffold was constructed, with only minor modifications possible. It also
is difficult to generate this type of construct on a large scale with consistent and reliable
properties. In fact, the use of decellularized scaffolds so greatly sacrifices engineering
control that it is debatable if these are truly tissue engineered constructs.
The use of cell sheets and natural materials, which are engineered in vitro, results
in improved control of scaffold properties. Remodeling of these scaffolds in animal
models also results in tissues with remarkable similarity to normal blood vessels [21, 34,
35]. However, with lengthy and elaborate ex vivo maturation (as long as 3-4 months
[22]) with autologous cells required, these scaffold materials are neither practical nor economical to produce. In fact, given that bypass graft operations are usually urgent, ex vivo cultured TEBVs likely are more useful as experimental model systems than as actual graft materials. Furthermore, while these approaches greatly improve the engineering control of graft design over decellularized scaffolds, graft designs are still limited by the relatively small number of suitable natural materials (collagen, fibrin, hyaluronic acid, etc) and the parameter space they confer (e.g. a fibrin gel with a 1 MPa modulus cannot be made). Chemical modification and hybrid natural-synthetic approaches have expanded the parameter space for these materials [58, 59]. However, like decellularized scaffolds, these materials, especially those that are (bio)chemically re-engineered, have an increased potential to elicit an immune response.
While biodegradable polymeric systems offer significantly more engineering control, animal studies with these systems tend to show poor capacity for remodeling, even when preconditioned in vitro [36, 53]. Rather than suggesting that natural scaffolds
102 are superior, these results reflect the fact that the science of polymeric scaffold design is still relatively young and incomplete. In this context, the successes of degradable polymeric materials for vascular applications are quite encouraging. Furthermore, because of their strength, biodegradable polymers can be implanted with intraoperative cell seeding that does not require in vitro preconditioning [39], suggesting the potential for these easily manufactured materials to act as off-the-shelf vascular grafts. The next generation of polymeric materials seeks to mimic the desirable properties of natural materials, such as cell adhesion and cell-mediated degradation [60, 61], while retaining a high degree of control of material properties. As our understanding of synthetic scaffold design improves, the superior engineering control available in the fabrication of these systems will make this a preferred approach. The development of cell-instructive scaffold systems that can guide the regeneration of vascular structures, yet be easily modified as new design criteria emerge, represents an important goal for vascular tissue engineering.
3.7. PEG-based Hydrogels for Tissue Engineering
Poly(ethylene glycol) (PEG) has biological properties that make it an excellent material for tissue engineering scaffolds. PEG is well hydrated in an aqueous environment which minimizes the adsorption of protein. For this reason, PEG is often classified as bioinert. As a result, cell-scaffold interactions with this material can be controlled specifically, even in the presence of serum [60]. PEGs of all molecular weights have high solubility in water, and, therefore, cannot maintain a scaffold structure in an aqueous environment without modifications. One strategy to generate PEG scaffolds is to cross-link PEG chains to generate an insoluble, swollen polymer network
103 known as a hydrogel. A variety of approaches have been utilized to this end, including
radiation cross-linking, modification with fumarate, acrylate, or methacrylate groups
followed by free radical photopolymerization, and by linking sulfone terminated
multiarm PEG with disulfhydryl cross-linkers via Michael addition [60, 62-65].
In particular photopolymerization using a suitable free radical photoinitiator
allows hydrogels to be formed into a variety of shapes in situ under relatively mild
polymerization conditions in aqueous environments (i.e. compatible with living cells). In
the presence of a photoinitiator and a suitable light source, this type of polymerization
proceeds via a chemistry analogous to the free radical polymerization of any vinyl
monomer. However, the kinetics of this process are complex and have two distinct
phases because the viscous and diffusive properties of the system rapidly change after the
onset of gelation [66, 67]. Immediately after the onset of polymerization, the viscosity of
the solution rapidly increases resulting in a phenomenon known as autoacceleration, where diffusion of propagating centers, which act as terminators by coupling with other centers, becomes much slower than diffusion of smaller PEG-based macromers, leading
to an increased rate of reaction. As the network becomes more robust throughout the
polymerization eventually the diffusion of monomer also becomes limited, leading to a
decreased polymerization rate known as autodeceleration. Since this polymerization is
occurring at low concentrations in an aqueous environment exposed to atmospheric oxygen, other termination/inhibition processes are also likely affecting the
polymerization [68].
For diacrylated PEG macromers, which are most typically used, the net result of
this complicated polymerization process is the formation of an interconnected PEG
104 network illustrated in Fig. 3.3 (also see Chapter 4, Fig. 4.1). This network contains poly([PEG-esterified] acrylic acid) kinetic chains that form multifunctional cross-linking nodes that are bridged by PEG cross-link chains. There is some evidence that these cross-linking points aggregate into dense microdomains [69]. During polymerization,
PEG chains can entangle to form additional cross-link nodes. The network also contains other defects, including partially unreacted macromer and cyclic structures, that do not contribute to the effective network.
The structure of these materials dictates their swelling and mechanical properties as well as the ability of solute to diffuse within the hydrogel [70]. The properties of these networks can be modified by adjusting the conditions of polymerization, altering the concentration of PEGDA at the time of polymerization, adjusting the length of PEGDA, altering the functionality of the PEG cross-linker, copolymerizing PEGDA with other acrylate compounds, or by introducing non-reactive additives such as linear PEG [71-75].
For tissue engineering applications, the PEG length and the initial concentration of macromer are used most commonly to alter the network structure. Increasing the initial macromer concentration or decreasing PEG length results in higher cross-linking, decreased swelling, increased mechanical modulus, and decreased apparent diffusivity of molecules within the gels [72, 76-79]. These properties alone can affect the behavior of cells on or within hydrogel tissue engineering scaffolds comprised of PEGDA [76-79].
PEGDA hydrogels are inherently non-adhesive due to the resistance of the PEG chains to protein adsorption. Hern and Hubbell first reported the modification of PEGDA with the RGD adhesive peptide in 1998 and subsequently numerous studies have incorporated various cell adhesive ligands for a variety of cell types [47, 48, 60, 65, 80-
105
I A O I O D O O I O O O O O O O O O O O O n O O O O O I n O O C O O O T B O I O O O O O O O O O O O O O n O n O O O O T O O O O O O O
T Figure 3.3-Chemistry of free radical polymerization of PEGDA materials. Photoinitiation generates a radical, I, which initiates the polymerization (A). Polymerization (B) of acrylate groups proceeds until termination to generate a polyacrylate polymer node (C). Each of these nodes is linked to others via PEG chains connecting the reactive acrylate groups (D) forming a polymer network.
106 82]. In this approach, RGD peptides were tethered to the PEGDA network via a PEG-
monoacrylate (PEGMA) [60]. This strategy has also been utilized to link growth factors, inorganic compounds and other biologically functional moieties to PEGDA gels [49, 83-
87]. As a result, PEGDA-co-PEGMA hydrogels have been employed for a variety of applications including tissue engineered bone, cartilage, and blood vessels [48, 60, 81,
82, 88].
Because the mesh size of these gels is likely on the order of 10-100 nm (e.g. see
Chapter 6), large proteins and cells cannot readily penetrate the intact network [65, 72].
Therefore, to allow 3D SMC in-growth, PEGDA systems also must be degradable.
PEGDA hydrogels have been created that are hydrolytically [89, 90] or proteolytically degradable [48, 61, 91]. Proteolytically degradable gels inherently respond to local cellular action, limiting degradation to sites of cellular invasion and allowing the unpopulated matrix to remain intact for as long as is necessary for cells to invade it. Such gels typically contain a peptide along the PEG backbone of a PEGDA molecule that is sensitive to cleavage by selected matrix metalloproteinases. After cleavage, the PEGDA molecules eventually cease to form a polymer network. Matrix metalloproteinase-1
(MMP-1, Collagenase-1) has been well characterized, is expressed by SMCs, and does not degrade laminin or elastin [92, 93]. Therefore, peptide sequences sensitive to this enzyme are an excellent choice for incorporation into degradable PEGDA for vascular tissue engineering. Such peptides have been used effectively in other PEG-based gel systems to allow cell-mediated gel degradation [62].
107 3.8. Specific Aims
The overall goal of this work was to develop a fully synthetic, engineered, and
cell-instructive hydrogel biomaterials to guide the development of the vascular tunica
media from smooth muscle cells. Such a system would not be designed to act as a stand-
alone scaffold, but would require the parallel development of a load bearing support.
Initially, this support is envisioned to be ePTFE because this material is already in
clinical use. Synthetic SMCs, generated as a consequence of graft placement or ex vivo culture, contribute to failure due to SMC hyperplasia. A key challenge for vascular tissue engineering is to establish strategies to re-differentiate synthetic, proliferating SMCs
toward a contractile phenotype, indicative of the SMCs in the native tunica media.
The specific hypothesis of this thesis is that SMC phenotype can be modulated by
a cell-instructive hydrogel scaffold. This thesis had three specific aims to investigate this hypothesis (Fig. 3.4):
Aim 1: Quantify the properties of poly(ethylene glycol) diacrylate (PEGDA)-based hydrogels. PEGDA hydrogels are typically copolymerized with poly(ethylene) glycol
(PEG) that is heterofunctionalized with acrylate and bioactive moieties (PEG monoacrylate, PEGMA). The swelling, mechanical, and network characteristics of these
copolymer networks were examined over a wide range of compositions to establish
engineering control over the physical properties of the system.
Aim 2: Quantify the ability of PEG-based hydrogels modified with ECM-mimetic
peptides to support the contractile SMC phenotype. SMC differentiation toward a
contractile phenotype, driven by external stimulation, was quantified on PEGDA
hydrogels copolymerized with various pendant ECM-mimetic peptides and compared
108 with the phenotype of SMCs on whole ECM proteins that have been previously
characterized.
Aim 3: Engineer heparin-releasing hydrogel scaffolds to modulate SMC phenotype.
Heparin releasing, RGD bearing PEG-based hydrogels were developed as first generation cell-instructive scaffold materials to promote contractile SMC phenotype. Hydrogel design was engineered to provide optimized heparin release, and the effect of released heparin on SMC phenotype was quantified.
The work presented here on these specific aims has resulted in a synthetic, engineered scaffold for smooth muscle tissue that promotes contractile phenotype.
109
Characterize PEGDA-co-PEGMA Aim 1: hydrogel properties
Photoinitiator + Light
Quantify SMC phenotype on Aim 2: PEGDA-co-PEGMA hydrogels
Engineer scaffolds to promote Aim 3: contractile SMC phenotype
Heparin LEGEND: PEGDA “Synthetic” SMC PEGMA “Contractile” (w/ ligand) SMC
Figure 3.4-Schematic overview of project specific aims.
110 3.9. References
1. Langer R, Vacanti JP. Tissue Engineering. Science 1993;260(5110):920-926.
2. Gartner LP, Hiatt JL. Color atlas of histology. 4th ed. Philadelphia: Lippincott Williams & Wilkins, 2006.
3. Boron WF, Boulpaep EL. Medical physiology : a cellular and molecular approach. Updated ed. Philadelphia, Pa.: Elsevier Saunders, 2005.
4. Abbott WM, Callow A, Moore W, Rutherford R, Veith F, Weinberg S. Evaluation and performance standards for arterial prostheses. J Vasc Surg 1993;17(4):746-756.
5. Chan-Park MB, Shen JY, Cao Y, Xiong Y, Liu Y, Rayatpisheh S, Kang GC, Greisler HP. Biomimetic control of vascular smooth muscle cell morphology and phenotype for functional tissue-engineered small-diameter blood vessels. J Biomed Mater Res A 2009;88(4):1104-1121.
6. Laube HR, Duwe J, Rutsch W, Konertz W. Clinical experience with autologous endothelial cell-seeded polytetrafluoroethylene coronary artery bypass grafts. J Thorac Cardiovasc Surg 2000;120(1):134-141.
7. Meinhart JG, Deutsch M, Fischlein T, Howanietz N, Froschl A, Zilla P. Clinical autologous in vitro endothelialization of 153 infrainguinal ePTFE grafts. Ann Thorac Surg 2001;71(5 Suppl):S327-331.
8. Hehrlein FW, Schlepper M, Loskot F, Scheld HH, Walter P, Mulch J. The use of expanded polytetrafluoroethylene (PTFE) grafts for myocardial revascularization. J Cardiovasc Surg (Torino) 1984;25(6):549-553.
9. Motwani JG, Topol EJ. Aortocoronary saphenous vein graft disease: pathogenesis, predisposition, and prevention. Circulation 1998;97(9):916-931.
10. Sayers RD, Raptis S, Berce M, Miller JH. Long-term results of femorotibial bypass with vein or polytetrafluoroethylene. Br J Surg 1998;85(7):934-938.
11. Kidd KR, Patula VB, Williams SK. Accelerated endothelialization of interpositional 1-mm vascular grafts. J Surg Res 2003;113(2):234-242.
12. Larsen CC, Kligman F, Tang C, Kottke-Marchant K, Marchant RE. A biomimetic peptide fluorosurfactant polymer for endothelialization of ePTFE with limited platelet adhesion. Biomaterials 2007;28(24):3537-3548.
13. Conklin BS, Richter ER, Kreutziger KL, Zhong DS, Chen C. Development and evaluation of a novel decellularized vascular xenograft. Medical Engineering & Physics 2002;24(3):173-183.
111 14. Borschel GH, Huang YC, Calve S, Arruda EM, Lynch JB, Dow DE, Kuzon WM, Dennis RG, Brown DL. Tissue engineering of recellularized small-diameter vascular grafts. Tissue Engineering 2005;11(5-6):778-786.
15. Kaushal S, Amiel GE, Guleserian KJ, Shapira OM, Perry T, Sutherland FW, Rabkin E, Moran AM, Schoen FJ, Atala A, Soker S, Bischoff J, Mayer JE. Functional small- diameter neovessels created using endothelial progenitor cells expanded ex vivo. Nature Medicine 2001;7(9):1035-1040.
16. Cho SW, Lim SH, Kim IK, Hong YS, Kim SS, Yoo KJ, Park HY, Jang Y, Chang BC, Choi CY, Hwang KC, Kim BS. Small-diameter blood vessels engineered with bone marrow-derived cells. Annals of Surgery 2005;241(3):506-515.
17. Cho SW, Kim IK, Kang JM, Song KW, Kim HS, Park CH, Yoo KJ, Kim BS. Evidence for in vivo growth potential and vascular remodeling of tissue-engineered artery. Tissue Eng Part A 2009;15(4):901-912.
18. Schaner PJ, Martin ND, Tulenko TN, Shapiro IM, Tarola NA, Leichter RF, Carabasi RA, DiMuzio PJ. Decellularized vein as a potential scaffold for vascular tissue engineering. Journal of Vascular Surgery 2004;40(1):146-153.
19. Daniel J, Abe K, McFetridge PS. Development of the human umbilical vein scaffold for cardiovascular tissue engineering applications. Asaio Journal 2005;51(3):252- 261.
20. Huynh T, Abraham G, Murray J, Brockbank K, Hagen PO, Sullivan S. Remodeling of an acellular collagen graft into a physiologically responsive neovessel. Nature Biotechnology 1999;17(11):1083-1086.
21. L'Heureux N, Paquet S, Labbe R, Germain L, Auger FA. A completely biological tissue-engineered human blood vessel. Faseb Journal 1998;12(1):47-56.
22. L'Heureux N, Dusserre N, Konig G, Victor B, Keire P, Wight TN, Chronos NA, Kyles AE, Gregory CR, Hoyt G, Robbins RC, McAllister TN. Human tissue- engineered blood vessels for adult arterial revascularization. Nat Med 2006;12(3):361-365.
23. McAllister TN, Maruszewski M, Garrido SA, Wystrychowski W, Dusserre N, Marini A, Zagalski K, Fiorillo A, Avila H, Manglano X, Antonelli J, Kocher A, Zembala M, Cierpka L, de la Fuente LM, L'Heureux N. Effectiveness of haemodialysis access with an autologous tissue-engineered vascular graft: a multicentre cohort study. The Lancet 2009;373(9673):1440-1446.
24. Weinberg CB, Bell E. A blood vessel model constructed from collagen and cultured vascular cells. Science 1986;231(4736):397-400.
112 25. Tranquillo RT, Girton TS, Bromberek BA, Triebes TG, Mooradian DL. Magnetically orientated tissue-equivalent tubes: Application to a circumferentially orientated media-equivalent. Biomaterials 1996;17(3):349-357.
26. Salacinski HJ, Goldner S, Giudiceandrea A, Hamilton G, Seifalian AM, Edwards A, Carson RJ. The mechanical behavior of vascular grafts: a review. J Biomater Appl 2001;15(3):241-278.
27. Walden R, L'Italien GJ, Megerman J, Abbott WM. Matched elastic properties and successful arterial grafting. Arch Surg 1980;115(10):1166-1169.
28. Girton TS, Oegema TR, Tranquillo RT. Exploiting glycation to stiffen and strengthen tissue equivalents for tissue engineering. Journal of Biomedical Materials Research 1999;46(1):87-92.
29. Seliktar D, Black RA, Vito RP, Nerem RM. Dynamic mechanical conditioning of collagen-gel blood vessel constructs induces remodeling in vitro. Ann Biomed Eng 2000;28(4):351-362.
30. Seliktar D, Nerem RM, Galis ZS. Mechanical strain-stimulated remodeling of tissue- engineered blood vessel constructs. Tissue Engineering 2003;9(4):657-666.
31. Stegemann JP, Nerem RM. Phenotype modulation in vascular tissue engineering using biochemical and mechanical stimulation. Ann Biomed Eng 2003;31(4):391- 402.
32. Grassl ED, Oegema TR, Tranquillo RT. A fibrin-based arterial media equivalent. Journal of Biomedical Materials Research Part A 2003;66A(3):550-561.
33. Swartz DD, Russell JA, Andreadis ST. Engineering of fibrin-based functional and implantable small-diameter blood vessels. American Journal of Physiology-Heart and Circulatory Physiology 2005;288(3):H1451-H1460.
34. Liu JY, Swartz DD, Peng HF, Gugino SF, Russell JA, Andreadis ST. Functional tissue-engineered blood vessels from bone marrow progenitor cells. Cardiovasc Res 2007;75(3):618-628.
35. Zavan B, Vindigni V, Lepidi S, Iacopetti I, Avruscio G, Abatangelo G, Cortivo R. Neoarteries grown in vivo using a tissue-engineered hyaluronan-based scaffold. Faseb Journal 2008;22(8):2853-2861.
36. Niklason LE, Gao J, Abbott WM, Hirschi KK, Houser S, Marini R, Langer R. Functional arteries grown in vitro. Science 1999;284(5413):489-493.
37. Shum-Tim D, Stock U, Hrkach J, Shinoka T, Lien J, Moses MA, Stamp A, Taylor G, Moran AM, Landis W, Langer R, Vacanti JP, Mayer JE. Tissue engineering of autologous aorta using a new biodegradable polymer. Ann Thorac Surg 1999;68(6):2298-2304.
113 38. Teebken OE, Pichlmaier AM, Haverich A. Cell seeded decellularised allogeneic matrix grafts and biodegradable polydioxanone-prostheses compared with arterial autografts in a porcine model. Eur J Vasc Endovasc Surg 2001;22(2):139-145.
39. Matsumura G, Hibino N, Ikada Y, Kurosawa H, Shin'oka T. Successful application of tissue engineered vascular autografts: clinical experience. Biomaterials 2003;24(13):2303-2308.
40. Shin'oka T, Matsumura G, Hibino N, Naito Y, Watanabe M, Konuma T, Sakamoto T, Nagatsu M, Kurosawa H. Midterm clinical result of tissue-engineered vascular autografts seeded with autologous bone marrow cells. J Thorac Cardiovasc Surg 2005;129(6):1330-1338.
41. Sill TJ, von Recum HA. Electrospinning: applications in drug delivery and tissue engineering. Biomaterials 2008;29(13):1989-2006.
42. Lee SJ, Liu J, Oh SH, Soker S, Atala A, Yoo JJ. Development of a composite vascular scaffolding system that withstands physiological vascular conditions. Biomaterials 2008;29(19):2891-2898.
43. Tillman BW, Yazdani SK, Lee SJ, Geary RL, Atala A, Yoo JJ. The in vivo stability of electrospun polycaprolactone-collagen scaffolds in vascular reconstruction. Biomaterials 2009;30(4):583-588.
44. Stitzel J, Liu L, Lee SJ, Komura M, Berry J, Soker S, Lim G, Van Dyke M, Czerw R, Yoo JJ, Atala A. Controlled fabrication of a biological vascular substitute. Biomaterials 2006;27(7):1088-1094.
45. Hashi CK, Zhu YQ, Yang GY, Young WL, Hsiao BS, Wang K, Chu B, Li S. Antithrombogenic property of bone marrow mesenchymal stem cells in nanofibrous vascular grafts. Proceedings of the National Academy of Sciences of the United States of America 2007;104(29):11915-11920.
46. Pektok E, Nottelet B, Tille JC, Gurny R, Kalangos A, Moeller M, Walpoth BH. Degradation and Healing Characteristics of Small-Diameter Poly(epsilon- Caprolactone) Vascular Grafts in the Rat Systemic Arterial Circulation. Circulation 2008;118(24):2563-2570.
47. Gobin AS, West JL. Val-ala-pro-gly, an elastin-derived non-integrin ligand: smooth muscle cell adhesion and specificity. J Biomed Mater Res A 2003;67(1):255-259.
48. Mann BK, Gobin AS, Tsai AT, Schmedlen RH, West JL. Smooth muscle cell growth in photopolymerized hydrogels with cell adhesive and proteolytically degradable domains: synthetic ECM analogs for tissue engineering. Biomaterials 2001;22(22):3045-3051.
114 49. Mann BK, Schmedlen RH, West JL. Tethered-TGF-beta increases extracellular matrix production of vascular smooth muscle cells. Biomaterials 2001;22(5):439- 444.
50. Mann BK, Tsai AT, Scott-Burden T, West JL. Modification of surfaces with cell adhesion peptides alters extracellular matrix deposition. Biomaterials 1999;20(23- 24):2281-2286.
51. Beamish JA, Fu A, Choi A, Haq N, Kottke-Marchant K, Marchant RE. The influence of RGD-bearing hydrogels on the re-expression of contractile vascular smooth muscle cell phenotype. Biomaterials 2009;in press.
52. Beamish JA, Zhu J, Kottke-Marchant K, Marchant RE. The effects of monoacrylated poly(ethylene glycol) on the properties of poly(ethylene glycol) diacrylate hydrogels used for tissue engineering. J Biomed Mater Res 2009;in press.
53. Hahn MS, McHale MK, Wang E, Schmedlen RH, West JL. Physiologic pulsatile flow bioreactor conditioning of poly(ethylene glycol)-based tissue engineered vascular grafts. Ann Biomed Eng 2007;35(2):190-200.
54. Bulick AS, Munoz-Pinto DJ, Qu X, Mani M, Cristancho D, Urban M, Hahn MS. Impact of endothelial cells and mechanical conditioning on smooth muscle cell extracellular matrix production and differentiation. Tissue Eng Part A 2009;15(4):815-825.
55. Poh M, Boyer M, Solan A, Dahl SLM, Pedrotty D, Banik SSR, McKee JA, Klinger RY, Counter CM, Niklason LE. Blood vessels engineered from human cells. Lancet 2005;365(9477):2122-2124.
56. Liu JY, Peng HF, Andreadis ST. Contractile smooth muscle cells derived from hair- follicle stem cells. Cardiovasc Res 2008;79(1):24-33.
57. Zilla P, Bezuidenhout D, Human P. Prosthetic vascular grafts: wrong models, wrong questions and no healing. Biomaterials 2007;28(34):5009-5027.
58. Almany L, Seliktar D. Biosynthetic hydrogel scaffolds made from fibrinogen and polyethylene glycol for 3D cell cultures. Biomaterials 2005;26(15):2467-2477.
59. Halstenberg S, Panitch A, Rizzi S, Hall H, Hubbell JA. Biologically engineered protein-graft-poly(ethylene glycol) hydrogels: A cell adhesive and plasm in- degradable biosynthetic material for tissue repair. Biomacromolecules 2002;3(4):710-723.
60. Hern DL, Hubbell JA. Incorporation of adhesion peptides into nonadhesive hydrogels useful for tissue resurfacing. J Biomed Mater Res 1998;39(2):266-276.
61. West JL, Hubbell JA. Polymeric biomaterials with degradation sites for proteases involved in cell migration. Macromolecules 1999;32(1):241-244.
115 62. Lutolf MP, Lauer-Fields JL, Schmoekel HG, Metters AT, Weber FE, Fields GB, Hubbell JA. Synthetic matrix metalloproteinase-sensitive hydrogels for the conduction of tissue regeneration: engineering cell-invasion characteristics. Proc Natl Acad Sci U S A 2003;100(9):5413-5418.
63. Merrill EW, Dennison KA, Sung C. Partitioning and diffusion of solutes in hydrogels of poly(ethylene oxide). Biomaterials 1993;14(15):1117-1126.
64. Lutolf MP, Hubbell JA. Synthesis and physicochemical characterization of end- linked poly(ethylene glycol)-co-peptide hydrogels formed by Michael-type addition. Biomacromolecules 2003;4(3):713-722.
65. Zhu JM, Beamish JA, Tang C, Kottke-Marchant K, Marchant RE. Extracellular matrix-like cell-adhesive hydrogels from RGD-containing poly(ethylene glycol) diacrylate. Macromolecules 2006;39(4):1305-1307.
66. Anseth KS, Kline LM, Walker TA, Anderson KJ, Bowman CN. Reaction-Kinetics and Volume Relaxation during Polymerizations of Multiethylene Glycol Dimethacrylates. Macromolecules 1995;28(7):2491-2499.
67. Burdick JA, Lovestead TM, Anseth KS. Kinetic chain lengths in highly cross-linked networks formed by the photoinitiated polymerization of divinyl monomers: A gel permeation chromatography investigation. Biomacromolecules 2003;4(1):149-156.
68. Kurdikar DL, Peppas NA. A Kinetic-Study of Diacrylate Photopolymerizations. Polymer 1994;35(5):1004-1011.
69. Lin-Gibson S, Jones RL, Washburn NR, Horkay F. Structure-property relationships of photopolymerizable poly(ethylene glycol) dimethacrylate hydrogels. Macromolecules 2005;38(7):2897-2902.
70. Peppas NA. Hydrogels in medicine and pharmacy. Boca Raton, Fla.: CRC Press, 1986.
71. Buxton AN, Zhu J, Marchant R, West JL, Yoo JU, Johnstone B. Design and characterization of poly(ethylene glycol) photopolymerizable semi-interpenetrating networks for chondrogenesis of human mesenchymal stem cells. Tissue Eng 2007;13(10):2549-2560.
72. Cruise GM, Scharp DS, Hubbell JA. Characterization of permeability and network structure of interfacially photopolymerized poly(ethylene glycol) diacrylate hydrogels. Biomaterials 1998;19(14):1287-1294.
73. Padmavathi NC, Chatterji PR. Structural characteristics and swelling behavior of poly(ethylene glycol) diacrylate hydrogels. Macromolecules 1996;29(6):1976-1979.
74. Keys KB, Andreopoulos FM, Peppas NA. Poly(ethylene glycol) star polymer hydrogels. Macromolecules 1998;31(23):8149-8156.
116 75. Ravi N, Mitra A, Hamilton P, Horkay F. Characterization of the network properties of poly(ethylene glycol)-acrylate hydrogels prepared by variations in the ethanol- water solvent composition during crosslinking copolymerization. Journal of Polymer Science Part B-Polymer Physics 2002;40(23):2677-2684.
76. Peyton SR, Raub CB, Keschrumrus VP, Putnam AJ. The use of poly(ethylene glycol) hydrogels to investigate the impact of ECM chemistry and mechanics on smooth muscle cells. Biomaterials 2006;27(28):4881-4893.
77. Bryant SJ, Bender RJ, Durand KL, Anseth KS. Encapsulating chondrocytes in degrading PEG hydrogels with high modulus: engineering gel structural changes to facilitate cartilaginous tissue production. Biotechnol Bioeng 2004;86(7):747-755.
78. Bryant SJ, Nuttelman CR, Anseth KS. The effects of crosslinking density on cartilage formation in photocrosslinkable hydrogels. Biomed Sci Instrum 1999;35:309-314.
79. Gunn JW, Turner SD, Mann BK. Adhesive and mechanical properties of hydrogels influence neurite extension. J Biomed Mater Res A 2005;72(1):91-97.
80. Benoit DS, Anseth KS. The effect on osteoblast function of colocalized RGD and PHSRN epitopes on PEG surfaces. Biomaterials 2005;26(25):5209-5220.
81. Burdick JA, Anseth KS. Photoencapsulation of osteoblasts in injectable RGD- modified PEG hydrogels for bone tissue engineering. Biomaterials 2002;23(22):4315-4323.
82. Nuttelman CR, Tripodi MC, Anseth KS. In vitro osteogenic differentiation of human mesenchymal stem cells photoencapsulated in PEG hydrogels. J Biomed Mater Res A 2004;68(4):773-782.
83. Gobin AS, West JL. Effects of epidermal growth factor on fibroblast migration through biomimetic hydrogels. Biotechnol Prog 2003;19(6):1781-1785.
84. DeLong SA, Moon JJ, West JL. Covalently immobilized gradients of bFGF on hydrogel scaffolds for directed cell migration. Biomaterials 2005;26(16):3227-3234.
85. Nuttelman CR, Benoit DS, Tripodi MC, Anseth KS. The effect of ethylene glycol methacrylate phosphate in PEG hydrogels on mineralization and viability of encapsulated hMSCs. Biomaterials 2006;27(8):1377-1386.
86. Cushing MC, Liao JT, Jaeggli MP, Anseth KS. Material-based regulation of the myofibroblast phenotype. Biomaterials 2007;28(23):3378-3387.
87. Benoit DS, Nuttelman CR, Collins SD, Anseth KS. Synthesis and characterization of a fluvastatin-releasing hydrogel delivery system to modulate hMSC differentiation and function for bone regeneration. Biomaterials 2006;27(36):6102-6110.
117 88. Hwang NS, Varghese S, Zhang Z, Elisseeff J. Chondrogenic differentiation of human embryonic stem cell-derived cells in arginine-glycine-aspartate-modified hydrogels. Tissue Eng 2006;12(9):2695-2706.
89. Sawhney AS, Pathak CP, Hubbell JA. Bioerodible Hydrogels Based on Photopolymerized Poly(Ethylene Glycol)-Co-Poly(Alpha-Hydroxy Acid) Diacrylate Macromers. Macromolecules 1993;26(4):581-587.
90. Mahoney MJ, Anseth KS. Three-dimensional growth and function of neural tissue in degradable polyethylene glycol hydrogels. Biomaterials 2006;27(10):2265-2274.
91. Gobin AS, West JL. Cell migration through defined, synthetic ECM analogs. Faseb J 2002;16(7):751-753.
92. Newby AC. Matrix metalloproteinases regulate migration, proliferation, and death of vascular smooth muscle cells by degrading matrix and non-matrix substrates. Cardiovascular Research 2006;69(3):614-624.
93. Parks WC, Mecham RP. Matrix metalloproteinases. San Diego: Academic Press, 1998.
118 CHAPTER 4: The Effects of Monoacrylated Poly(Ethylene Glycol) on the Properties of Poly(Ethylene Glycol) Diacrylate Hydrogels Used for Tissue Engineering
Based on: Beamish JA, Zhu J, Kottke-Marchant K, Marchant RE. The effects of monoacrylated poly(ethylene glycol) on the properties of poly(ethylene glycol) diacrylate hydrogels used for tissue engineering. J Biomed Mater Res 2009; in press.
4.1. Introduction
Poly(ethylene glycol) (PEG)-based hydrogels have attracted broad interest as a
scaffold material for tissue engineering applications [1-3]. The PEG backbone of these
materials resists the adsorption of exogenous proteins and does not support cell
attachment. However, the inclusion of biological ligands allows cell interactions with the
scaffold that can be controlled quantitatively. PEG networks have been formed using a
variety of approaches including radiation cross-linking, modification with fumarate,
acrylate or methacrylate groups followed by free radical photopolymerization, and by
linking sulfone terminated multiarm PEG with disulfhydryl cross-linkers via Michael
addition [1-4]. Photopolymerization, including photolithography, of PEG
di(meth)acrylate (PEGDA) has been employed specifically for tissue engineering
applications because these gels can be formed readily in situ [5]. This allows scaffolds to
be engineered in a variety of 3-dimensional structures and with gradients of biological
agents. Proteolytically or hydrolytically degradable PEGDA derivatives have been
synthesized to enhance the utility of this system as a scaffold material further [6, 7].
Acrylation followed by photoinitiated chain polymerization also has been used widely to
form networks from a variety of biological and synthetic materials used for tissue
engineering applications [8-11].
119 The network structure of PEGDA hydrogels is distinct from traditional polymer
networks that are formed by cross-linking preformed polymer chains or by
copolymerizing monomer with a small fraction of a cross-linker. PEGDA hydrogels are
formed typically from a single component that serves both as cross-linker and cross-
linked chain. The resulting network consists of PEG chains esterified to dense,
multifunctional cross-link regions formed from long poly(acrylic acid) kinetic chains
[12]. Physical chain entanglements also contribute to the network (Fig. 4.1). It is also
possible for PEGDA molecules to form intra-chain cycles and dangling ends, but these do
not contribute to the network-dependent properties.
Numerous studies have attempted to develop effective strategies to control
PEGDA gel properties, including increasing the ratio of PEGDA to water at the time of
polymerization or decreasing the PEG chain length. These approaches result in
decreased hydrogel swelling [13-15], increased mechanical modulus [14, 15], and
decreased effective transport of non-gaseous molecules diffusing through the gel [13].
Alterations in PEGDA concentration or size alter the kinetics of polymerization, which
affect the chain length of the poly(acrylic acid) cross-linking nodes and the extent of
network defects such as intra-chain cyclic structures or unreacted PEGDA. Many studies
also have attempted to estimate the molecular weight between cross-links (Mc) and the
mesh size (ε) from swelling and mechanical data [6, 13, 14] using variations on the Flory-
Rehner model [16]. These models provide estimates that generally correlate well with
observed physical properties. However, by using equations derived for networks formed
from long polymers with tetrafunctional junctions [13] these analyses neglect the complex nature of the PEGDA network.
120 When used as a tissue engineering scaffold, the physical properties of PEGDA hydrogel scaffolds can alter the behavior of a variety of tissues including cartilage, smooth muscle, and nerve. Increasing the initial macromer concentration or decreasing
PEG length results in altered gel properties that affect cell behavior directly including diminished access to proteins in the extracellular space [13], altered extracellular matrix production and cell phenotype [6, 17], or decreased neurite extension [18]. The physical properties of a tissue engineering scaffold can also limit the diffusion of nutrients and growth factors and can alter the rate of degradation of degradable PEGDA derivatives.
Without modification, PEGDA hydrogels do not support cell attachment [19].
Since Hern and Hubbell first reported the modification of PEGDA gels with the RGD adhesive peptide, numerous studies have developed this system as a scaffold for a variety of cell types [3, 7, 19-21]. In these studies, adhesive peptides were linked to the PEGDA network via PEG-monoacrylates (PEGMA) which participate in the free radical cross- linking of the gel, resulting in non-network dangling ends (Fig 4.1.). This strategy has also been utilized to link growth factors, inorganic compounds and other biologically functional moieties to PEGDA gels [22-25]. PEGMA conjugates also have been copolymerized with PEGDA to impart pH sensitivity to gel swelling or to alter the network properties for non-tissue engineering purposes [14, 26]. Despite the wealth of applications, the effects of PEGMA on PEGDA network structure and gel properties have not been studied in detail. Understanding the effect of incorporating biologically active
PEGMA will facilitate the design of PEGDA scaffolds which provide optimal biochemical and mechanical environments. In this report, we examine the effect of
121
Figure 4.1-Schematic representation of the PEGDA network showing (a) cross-linked PEG chains and network defects including (b) unreacted acrylate moieties, (c) PEG cycles, (d) chain entanglements, and (e) PEG tethered biological moieties. Shaded arrow shapes represent reacted acrylates in a poly(acrylic acid) cross-linking node, unshaded arrow shapes represent unreacted acrylate, dark lines represent PEG chains esterified to the acrylic acid, and the shaded hexagon represents a biological moieity. For clarity, short poly(acrylic acid) chains are shown, but in actual gels these chain lengths may be much longer.
122 PEGMA on the network structure and physical properties of PEGDA hydrogels generated from macromer materials similar to those used for tissue engineering.
4.2. Materials and Methods
4.2.1. Materials
Poly(ethylene glycol) (PEG, Mn 6000), monomethoxy-PEG (Mn 2000 or 5000), triethylamine, acryloyl chloride, and anhydrous dichloromethane were purchased from
Aldrich and used as received. Irgacure 2959 (1-[4-[2-Hydroxyethoxy]-phenyl]-2- hydroxy-2-methyl-1-propane-1-one) was purchased from Ciba Specialty Chemicals
(Tarrytown, NY). Phosphate buffered saline (PBS) was prepared in distilled water and adjusted to pH 7.4. Chemical structures of relevant compounds are shown in Table 4.1.
4.2.2. Synthesis of Hydrogel Precursors
PEGDA (Mn ≈ 6000) and PEGMA (Mn ≈ 2000 or 5000) were prepared by conjugation of PEG-diol or monomethoxy-PEG-OH, respectively, with a 2-fold molar excess of acryloyl chloride relative to free hydroxyls in the presence of triethylamine in anhydrous dichloromethane under nitrogen overnight. The resulting products were precipitated in ice cold ether, filtered, dried in a vacuum oven at room temperature, and stored at -20 ºC. The resulting product was characterized by matrix assisted laser desorption/ionization time of flight mass spectroscopy (MALDI-MS) and proton NMR.
4.2.3. Hydrogel Formation
PEGDA-co-PEGMA hydrogels were formed at various compositions (PEGMA: 0-20 % w/w, PEGDA: 0-20% w/w, PBS: 60-95% w/w) using 0.1% (w/v) Irgacure 2959 photoinitiator. Hydrogel precursors were formed from stock solutions of PEGDA (44% w/w), PEGMA (44% w/w), PBS, and Irgacure 2959 (1% w/v in PBS). The density of
123
Table 4.1-Chemical Structures of PEGDA and PEGMA
Chemical Structure poly(ethylene glycol) diacrylate (PEGDA, MW = 6 kD) poly(ethylene glycol) monoacrylate methyl ether (PEGMA, MW = 5 kD)
124 PEG solutions, which is linear and independent of MW for 2-12 kD PEG in this
composition range, was used to adjust the volumes of each solution to achieve the
appropriate final mass composition for each hydrogel. Each precursor solution (100 μl)
was dispensed into a stainless steel mold (D = 10 mm, H = 1.2 mm) and polymerized for
10.0 min at 0.4-0.5 mW/cm2 under a 365 nm UV lamp at room temperature. Three gels
were formed for each composition studied.
4.2.4. Properties Testing
Hydrogels were allowed to swell in excess PBS (pH 7.4) at room temperature (22-
23 ºC) for 4 days. A 10 mm circle was punched from the center of each swollen gel,
massed, and mounted between 10 mm circular compression platens in a Dynamic
Mechanical Analyzer 7e (Perkin-Elmer). The mechanical properties of each gel were
evaluated in unconfined compression in a PBS bath at a rate of 50 mN/min. The shear
modulus was determined from the slope of σ vs. –(λ-λ-2) where λ is the ratio of the
deformed height of the gel to the pre-deformation height[14]. The linear region used to
determine the shear modulus was typically between 2 and 25% compressive strain. Non-
linear regions of the σ vs. –(λ-λ-2) plot for small values of –(λ-λ-2) (during initial platen- gel contact) were excluded from the analysis. For the strongest and weakest gels, the tests were repeated after 1 day of re-equilibration with PBS at a rate of 25 mN/min to assess the effects of stress-rate on the measured modulus.
After mechanical testing, each gel was incubated with excess distilled water for an additional day to leach buffer salts and lyophilized. The mass swelling ratio, q, was determined using the following equation:
m q = s (1) mp
125
where ms and mp are the mass of the swollen gel in PBS and the mass of the polymer network, respectively.
4.2.5. Determination of minimal PEGDA concentration for gel formation
To investigate the properties of PEGDA-co-PEGMA gels with minimal PEGDA
content, hydrogels with PEGMA (20% w/w) were polymerized with a range of low
concentrations of PEGDA (0.1-5%, w/w) as described above. The mass swelling ratio
was determined gravimetrically.
4.2.6. Hydrogel Network Composition
After testing gel properties, selected networks were degraded in 1 N sodium
hydroxide, dialyzed (500 MWCO), and lyophilized yielding samples containing monomethoxy-PEG (mPEG5k) and diol-PEG (PEG6k) corresponding to polymerized
PEGMA and PEGDA, respectively. Relative PEGMA vs. PEGDA incorporation was determined using the relative size of the mPEG5k vs. PEG6k distributions on MALDI-
MS spectra of the degraded gel samples. Spectra of samples with known
mPEG5k/PEG6k compositions were used to form a linear calibration curve (r2 > 0.99).
4.2.7. Analysis of Hydrogel Network Structure
Since the hydrogels studied here were formed in solution, the method of Bray and
Merrill was adapted to calculate ve, the effective cross-linked chains per gram of polymer, using the following equation [27]:
2 ln( 1−++vvps,,) psχ v ps , v = (2) e ⎛⎞1 ⎡ ⎜⎟ ⎤ ⎢⎛⎞vv⎝⎠3 2 ⎛⎞⎥ −−Vvρ ⎜⎟ps,, ⎜⎟ ps sppr, ⎢⎜⎟ ⎜⎟⎥ ⎢⎝⎠vFvpr,, ⎝⎠ pr ⎥ ⎣ ⎦
126
where:
1 v = (3) ps, ρ p ()q −11+ ρs
1 vpr, = (4) ρ p ⎛⎞1 ⎜⎟−11+ ρs ⎝⎠f
and χ is the Flory-Huggins interaction parameter for PEG and water (0.426) [4], Vs is the
3 -1 molar volume of the solvent, water (18.0 cm mol ), ρ p is the bulk density of PEG (1.18
-1 -1 g ml ) [14], F is the junction functionality, ρs is the density of PBS (1.01 g ml ), and f is
the mass fraction of polymer in solution at the time of cross-linking.
The PEGMA-co-PEGDA hydrogels contained various compositions of cross- linking (PEGDA) and non-cross-linking elements (PEGMA). To characterize the nature of these networks, the number of effective cross-links was normalized to the number of cross-link forming PEGDA molecules in each gel at the time of cross-linking to give a
* new quantity, ve defined as:
* f vve =⋅e ⋅ MWPEGDA (5) fPEGDA
where fPEGDA is the mass fraction of PEGDA at the time of polymerization and MWPEGDA
is the number averaged molecular weight of the PEGDA. In a perfect network with no
* entanglements, ve is equal to 1 since all of the PEGDA chains participate in the network.
* Because F is not explicitly determined by the chemical structure of the gels, ve was fit to
swelling data for 4 < F ≤ 106.
127 The network structure was also calculated independently from mechanical data using an affine model of rubber elasticity for isovolumetric compression [16]. After normalizing to the number of PEGDA molecules, the relationship between the shear
* modulus and ve was:
* GMW⋅ PEGDA ve = (6) 6 ⎛⎞fPEGDA RTvpp()ρ ⋅10 ⎜⎟ ⎝⎠f where G is the shear modulus (in N m-2), R is the gas constant (8.413 J K-1mol-1), and T is
-3 -1 the absolute temperature (K), and the units of ρp and MWPEGDA are g cm and g mol , respectively. The 106 term converts density from g cm-3 to g m-3.
4.2.8. Photopolymerization of PEGMA
Poly(PEGMA) homopolymers were prepared by polymerizing various concentrations of PEGMA (Mn = 2 kD, 10-50 % w/w) as described for the PEGDA-co-
PEGMA hydrogels. After polymerization the viscous liquids were transferred from the mold and lyophilized.
4.2.9. Gel Permeation Chromatography
Lyophilized poly(PEGMA) was reconstituted at 5 mg/ml in deionized water and filtered through a 0.22 μm filter. GPC was performed using 3 HEMA BIO 100 (Tessek
SEPARO) size exclusion columns in series with a flow rate of 1 ml/min and 100 μl sample volumes. Light scattering and refractive index measurements were acquired using a Waters 2690 separations module with a Waters 410 refractive index detector and a Wyatt MiniDAWN light scattering detector. The software package associated with the system (ASTRA, version 4.0) was used to calculate the molecular weight (Mn and Mw) of the PEGMA homopolymers using a dn/dc for PEG of 0.1360. The extent of
128 polymerization was determined by dividing Mn of the homopolymer by the Mn of the
PEGMA (2 kD). 2 kD PEGMA was substituted for 5 kD PEGMA to bring the molecular weight of the resulting branched polymer into a range suitable for our separation columns.
4.2.10. Statistics
Data are represented as mean ± standard deviation of at least 3 samples. Statistical analysis was performed using student’s t-tests for single comparisons. A p value < 0.05 was considered significant.
4.3. Results
4.3.1. PEGDA, PEGMA Synthesis
MALDI-MS spectra of PEGDA and PEGMA showed a shift to higher molecular weight of roughly 108 and 54 respectively corresponding to the conjugation of two or one acrylate groups. The PEGMA spectra showed two families of peaks consistent with the presence of acrylated and free hydroxyl PEGs and did not show evidence for the presence of any diacrylated PEG. Proton NMR demonstrated approximately 75% and 95% acrylation for 2 kD PEGMA and 5 kD PEGMA, respectively.
4.3.2. Hydrogel Network Composition
The recovered mass of the copolymer hydrogel networks increased proportionally with PEG concentration in the initial macromer solution (PEGMA+PEGDA) with no dependence on the PEGMA:PEGDA ratio, suggesting that the incorporation of PEGMA and PEGDA approximately mirrored the composition of the initial macromer solution.
The overall conversion increased from 78±5% toward 100% as the PEG concentration increased from 5 to 40% w/w. MALDI-MS analysis of selected
129 10%PEGDA/10%PEGMA and 10%PEGDA/20%PEGMA gels revealed estimated fractions of PEGMA in the final polymer network of 0.57±0.01 and 0.71±0.004 respectively.
4.3.3. Hydrogel Swelling
For fixed concentrations of PEG (PEGDA+PEGMA), increasing the mol fraction of PEGDA in the macromer feed resulted in a decreased swelling ratio, as expected (Fig
4.2A). The swelling ratio of PEGDA-co-PEGMA hydrogels decreased substantially with increasing PEGDA composition (Fig. 4.3A). The substitution of polymerization-inactive
PBS with polymerization-active PEGMA decreased the swelling ratio by up to 42±1.6%.
The effect was most pronounced for gels with low PEGDA compositions. PEGDA gels used for tissue engineering applications are often supplemented with biologically active
PEGMA. The addition of PEGMA (shown in units of mM) to gels with fixed PEGDA composition resulted in a decreased swelling ratio only when the concentration of
PEGMA was high relative to the concentrations used to promote cell attachment (< 7 mM) and the PEGDA concentration was low (Fig. 4.3B)[3, 19, 28].
As the concentration of cross-linking PEGDA was decreased in PEGDA-co-
PEGMA gels containing fixed PEGMA (5 kD, 20% w/w), the swelling ratio of the resulting gels increased asymptotically (Fig. 4.4). A minimum concentration of 1%
PEGDA (1:20 PEGDA:PEGMA) was necessary to induce gel formation. Controls formed from 0% PEGDA/20% PEGMA did not form gel networks.
4.3.4. Hydrogel Mechanical Properties
The shear modulus of the gels was calculated from the slope of σ vs. -(λ-λ-2). The value of r2 was greater than 0.988 for all regression fits. For fixed concentrations of PEG
130
Figure 4.2-Mass swelling ratios (A) and shear moduli (B) of PEGDA-co-PEGMA hydrogels shown as a function of the mole fraction of PEGDA in the macromer feed (mol PEGDA/[mol PEGDA+ mol PEGMA]). Each curve shows a fixed overall concentration of PEG in PBS (% PEGDA + % PEGMA, w/w) at the time of polymerization.
131
Figure 4.3-Mass swelling ratios of PEGDA-co-PEGMA hydrogels shown as a function of (A) the concentration of PEGDA (% w/w) at the time of cross-linking supplemented with various concentrations of PEGMA (% w/w), or (B) the molar concentration of PEGMA (mM) in the solution at the time of cross-linking for various PEGDA compositions (% w/w). PEGMA concentration was expressed in millimoles per liter to facilitate comparison with the tissue engineering literature.
Figure 4.4-Mass swelling ratio of PEGDA-co-PEGMA hydrogels formed from 20% PEGMA (w/w) and decreasing amounts of PEGDA. The shaded area indicates PEGDA compositions which did not support the formation of networks. Hydrogels formed from PEGDA alone are shown for comparison.
132 (PEGDA+PEGMA), increasing the mol fraction of PEGDA in the macromer feed resulted in an increased shear modulus, as expected (Fig 4.2B). The shear modulus increased with increasing PEGDA composition (Fig 4.5). The addition of PEGMA generally resulted in an increase in shear modulus that was proportionately similar, relative to the 0% PEGMA, for each PEGDA concentration studied (up to 167±29.3%).
After one day of re-equilibration in PBS, a reduction in the rate of stress application from 50 mN/min to 25 mN/min resulted in a decrease in the apparent modulus from 4.0±0.6 kPa to 3.4±0.3 kPa (14.7%, p=0.047) for the 5% PEGDA/0%
PEGMA gels and from 104.2±4.0 kPa to 102.6±18.5 kPa (1.6 %, p=0.861) for the 20%
PEGDA/20% PEGMA gels.
4.3.5. Hydrogel Network Structure
* The effective number of cross-linked chains per PEGDA molecule, ve , calculated with Eq. 5, was used to estimate the network structure from swelling data. A sensitivity
* * analysis on the effect of the unknown parameter F on ve showed that ve decreased asymptotically with increasing F, approaching a value 20-40% less than the F=4 calculation for PEGDA gels containing 0% PEGMA (Fig. 4.6). The results for hydrogels containing 5-20% (w/w) PEGMA followed a similar trend (not shown).
* To assess the network structure of PEGDA-co-PEGMA gels, ve was calculated from both swelling and mechanical data using F=4 (Figs. 4.7 and 4.8, respectively). The
* value of ve was directly proportional to the PEGDA composition (Figs. 4.7A and 4.8A).
* The addition of PEGMA increased ve while having a minimal impact on the slope of the
* ve vs. PEGDA relationship. The addition of PEGMA to gels with fixed PEGDA
* * composition resulted in an increase in ve (Figs. 4.7B and 4.8B). ve estimated from
133
Figure 4.5-Shear moduli of PEGDA-co-PEGMA hydrogels determined using an affine network model shown as a function of (A) concentrations of PEGDA (% w/w) at the time of cross-linking supplemented with various concentrations of PEGMA (% w/w) or (B) the molar concentration of PEGMA (mM) in the solution at the time of cross-linking for various PEGDA concentrations (% w/w). PEGMA concentration was expressed in millimoles per liter to facilitate comparison with the tissue engineering literature.
* Figure 4.6-ve , the number of effective cross-links per PEGDA molecule for PEGDA (0% PEGMA) hydrogels calculated from swelling data using the equation 5 for various values of F, the junction functionality. Calculations performed for 100 < F ≤ 106 are not * shown for clarity but were within 1.7% of the F = 100 values. ve calculated independently from mechanical data is also shown for reference.
134
* Figure 4.7-ve , the number of effective cross-links per PEGDA molecule for various PEGDA-co-PEGMA hydrogels calculated from swelling data shown as a function of (A) the concentration of PEGDA (% w/w) at the time of cross-linking supplemented with various concentrations of PEGMA (% w/w) or (B) the molar concentration of PEGMA (mM) in the solution at the time of cross-linking for various PEGDA concentrations (% * w/w). ve was estimated using equation 5 with a junction functionality (F) of 4.
* Figure 4.8-ve , the number of effective cross-links per PEGDA molecule for various PEGDA-co-PEGMA hydrogels calculated from mechanical data shown as a function of (A) the concentration of PEGDA (% w/w) at the time of cross-linking supplemented with various concentrations of PEGMA (% w/w) or (B) the molar concentration of PEGMA (mM) in the solution at the time of cross-linking for various PEGDA concentrations (% * w/w). ve was estimated using equation 6.
135 mechanical data, which did not depend on junction functionality, was generally well
* below the range of possible ve values calculated from swelling data, but followed the same trends.
* The value of ve gives an estimate of the type of cross-linking occurring in these
* network structures. For most of the gels ve was greater than 1 when assessed using both swelling and mechanical data (even in the limiting case F→∞), and for many of the gels
* with high compositions of PEGMA or PEGDA, ve was greater than 2, suggesting strongly that PEGDA entanglements play a critical role in the network structures of these materials.
4.3.6. Monoacrylated PEG Homopolymerization
The degree of polymerization of PEGMA macromers polymerized without
PEGDA under the same conditions as the gels was determined using GPC with absolute molecular weight determined by light scattering and refractometry data. The degree of polymerization increased monotonically with PEGMA concentration (Fig. 4.9).
4.4. Discussion
PEG di(meth)acrylate hydrogels copolymerized with PEGMA have been employed as tissue engineering scaffolds for many applications including bone, cartilage, nerve, and vascular tissues [3, 17, 18, 21]. The goal of this work was to examine the effects of this copolymerization on the properties and network structure of the resulting hydrogels.
We focused our investigation on a single combination of PEGDA and PEGMA macromers. 6 kD PEGDA is typical of the range commonly used for tissue engineering
(3.4 kD-8 kD) [3, 20, 29, 30] and 5 kD PEGMA is similar in size to peptide-PEG-
136
Figure 4.9-Number averaged degree of polymerization of PEGMA (2 kD) homopolymer formed under the same conditions as the hydrogels studied here as determined by gel permeation chromatography.
137 acrylate (for example, GRGDSP-PEGMA has a MW of ~4200). A neutral methyl-PEG terminus was employed to avoid confounding effects from specific peptide-PEGMA- solvent interactions which can alter gel properties [26].
The PEGDA composition at the time of cross-linking played a dominant role in determining the network structure (Figs. 4.3A and 4.5A). However, the substitution of
PEGMA for PBS in the prepolymerization macromer solution resulted in a decreased swelling ratio and an increased shear modulus, suggesting that the additional PEGMA increased the network cross-linking. Since the PEGMA does not contribute to the elastic component of the network, this result was unexpected.
* A new parameter ve , the number of effective cross-links per PEGDA molecule, was defined to better describe the network of PEGDA-co-PEGMA gels. Previous reports characterized PEGDA networks by calculating Mc [6, 13, 14, 29] using variations of the Bray-Merrill model [27], which was derived for networks formed from long polymer chains cross-linked with tetrafunctional linkages in solution. Given that (1)
PEGDA networks are formed from short polymer chains cross-linked by polymerization of junctions of unspecified functionality (Fig. 4.1), (2) there is no appropriate Mn to characterize the long polymer chains from which the network is to be sub-divided, and
(3) the Bray-Merrill model does not account for the PEGMA component of the
* copolymer gels, ve better characterizes PEGDA and PEGDA-co-PEGMA systems.
However, like original the Bray-Merrill model, our approach relies upon a statistical thermodynamic-based derivation of the entropies of solvation and distension. Thus, observations that the cross-linking nodes may form discrete microdomains within the gels[12] suggest there may be limitations to this approach.
138 Because the value of F was unknown in our experiments, we conducted a sensitivity analysis for this parameter, which indicated that the value of F had minimal
* impact on the magnitude and trends for ve , even if F was allowed to vary as a function of
PEGDA composition (F=4 for 5% PEGDA to F→∞ for 20% PEGDA). Since the trends were unaltered by F, we performed subsequent calculations using F=4. Although this
* over estimates ve for these gels, this value has been used implicitly to model PEGDA
* hydrogels by others [13] and serves as an upper limit on the calculated value of ve .
This analysis of the data supports the conclusion that the addition of PEGMA to
PEGDA polymerizations results in increased cross-linking of the network. Others have reported that replacement of buffer with PEGMA did not affect the properties of gel networks made with 743 Da PEGDA and 460 Da PEGMA [14]. For similar mass compositions, these smaller molecular weight materials resulted in gels with a cross-link density (effective chains per volume of polymer) 1-2 orders of magnitude higher than those observed in this study. This difference suggests that gels formed in solution with lower concentrations of reactive acrylate groups are more susceptible to the increased cross-linking via the addition of PEGMA, supporting a polymerization kinetics-based explanation for the observed effects.
* Calculations of ve from swelling data (for the limiting case of F→∞) and mechanical data, predicted a greater number of cross-link units in the gels than the number of PEGDA units for many of the compositions studied (Figs. 4.7 and 4.8). This result is highly suggestive that chain entanglements play a critical role in the network structure of these materials. Entanglements, if present, would represent a second set of cross-links in the network with a junction functionality of ~4 and a second distribution of
139 chain lengths which are subdivisions of the PEGDA chains, suggesting that a simple statistical thermodynamic model is insufficient to fully describe the network structure of these PEGDA gels and may explain the discrepancy between mechanical and swelling data. The potential for a bimodal distribution of chain lengths and junction functionalities in the entangled PEGDA networks further suggests that Mc derived from these network models (which for PEGDA-co-PEGMA gels is most appropriately calculated by dividing the mass of cross-linking polymer by the number of cross-linking
* chains in moles, equivalent to dividing Mn of the PEGDA by ve ) does not adequately represent the physical dimension inside these copolymerized gels and may not correlate well with actual gel mesh size or transport properties, as is the case with traditionally cross-linked networks [31].
Although the polymerization kinetics within these diacrylate networks is complex
[32, 33], a simplified model of the free radical polymerization suggests that the kinetic chain length of the resulting poly(acrylic acid) nodes should be roughly proportional to initial concentration of acrylate groups, which is increased with increasing PEGMA [34].
In the absence of gelation, we observed a direct relationship between the initial PEGMA concentration and the resulting polymer chain length, with a degree of polymerization on the order of 100’s of repeat units. These results are consistent with Burdick et al. who reported chain lengths on the order of 1000’s for more highly cross-linked polyanhydride diacrylate networks [11]. While the degree of polymerization seen in this experiment cannot be related to the polyacrylic acid nodes within the PEGDA hydrogels directly, these results suggest that the functionality of the nodes could be quite high. An increased kinetic chain length of the resulting nodes would be expected to increase the junction
140 functionality F, although this increase is not 1:1 since both PEGDA and non-network forming PEGMA are incorporated into the chain. The PEGMA-extended poly(acrylic acid) nodes, if present, would also fix additional chain entanglements in the network, as
* suggested by the analysis of ve . These results also suggest that copolymerization of other acrylate functionalized materials like poly(vinyl alcohol), hyaluronic acid, or fibrinogen[8-10] with additional monofunctional material likewise may enhance the cross- linking of these systems.
In tissue engineering applications, the concentration of PEGMA is much lower than PEGDA, typically 7 mM or less [3, 19, 28]. The effects of PEGMA demonstrated in this study were most apparent as the concentration approached or exceeded the concentration of PEGDA (Figs. 4.3B, 4.5B, 4.7B, and 4.8B). These results suggest that the low concentration of PEGMA (conjugated with a high affinity ligand like RGD) currently used for tissue engineering applications has a minimal effect on the gel’s network. However, several groups are beginning to report the incorporation of multiple peptides or growth factors [30, 35]. Others have reported the incorporation of inorganic groups or drugs to enhance the biological activity of these scaffolds [23, 25]. A tissue engineering scaffold that can incorporate a wide variety of biologically active factors at high concentration may more effectively mimic the natural extracellular space or provide more potent cues for cell behavior. Our results suggest that the formation of robust hydrogel scaffolds with a high concentration of biologically active moieties is feasible using a copolymerization approach.
141 4.5. Conclusions
The addition of PEGMA to photopolymerized PEGDA hydrogels similar to those used in tissue engineering applications enhanced the network cross-linking, as evidenced by a decreased swelling ratio, increased mechanical modulus, and an increase in the
* calculated effective chains per cross-linking PEGDA molecule, ve . Further analysis of the network indicated that this enhanced cross-linking occurred via the formation of entanglements which were fixed by extended poly(acrylic acid) cross-linking nodes in the networks. The results suggest that PEGMA-co-PEGDA gels can be formed with higher concentrations of PEGMA-tethered ligands than previously reported allowing the formation of scaffolds with a rich diversity of biological functionalities without sacrificing the integrity of the gel network.
4.6. Acknowledgements
The project described was supported by Grant Number 5R01EB002067 for the
National Institute of Biomedical Imaging and Bioengineering. The authors would like to acknowledge the facilities provided by the Center for Cardiovascular Biomaterials. JAB was also supported by NIH T32GM07250 and American Heart Association Predoctoral
Fellowship 0715422B.
142 4.7. References
1. Shin H, Jo S, Mikos AG. Modulation of marrow stromal osteoblast adhesion on biomimetic oligo[poly(ethylene glycol) fumarate] hydrogels modified with Arg-Gly- Asp peptides and a poly(ethyleneglycol) spacer. J Biomed Mater Res 2002;61(2):169-179.
2. Lutolf MP, Lauer-Fields JL, Schmoekel HG, Metters AT, Weber FE, Fields GB, Hubbell JA. Synthetic matrix metalloproteinase-sensitive hydrogels for the conduction of tissue regeneration: engineering cell-invasion characteristics. Proc Natl Acad Sci U S A 2003;100(9):5413-5418.
3. Burdick JA, Anseth KS. Photoencapsulation of osteoblasts in injectable RGD- modified PEG hydrogels for bone tissue engineering. Biomaterials 2002;23(22):4315-4323.
4. Merrill EW, Dennison KA, Sung C. Partitioning and diffusion of solutes in hydrogels of poly(ethylene oxide). Biomaterials 1993;14(15):1117-1126.
5. Hahn MS, Taite LJ, Moon JJ, Rowland MC, Ruffino KA, West JL. Photolithographic patterning of polyethylene glycol hydrogels. Biomaterials 2006;27(12):2519-2524.
6. Bryant SJ, Anseth KS. Hydrogel properties influence ECM production by chondrocytes photoencapsulated in poly(ethylene glycol) hydrogels. J Biomed Mater Res 2002;59(1):63-72.
7. Mann BK, Gobin AS, Tsai AT, Schmedlen RH, West JL. Smooth muscle cell growth in photopolymerized hydrogels with cell adhesive and proteolytically degradable domains: synthetic ECM analogs for tissue engineering. Biomaterials 2001;22(22):3045-3051.
8. Masters KS, Shah DN, Leinwand LA, Anseth KS. Crosslinked hyaluronan scaffolds as a biologically active carrier for valvular interstitial cells. Biomaterials 2005;26(15):2517-2525.
9. Almany L, Seliktar D. Biosynthetic hydrogel scaffolds made from fibrinogen and polyethylene glycol for 3D cell cultures. Biomaterials 2005;26(15):2467-2477.
10. Martens P, Anseth KS. Characterization of hydrogels formed from acrylate modified poly(vinyl alcohol) macromers. Polymer 2000;41(21):7715-7722.
11. Burdick JA, Lovestead TM, Anseth KS. Kinetic chain lengths in highly cross-linked networks formed by the photoinitiated polymerization of divinyl monomers: A gel permeation chromatography investigation. Biomacromolecules 2003;4(1):149-156.
143 12. Lin-Gibson S, Jones RL, Washburn NR, Horkay F. Structure-property relationships of photopolymerizable poly(ethylene glycol) dimethacrylate hydrogels. Macromolecules 2005;38(7):2897-2902.
13. Cruise GM, Scharp DS, Hubbell JA. Characterization of permeability and network structure of interfacially photopolymerized poly(ethylene glycol) diacrylate hydrogels. Biomaterials 1998;19(14):1287-1294.
14. Lin HQ, Kai T, Freeman BD, Kalakkunnath S, Kalika DS. The effect of cross- linking on gas permeability in cross-linked poly(ethylene glycol diacrylate). Macromolecules 2005;38(20):8381-8393.
15. Lin-Gibson S, Bencherif S, Cooper JA, Wetzel SJ, Antonucci JM, Vogel BM, Horkay F, Washburn NR. Synthesis and characterization of PEG dimethacrylates and their hydrogels. Biomacromolecules 2004;5(4):1280-1287.
16. Flory PJ. Principles of polymer chemistry. Ithaca,: Cornell University Press, 1953.
17. Peyton SR, Raub CB, Keschrumrus VP, Putnam AJ. The use of poly(ethylene glycol) hydrogels to investigate the impact of ECM chemistry and mechanics on smooth muscle cells. Biomaterials 2006;27(28):4881-4893.
18. Gunn JW, Turner SD, Mann BK. Adhesive and mechanical properties of hydrogels influence neurite extension. J Biomed Mater Res A 2005;72(1):91-97.
19. Hern DL, Hubbell JA. Incorporation of adhesion peptides into nonadhesive hydrogels useful for tissue resurfacing. J Biomed Mater Res 1998;39(2):266-276.
20. Nuttelman CR, Tripodi MC, Anseth KS. In vitro osteogenic differentiation of human mesenchymal stem cells photoencapsulated in PEG hydrogels. J Biomed Mater Res A 2004;68(4):773-782.
21. Hwang NS, Varghese S, Zhang Z, Elisseeff J. Chondrogenic differentiation of human embryonic stem cell-derived cells in arginine-glycine-aspartate-modified hydrogels. Tissue Eng 2006;12(9):2695-2706.
22. DeLong SA, Moon JJ, West JL. Covalently immobilized gradients of bFGF on hydrogel scaffolds for directed cell migration. Biomaterials 2005;26(16):3227-3234.
23. Nuttelman CR, Benoit DS, Tripodi MC, Anseth KS. The effect of ethylene glycol methacrylate phosphate in PEG hydrogels on mineralization and viability of encapsulated hMSCs. Biomaterials 2006;27(8):1377-1386.
24. Cushing MC, Liao JT, Jaeggli MP, Anseth KS. Material-based regulation of the myofibroblast phenotype. Biomaterials 2007;28(23):3378-3387.
144 25. Benoit DS, Nuttelman CR, Collins SD, Anseth KS. Synthesis and characterization of a fluvastatin-releasing hydrogel delivery system to modulate hMSC differentiation and function for bone regeneration. Biomaterials 2006;27(36):6102-6110.
26. Kim JH, Kim JG, Kim D, Kim YH. Preparation and properties of PEG hydrogel from PEG macromonomer with sulfonate end group. Journal of Applied Polymer Science 2005;96(1):56-61.
27. Bray JC, Merrill EW. Poly(vinyl alcohol) hydrogels. Formation by electron beam irradiation of aqueous solutions and subsequent crystallization. Journal of Applied Polymer Science 1973;17:3779-3794.
28. Mann BK, West JL. Cell adhesion peptides alter smooth muscle cell adhesion, proliferation, migration, and matrix protein synthesis on modified surfaces and in polymer scaffolds. J Biomed Mater Res 2002;60(1):86-93.
29. Buxton AN, Zhu J, Marchant R, West JL, Yoo JU, Johnstone B. Design and characterization of poly(ethylene glycol) photopolymerizable semi-interpenetrating networks for chondrogenesis of human mesenchymal stem cells. Tissue Eng 2007;13(10):2549-2560.
30. Mann BK, Schmedlen RH, West JL. Tethered-TGF-beta increases extracellular matrix production of vascular smooth muscle cells. Biomaterials 2001;22(5):439- 444.
31. Peppas NA. Hydrogels in medicine and pharmacy. Boca Raton, Fla.: CRC Press, 1986.
32. Anseth KS, Kline LM, Walker TA, Anderson KJ, Bowman CN. Reaction-Kinetics and Volume Relaxation during Polymerizations of Multiethylene Glycol Dimethacrylates. Macromolecules 1995;28(7):2491-2499.
33. Ward JH, Furman K, Peppas NA. Effect of monomer type and dangling end size on polymer network synthesis. Journal of Applied Polymer Science 2003;89(13):3506- 3519.
34. Odian GG. Principles of polymerization. 4th ed. Hoboken, N.J.: Wiley-Interscience, 2004.
35. Weber LM, Hayda KN, Haskins K, Anseth KS. The effects of cell-matrix interactions on encapsulated beta-cell function within hydrogels functionalized with matrix-derived adhesive peptides. Biomaterials 2007;28(19):3004-3011.
145 CHAPTER 5: The Influence of RGD-Bearing Hydrogels on the Re- expression of Contractile Vascular Smooth Muscle Cell Phenotype
Based on: Beamish JA, Fu AY, Choi A, Haq NA, Kottke-Marchant K, Marchant RE. The influence of RGD-bearing hydrogels on the re-expression of contractile vascular smooth muscle cell phenotype. Biomaterials 2009; in press.
5.1. Introduction
Tissue engineering approaches have the potential to drastically improve the function of synthetic vascular grafts by employing cells and tissues to provide the complex set of responses necessary to maintain long-term patency [1]. Over the past two decades, much progress has been made generating biologically functional tissue engineered blood vessels (TEBV) [2-4]. Vascular smooth muscle cells (SMCs) and the surrounding extracellular matrix (ECM) provide mechanical support and mediating changes in vascular tone. Therefore SMCs are a critical part of TEBVs. However, the mere presence of SMCs is not sufficient to recapitulate their function. In a variety of vascular pathologies, including restenosis after angioplasty and anastomotic intimal hyperplasia, normal SMCs, which exhibit a quiescent, contractile phenotype, de- differentiate into a phenotype characterized by proliferation and excessive synthesis of
ECM [5]. These “synthetic” SMCs are implicated in the stenosis of the vascular reconstruction. De-differentiation to the synthetic phenotype also occurs rapidly in the expansion culture of primary SMCs [6], such as is used for ex vivo graft preparation.
Scaffold materials designed to stimulate vascular regeneration must be able to regulate
SMC phenotype to prevent these failure mechanisms.
Regulation of SMC phenotype is complex and related to soluble signaling factors, mechanical stimuli, and cell substrate [7]. A wide variety of signaling factors have been implicated that affect the balance between contractile and synthetic phenotype including
146 platelet derived growth factor, basic fibroblast growth factor, insulin-like growth factors
(IGFs), heparin, and transforming growth factor β1 among many others [8-14]. It is also well known that mechanical stress can affect the phenotype of SMCs [7, 15, 16].
Furthermore, primary SMCs seeded on fibronectin (FN) de-differentiate more quickly in culture than cells seeded on laminin (LN) [13, 17-20] and the RGD peptide alone is sufficient to mimic the effects of FN [21]. In general, these studies have focused on factors that mediate de-differentiation of SMCs in culture. Strategies to re-differentiate cultured SMCs toward a contractile phenotype have been investigated less extensively.
The use of synthetic polymeric scaffolds with peptide cell-adhesive ligands allows quantitative control of cell-scaffold interactions and minimizes the possibility of confounding results from ECM bound growth factors. In particular, photopolymerized, poly(ethylene glycol) diacrylate (PEGDA)-based hydrogels have been employed widely as tissue engineering scaffolds for a variety of tissues including bone, cartilage, and blood vessels [22-26]. PEGDA hydrogels resist the adsorption of exogenous proteins and non- specific cell attachment. However, the inclusion of synthetic peptides that are copolymerized with the network can provide sites for specific cell adhesion [23]. In particular, RGD-based peptides, derived from integrin-binding proteins such as fibronectin, have been used widely to mediate adhesion [22-24, 26].
PEGDA and other PEG-based hydrogel systems have been used as scaffolds to study SMC biology by several groups [24, 27-29]. This work has established the ability of these scaffolds to support basic cell functions, such as attachment, migration, and growth [28, 29]. However, prior work has not examined the effect these systems have on the phenotype of vascular SMCs in detail, focusing on indirect markers of phenotypic
147 state, such as collagen production [24, 29], or examining phenotypic markers using qualitative immunostaining techniques [27, 30]. In this report, our goal was to assess the ability of RGD-bearing PEGDA hydrogels (RGD-gels) to support an in vitro transition of cultured human vascular SMCs toward a contractile phenotype.
5.2. Materials and Methods
5.2.1. Materials
All reagents were obtained from Sigma-Aldrich (St. Louis, MO) and used as received unless otherwise stated.
5.2.2. Preparation of PEGDA
PEG (MW 6000, dried in vacuo at 45 °C) was dissolved in anhydrous dichloromethane (0.2 g/ml), purged with argon, and placed on ice. Triethylamine (dried under molecular sieves, 1.1 molar excess per OH), and then acryloyl chloride (1.1 molar excess per OH) were added drop-wise. The reaction was stirred overnight at 4 °C. The acrylated PEG product was filtered to remove triethylamine hydrochloride salt, precipitated in diethyl ether, collected by filtration, and dried in vacuo. Substitution was approximately 72%, as determined by 1H NMR (in CDCl3) using the magnitude of peaks at 5.0-6.5 ppm (vinyl protons) and 3.7 ppm (PEG backbone).
5.2.3. Preparation of Peptide-PEG-acrylate Derivatives
The following peptides were synthesized as candidates to mediate cell attachment to PEGDA hydrogels: GRGDSP (fibronectin-derived)[31], KQAGDV (fibrinogen- derived) [27, 32], GSWSGSPPRRARVT (fibronectin-derived, syndecan-binding peptide)
[33], GKDGEA (collagen I-derived) [34], VAPG (elastin-derived) [35], and YIGSR
(laminin-derived) [36]. Peptides were synthesized using a solid phase peptide synthesizer
148 (Applied Biosystems, Model 433A, Foster City, CA) using standard Fmoc chemistry on an amide (Knorr) resin. Peptides were cleaved and deprotected using trifluoroacetic acid, precipitated in ether, dried, purified using reverse phase high performance liquid chromatography (HPLC), lyophilized, and stored at -20 °C. Successful peptide synthesis was confirmed by matrix assisted laser desorption/ionization mass spectroscopy
(MALDI-MS).
Peptides (added dropwise, 0-15 % molar excess) were reacted with acrylate-PEG-
N-hydroxysuccinimide (ACRL-PEG-NHS, 15-55 mg/ml, approximate MW 3400, Laysan
Bio, Huntsville, AL) in aqueous sodium bicarbonate (pH = 8.4) under argon for at least 2 h. Salts and unreacted peptide were removed by dialysis against water for 1 day (1:400 volume ratio, 3 exchanges minimum). The purified product was lyophilized and stored at
-20 °C. Conjugation was confirmed by comparison of MALDI-MS spectra of conjugates with the corresponding acrylate-PEG-COOH formed from hydrolyzed acrylate-PEG-
NHS starting material.
5.2.4. Synthesis of Hydrogel Substrates
Glass coverslips were cleaned by sonication in chloroform followed by 15 min treatment per surface by radio frequency (RF) glow discharge from argon bubbled through water. Glass was then coated with γ-methacryloxypropyl trimethoxysilane in
95% ethanol/5% water (v/v) adjusted to pH 5 with glacial acetic acid (~0.7% v/v) for 2 h per side, rinsed with copious amounts of pure ethanol, and dried in vacuo in covered glass petri dishes at 110 °C to anneal and sterilize the silanized coverslips. A uniform water-air contact angle greater than 60° confirmed successful modification. Flexible poly(ethylene terephthalate) (PET) sheets (McMaster-Carr, Cleveland, OH) were cleaned
149 by sonication in water and then in ethanol, treated by RF glow discharge, and sterilized with ethylene oxide.
Hydrogel films were formed on the silanized glass substrates. Hydrogel precursor solutions contained PEGDA (20% w/w), peptide-PEG-acrylate (5-10 mM, 2-4% w/w), and Irgacure 2959 (0.1% w/v, 1-[4-[2-Hydroxyethoxy]-phenyl]-2-hydroxy-2-methyl-1- propane-1-one, Ciba Specialty Chemicals, Tarrytown, NY) dissolved in phosphate buffered saline (PBS, pH 7.4). The resulting solution was sterilized by filtration (0.22
μm pore). In a sterile field, drops of precursor solution (1 drop per coverslip, 9 μl/cm2) were placed on PET sheets, covered with silanized glass coverslips, and polymerized for
10 min under ultraviolet irradiation (365 nm, 0.4-0.5 mW/cm2). The assembly was inverted and submerged in PBS. The PET sheet was peeled away leaving a thin hydrogel film covalently linked to the glass coverslip. The gel films were incubated in excess PBS for at least 2 h to leach unreacted material from the gels before use in cell experiments.
5.2.5. Cell Culture
Human coronary artery SMCs (HCASMCs, Lonza, Walkersville, MD) were routinely cultured in SmGM-2 (Lonza) growth medium which contains 5% fetal bovine serum (FBS) and proprietary amounts of basic fibroblast growth factor, epidermal growth factor, and insulin. Supplied antimicrobials were not added. All cell culture was performed at 37 °C, 5% CO2. All studies presented in this paper were performed at passage 7 with a single lot of cells (Lonza lot 6F4008) but similar studies (not shown) were performed with other cell lots and other passages (6-8) with similar results.
150 5.2.6. Differentiation of HCASMCs
Hydrogel films (D = 25 mm) formed with peptide-PEG-acrylate (5 mM) were placed in 6-well culture plates and secured with a ring of silicone tubing. Human fibronectin (FN, 1 μg/cm2) or murine laminin-1 (LN, 2 μg/cm2 from Engelbreth-Holm-
Swarm sarcoma) was coated (1-3 h) on tissue culture polystyrene (TCPS) as controls.
For immunofluorescence (IF) experiments FN and LN coatings were done on glass coverslips rather than TCPS. Prior to seeding, each surface was washed with PBS and preincubated with serum-free attachment medium (SFAM: insulin-transferrin-selenium supplement [ITS-X, 1X, Invitrogen, Carlsbad, CA], taurine [5 mM], bovine serum albumin [BSA, 1 mg/ml] in Dulbecco's Modified Eagle Medium [DMEM, Invitrogen]).
HCASMCs were dissociated with trypsin, resuspended in SFAM, and seeded at a density of 2.2-4.0 x 104 cells/cm2 for mRNA and protein experiments or 1.5 x 104 cells/cm2 for
IF experiments. After overnight seeding (14-17 h), SFAM was replaced with low serum
(2% FBS) DMEM containing heparin (400 μg/ml) to induce differentiation (LSM+H) or
SmGM-2 as a control. HCASMCs were cultured under these conditions for up to 6 d.
Medium was exchanged every 2 d. To assess ligand induced contractility, HCASMCs were exposed to 0-4 mM carbachol after 6 d of culture in LSM+H. HCASMC morphology was observed under phase contrast microscopy for 5 min after the addition of carbachol as described by others [13].
5.2.7. Real-time Reverse Transcription Polymerase Chain Reaction (RT-PCR)
RNA was collected from experimental samples immediately after seeding (14-17 h) and after 2 and 6 d of culture in LSM+H or SmGM using RNeasy spin columns
(Qiagen, Valencia, CA) following the manufacturer’s instructions. Total RNA yield was
151 determined spectroscopically and 130-320 ng RNA was reverse transcribed using the
Superscript First Strand Synthesis System (Invitrogen). The resulting cDNA template was diluted 2:16.5 in ultra pure water (Millipore filtration system, resistivity ≥ 18.2 MΩ- cm) and was used for real-time PCR analysis using the iQ SYBR Green Supermix (Bio- rad, Hercules, CA) and an iCycler fluorescence detection system (Bio-rad). Gene specific primers (Table 5.1) were designed using Oligoperfect software (Invitrogen). All primers sets were designed to amplify all known transcript variants of each gene.
A standard curve of pooled cDNA and a no template control were run on each plate for each gene to relate threshold amplification cycle (Ct) to relative transcript abundance. Melt curve analysis was performed after each PCR to ensure single product amplification. Reaction wells with aberrant results were excluded from further analysis
(generally fewer than 1 well/plate). Relative transcript abundance was normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) expression assuming 100% PCR efficiency.
5.2.8. Western Blotting
Protein was collected from experimental samples after seeding (14-17 h) and after
2 and 6 d of culture in LSM+H or SmGM-2 in radioimmunoprecipitation assay (RIPA) lysis buffer (Santa Cruz Biotechnology, Santa Cruz, CA) on ice. The soluble protein content was determined using the bicinchoninic acid (BCA) protein assay (Pierce
Biotechnologies, Rockford, IL). Samples were suspended in sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer (final composition: 62.5 mM Tris [pH 6.8], 2% w/v SDS, 10% v/v glycerol, 5% v/v β-mercaptoethanol, 25 μg/ml bromophenol blue). Protein samples (5 μg per lane) were separated on a Tris-HCl Ready
152
Table 5.1-Primers used for real-time RT-PCR
Product Size Name Gene Primers (BP) GGCCGAGATCTCACTGACTA Smooth Muscle α-Actin ACTA2 144 AGTGGCCATCTCATTTTCAA Smooth Muscle Basic CCCAGAAGTATGACCACCAG CNN1 226 Calponin TACTTGGTGATGGCCTTGAT CCAACAAGGGTCCTTCCTAT SM22α TAGLN 112 CCACACTGCACTATGATCCA Smooth Muscle Myosin CTGCAGCTTGGAAATATCGT MYH11 136 Heavy Chain GAGTGAGGATGGATCTGGTG Glyceraldehyde-3-Phosphate GACCTGACCTGCCGTCTA GAPDH 237 Dehydrogenase GTTGCTGTAGCCAAATTCGTT
153 Gel (12% resolving, 4% stacking, Bio-rad) and transferred to a nitrocellulose membrane.
Membranes were blocked in 4% (w/v) dried milk and probed for smooth muscle α-actin
(SMαA, 200-400 ng/ml, clone 1A4, Santa Cruz), calponin (400 ng/ml, clone CALP,
Santa Cruz), and SM-22α (800 ng/ml, polyclonal, H-75, Santa Cruz). The blot was visualized using horseradish peroxidase conjugated secondary antibodies (goat-anti- mouse: 80 ng/ml, goat-anti-rabbit: 100 ng/ml, Santa Cruz) and the SuperSignal West Pico
Chemiluminescent detection system (Pierce) using CL-XPosure film (Pierce).
Membranes were then stripped in stripping buffer (62.5 mM Tris, 2% SDS, 100 mM β- mercaptoethanol) for 20 min at 50 °C and reprobed for GAPDH (400 ng/ml, clone 0411,
Santa Cruz) using the same procedure. Films were scanned using a Hewlett-Packard
ScanJet IIcx and densitometry was performed on the digitized images using ImageJ
(NIH). Optical densities were normalized to lane 4 for each probe, then again to GAPDH for each lane before pooling data between experiments.
5.2.9. Immunofluorescent Staining
After 6 d of culture, surfaces were rinsed twice with PBS, fixed in 4% paraformaldehyde in PBS for 10 min, permeabilized in 0.1% (v/v) Triton-X 100, and blocked in blocking buffer (10 mM HEPES [pH 7.4], 150 mM NaCl, 2% BSA).
Immunofluorescence (IF) experiments utilized the same primary antibodies as western blotting. For SMαA and calponin co-staining, surfaces were incubated with primary antibodies (500 ng/ml), washed, and visualized with Alexafluor 568 goat-anti-mouse
IgG2a (Invitrogen) and Alexafluor 488 goat-anti-mouse IgG1 (Invitrogen), respectively.
For visualization of SM-22α, substrates were also blocked with an avidin/biotin blocking kit (Invitrogen), then incubated with anti-SM-22α antibodies (2 μg/ml), washed,
154 incubated with biotinlyated goat-anti-rabbit Fab fragment (1:400 dilution, Jackson
Immunoresearch, West Grove, PA), washed, and visualized with Alexafluor-488 conjugated streptavidin (5 μg/ml, Invitrogen). Co-staining for SMαA followed the same procedure as above. Nuclei were counterstained with 4’,6-diamidino-2-phenylindole
(DAPI, 1 μM). Fluorescent images were acquired on a Nikon Diphot 100 inverted microscope using a 10X objective with fixed acquisition settings to allow qualitative assessment of protein levels and using a 40X objective with individually optimized acquisition settings to assess intracellular protein organization. Control samples were stained using the above protocols without primary antibodies.
5.2.10. Attachment to hydrogel substrates
Hydrogel films (D = 15 mm) formed with 10 mM peptide-PEG-acrylate were placed in 24-well culture plates and retained with a small ring of silicone tubing to ensure that the well bottom was covered by only hydrogel film. Peptide-PEG-acrylate incorporation, and therefore the final peptide concentration in the network, was assumed to be equivalent for all peptides examined. FN and LN were coated on TCPS as controls.
Prior to seeding, each surface was washed with PBS and preincubated with SFAM.
HCASMCs were non-enzymatically detached (Cellstripper, Mediatech, Manassas, VA), resuspended in SFAM, and seeded near confluence for 16 h. Each substrate was washed gently with Hank’s balanced salt solution supplemented with HEPES (25 mM), MgCl2 (1 mM), CaCl2 (1 mM) and BSA (1 mg/ml) and imaged by phase contrast microscopy. The wash buffer was removed and the samples frozen at -80 °C. Frozen cells were lysed at room temperature using CyQUANT lysis buffer (Invitrogen) containing PicoGreen
(Invitrogen). DNA content of the lysate was measured using a fluorescent microplate
155 reader. To account for variations in cell seeding density, results for each experiment were normalized to FN attachment before pooling data between experiments.
For GRGDSP-PEG-acrylate (10 mM) containing hydrogels only, inhibition of attachment by soluble peptide was determined. HCASMCs were incubated with free
GRGDSP peptide (2 mM) for 30 min before seeding. Seeding duration was reduced to 1 h. Cell morphology and attachment were determined as described above.
5.2.11. Statistics
Statistical analysis was done using Microsoft Excel and Minitab 15. Data are represented as mean ± standard deviation of at least triplicate independent experiments.
Single comparisons were made using an un-paired student’s t-test. Analysis of variance
(ANOVA) followed by Tukey’s post hoc test was used for data sets with multiple comparisons. Statistical analysis of real-time PCR data was performed in the logarithmic domain. A value of α < 0.05 was considered significant.
5.3. Results
5.3.1. Analysis of Functionalized PEG Materials.
Conjugation of peptides to ACRL-PEG-NHS resulted in a shift in the polymer chain distribution to higher molecular weight on MALDI-MS spectra compared with hydrolyzed ACRL-PEG-NHS approximately equal to the molecular weight of peptide.
Hydrolyzed ACRL-PEG-NHS that was dialyzed, but not conjugated, did not result in a right shift of the distribution. Conjugation was also confirmed by a shift in the location of each peak relative to hydrolyzed ACRL-PEG-NHS, although it was not possible to resolve individual chain peaks in the MALDI-MS spectra for all of the conjugates studied.
156 5.3.2. Cell Morphology During Differentiation
Cell morphology was monitored over the course of culture in either LSM+H or
SmGM-2 medium. Within 2 d of medium change, HCASMCs cultured in LSM+H became less-spindle shaped with a hypertrophied cytoplasm that spread to fill gaps between cells and self-oriented in aligned band patterns. HCASMCs cultured in SmGM also self oriented into aligned patterns, but remained smaller and more fusiform in shape.
No differences in cell morphology were observed between substrates.
After 6 d of culture in LSM+H, HCASMCs on FN only were exposed to 0-4 mM carbachol. No changes in cell morphology were noted after 5 min of observation.
5.3.3. Expression of Contractile Marker mRNA
We observed decreased mRNA expression of SMαA (2.8-fold, p = 0.042), calponin (3.2-fold, p = 0.0012) and SM-22α (1.4-fold, p = 0.053) in HCASMCs seeded on LN compared with FN immediately after the seeding period in serum-free medium
(Fig. 5.1). Expression of SMαA, calponin, and SM-22α was also decreased on LN compared with RGD-gels (1.9-, 2.1-, and 1.2-fold, respectively), but, with the exception of calponin (p = 0.01), these differences were not significant. No significant differences were observed in SM-MHC expression (p > 0.82) or in SMαA, calponin, and SM-22α expression between RGD-gels and FN (p > 0.12).
Over 6 d of culture, the pattern of SMαA, calponin, and SM-22α expression followed a similar pattern (Fig. 5.2ABC). On each substrate, the expression of these markers increased substantially after 2 d in culture in LSM+H compared with expression at the end of the seeding period (Fig. 5.2ABC, p < 0.0003). Marker expression continued to increase over the remaining 4 d of culture, but was not statistically significant (p >
157
Figure 5.1-Expression of contractile marker gene mRNA after a 14-17 h seeding period in SFAM. The expression of each gene was normalized to the relative (to GAPDH) expression on fibronectin (FN). *: p < 0.05 with respect to FN, #: p < 0.05 with respect to RGD-gels.
158 0.89). The overall level of expression of each gene, relative to GAPDH, was indistinguishable between substrates after 2 and 6 d of culture in LSM+H (p > 0.95).
Expression of markers in LSM+H also was significantly higher than cells cultured in parallel on the same substrates in SmGM-2 (Fig. 5.2ABC, p < 0.0003)
Expression of SM-MHC also increased over 6 d of culture in LSM+H, but did so more slowly than SMαA, calponin, and SM-22α (Fig. 5.2D). After 2 d of culture in
LSM+H, SM-MHC expression increased slightly (1.7-2.9-fold, p > 0.59), but did not become statistically greater than the post-seeding level until the end of the 6-d culture period (6.7-16-fold increase, p < 0.03). SM-MHC expression in LSM+H also was greater than SmGM-2 parallel cultures (Fig. 5.4D, p < 0.03). SM-MHC transcript abundance was 4-6 orders of magnitude less than GAPDH.
Expression of all marker proteins decreased slightly from the post-seeding level over 6 d of culture in SmGM-2 on all substrates (Fig. 5.2, p < 0.05 for several conditions). There were no differences in the level of SMαA, calponin, SM-22α, or SM-
MHC expression between substrates for SMCs cultured in SmGM (p > 0.95).
5.3.4. Expression of Contractile Marker Proteins
Changes in SMαA, calponin, and SM-22α protein, assessed by Western blot (Fig.
5.3), generally were consistent with observed changes in mRNA expression (Fig. 5.2).
After 2 d of culture in LSM+H, we observed a qualitative increase in band intensity for
SMαA, calponin, and SM-22α (Fig. 5.3A). Semi-quantitative densitometry of these blots revealed a significant increase in SMαA only (p < 0.02) and extensive variability between samples. After 6 d of culture in LSM+H, we also observed a qualitative increase in band intensity for SMαA, calponin, and SM-22α (Fig. 5.3B) compared with the
159
Figure 5.2-Expression of SMαA (A), calponin (B), SM-22α (C), and SM-MHC (D) mRNA over 6 d of culture in LSM+H or SmGM. Expression levels shown are relative to GAPDH expression in the same sample. †: p < 0.0003 vs. post-seeding level, *: p < 0.05 vs. post-seeding level, ‡: p < 0.0003 vs. SmGM-2 (same day, same substrate), §: p < 0.023 vs. SmGM-2 (same day, same substrate), # p < 0.05 vs. Day 2 (same medium, same substrate).
160 parallel cultures in SmGM-2 or HCASMCs after seeding. Changes in SM-22α band intensity were subtle, consistent with the modest 2-3-fold change in mRNA levels observed by RT-PCR (Fig. 5.3AB). Semi-quantitative densitometry of these blots revealed significant increases in SMαA and calponin protein compared with post-seeding levels and with parallel cultures in SmGM-2 (p < 0.0012) and a slight increase in SM-
22α levels (Fig. 5.3C). No qualitative or semi-quantitative differences were observed between substrates for any time-points or culture conditions for all three proteins (p >
0.25).
IF staining for SMαA and calponin also revealed substantially more protein in cells cultured in LSM+H for 6 d compared with SmGM-2 (Fig. 5.4). No difference in the intensity of staining was observed between HCASMCs on FN, LN, or RGD-gels.
Moderate variations in expression were observed within the cell population, but IF staining was still strong in nearly all of the cells, suggesting that the entire population was responding uniformly. In wide field images, no differences in SM-22α IF staining intensity between samples were observed (Fig. 5.4, right column).
5.3.5. Intracellular Organization of Contractile Marker Proteins
High magnification IF staining for SMαA, calponin, and SM-22α revealed that expressed proteins co-localized into filaments within the cytoplasm of HCASMCs cultured in LSM+H for 6 d (Fig. 5.5). Filaments of SMαA were aligned with the long axis of the cells. Calponin co-localized most strongly with SMαA stress fibers in the central region of the cells and diminished in staining intensity towards the cell borders
(Fig. 5.5A). SM-22α stained more weakly than calponin, but co-localized with SMαA stress fibers evenly throughout the entire cytoplasm (Fig. 5.5B). SM-22α staining was
161
Figure 5.3-Relative expression of SMαA, calponin, and SM-22α protein after 2 d (A) or 6 d (B) of culture in LSM+H or SmGM-2 on fibronectin (FN), laminin (LN) and RGD- gels (Gel). Densitometry (C) was performed on 6-d blots from independent experiments (N = 5) and normalized to lane 4 (lighter shading). *: p < 0.05 vs. post-seeding levels, †: p < 0.001 vs. Day 6-SmGM-2 levels.
162
Figure 5.4-Expression and cellular distribution of SMαA (red, left column), calponin (green, center column), and SM-22α (green, right column) protein after 6 d of culture in LSM+H or SmGM-2 determined by immunofluorescent staining. Immunofluorescent stains of SMCs cultured with SmGM-2 on fibronectin and laminin (not shown) were similar to RGD-gels and showed substantially less staining than SMCs cultured in LSM+H. Staining and acquisition protocols were identical for all images for each protein. Nuclei were counterstained with DAPI (blue). Scale bar = 200μm.
163
Figure 5.5-Intracellular organization of SMαA (red), calponin (green, top row), and SM- 22α (green, bottom row) protein after 6 d of culture in LSM+H on RGD-gels determined by immunofluorescent staining. Co-localization is indicated by yellow in the right column. Nuclei were counterstained with DAPI (blue). Scale bar = 50 μm.
164 not homogenous through the cell population, but was more prominent in a small subset of cells, with many cells showing minimal staining. No differences in co-localization were observed between FN, LN, and RGD-gel substrates. Controls stained without primary antibody resulted in weak, diffuse background staining with no filamentous SMαA, calponin, or SM-22α structures.
5.3.6. Attachment to Hydrogel Substrates
After a 16 h seeding period, SMCs attached and spread only on GRGDSP-PEG-
ACRL containing hydrogels (RGD-gels). After washing, the adherent cells were well- spread on RGD-gels, FN, and LN substrates, but rounded cell morphology was observed for the sparse population of adherent cells on the other substrates (Fig. 5.6A). Compared with FN, LN, and RGD-gels, the DNA content of all of the other hydrogel substrates was reduced by at least 84% (p < 0.0001) (Fig. 5.6B). Among the non-RGD-gels, there were no significant differences in DNA content (p > 0.887). We also observed a small decrease (~35%) in measured DNA content on the FN and LN samples compared with the RGD-gels (p < 0.0001), which was not attributable directly to differences in the
PicoGreen assay response to the hydrogel substrate compared with protein coated TCPS.
Soluble GRGDSP peptide (2 mM) reduced the post-seeding DNA content on
GRGDSP-containing (10 mM) hydrogels by 60% (p < 0.03), which was similar to attachment to hydrogels with no peptide (p = 0.70). In the presence of GRGDSP peptide, the number of cells on the surface with a spread morphology was drastically reduced, with cell morphology generally more similar to that on unmodified hydrogels. Overall, cell attachment after 1 h incubation was reduced compared with overnight seeding, and attachment to GRGDSP hydrogels was less than FN controls.
165
Figure 5.6-Cell morphology (A) and attachment density (B) on additional peptide modified PEGDA hydrogels after 16 h in SFAM. Phase contrast micrographs (A) for peptide-gels not shown were similar to KQAGDV containing gels and gels containing no ACRL-PEG. Cell attachment and spreading on laminin was similar to fibronectin. Attachment was quantified by measuring DNA content (B) reported relative to GRGDSP-containing (10 mM) hydrogels. †: p < 0.0001 vs. GRGDSP containing gels, ‡: p < 0.0001 vs. FN or LN coated tissue culture polystyrene. Scale bar = 200 μm.
166 5.4. Discussion
The aim of this study was to investigate the ability of synthetic, peptide bearing hydrogel scaffolds to support the re-differentiation of cultured vascular SMCs toward a contractile cell phenotype. A photopolymerizable PEGDA hydrogel modified with pendant GRGDSP peptides was utilized as a scaffold material with well-defined SMC- material interactions. This system allowed us to explore the role that a specific RGD ligand-cell interaction has on the ability of cultured HCASMCs to re-differentiate to a contractile phenotype in vitro. Once seeded on these substrates, we utilized low serum culture conditions and soluble heparin to drive SMC differentiation and monitored phenotype using a panel of markers at the level of mRNA, protein, and intracellular organization.
Here we showed that FN, LN, and RGD-gels support equivalent levels of expression of the contractile markers, SMαA, calponin, and SM-22α, over a 6-d culture period at the transcriptional and translational level and that these markers are organized into filaments within the cell. By these measures, we achieved a significant shift in cell phenotype on our chemically well-defined RGD-gel system. These results are consistent with other studies of vascular SMC phenotype on PEG-based gels [27, 30] which demonstrate the qualitative presence of contractile SMC markers such as SMαA, calponin, and caldesmon detected by immunostaining. Our results quantify these changes at the mRNA and protein level, and indicate that with appropriate culture conditions, phenotype switching can occur rapidly, with significant changes in the transcriptional profile and protein levels after only 2 d of culture. Furthermore, we demonstrated that changes in marker levels were also associated with distinct
167 organization of contractile proteins into fibrillar structures in the cell cytoplasm. This organization was critical to distinguish positive staining from diffuse intracellular background in IF studies, which was very similar to control slides without primary antibodies. We also noted weak fibrillar staining for SMαA, calponin, and SM-22α (Fig.
5.4) in SMCs cultured in SmGM-2, which without comparison to cells cultured in
LSM+H may have caused them to appear differentiated. These issues suggest that IF studies alone are not sufficient to study the phenotype of cultured SMCs. While our study examined the contractile state of SMCs on selected substrates, we did not examine other issues that might impact the differentiation of SMCs such as variation in the ligand density of RGD or the mechanical properties of the underlying substrate. Since these issues have been shown to affect cell proliferation and collagen production [27, 29], the roles these factors play on SMC differentiation will be excellent candidates for additional study using the quantitative methods described here.
While dramatically bringing their marker profile closer to a contractile phenotype, the cells did not show evidence of contraction in response to carbachol. Furthermore, the overall level of expression of SM-MHC mRNA was significantly lower than the other markers and the SM-MHC protein was not detectable by western blotting or IF (not shown). This molecule is not only a marker of cell phenotype, but also a critical component of the contractile machinery. Taken together, these observations suggest the contractile apparatus was not reformed completely. These results also illustrate the importance of completely characterizing cell phenotype quantitatively and that even with the achievement of quantitative increases in several marker proteins, more work is needed to restore a truly contractile phenotype to cultured SMCs.
168 Extensive literature suggests that FN plays an important role in the loss of contractile SMC phenotype during primary culture while LN may delay this transition
[13, 17-20]. The effect of FN could also be mimicked by the RGD-ligand alone [21].
Our experiments were designed to assess the roles these ECM ligands have on the re- differentiation of cultured vascular SMCs, which has not been studied extensively. We observed decreased expression of contractile marker mRNA (with no differences in protein levels) on the LN substrate compared with FN and RGD-gels after seeding in serum-free medium. This result was the opposite of that anticipated if cultured cells behaved similarly to primary vascular SMCs. These results suggest that during expansion culture, the signaling machinery in native SMCs may be reprogrammed to respond differently to LN. Such changes in signaling machinery have been observed in cultured SMCs with IGF-1, which initially helps to maintain the contractile phenotype of primary rat aortic SMCs, but after de-differentiation in culture, stimulates proliferation and migration [37].
However, it is also possible that low initial contractile marker expression on LN may have resulted from poor initial adhesion on LN rather than altered response to SMC-
LN interactions per se. Others have suggested that there may be a correlation between focal adhesion formation and SMC differentiation [27]. We observed differences in SMC morphology between LN and FN during the early stages of the adhesion process (after 0-
4 h, not shown), but we found that after 4 h cells seeded LN did not form attachments sufficient to withstand a change in medium. However, cell morphology was indistinguishable between the substrates after overnight seeding. Others have shown that cultured baboon SMCs plated on LN developed a well-formed provisional FN matrix
169 after 4 h of culture, effectively masking the LN substrate [38]. Although we did not measure FN production directly, our morphological observations were consistent with our human SMCs also organizing a FN matrix within the first few hours of culture, which subsequently provided robust cell adhesion but delayed SMC marker expression. The production and organization of endogenous FN also may have explained why the initial differences in cell phenotype on LN did not persist during the remainder of the differentiation period, especially after the addition of LSM+H, which contains exogenous cell-adhesive serum proteins including FN.
To address this issue, we prepared hydrogels containing peptides from LN and other ECM proteins in an effort to directly assess the role of ligand identity on modulation of SMC differentiation. In contrast to LN coated TCPS, the hydrogel substrates were designed to resist the adsorption of exogenous protein. We screened a variety of peptides derived from ECM proteins including those known to modulate SMC phenotype: KQAGDV (fibrinogen) [27, 32], GKDGEA (type I collagen) [34], YIGSR
(laminin) [36], and VAPG (elastin) [35]. We also assessed a heparin binding peptide, -
SPPRRARVT [33], in an attempt to engage the syndecan family of receptors, which is known to be important for SMC adhesion [39]. Except for GRGDSP, we did not observe attachment or spreading on any of the peptide-containing hydrogels, which precluded us from performing differentiation studies on these surfaces. This result was unanticipated since all of the above peptides have documented interactions with receptors known to be expressed on SMCs [39] or have been empirically shown to bind SMCs [27, 32]. Many of the published cell attachment studies used a solid substrate to present the peptides that does not resist protein adsorption to the same degree as PEG. In the PEGDA hydrogels
170 we employed, no cell adhesion was observed unless the hydrogels were modified with a pendant peptide, making this a more rigorous system for assessing cell-peptide interactions in the context of cell adhesion.
However, in the case of KQAGDV and VAPG, SMC attachment has been reported on PEGDA hydrogels modified with these peptides [27, 40]. In these studies, specific attachment either was not confirmed by inhibition of attachment with soluble peptide [27] or soluble peptide did not inhibit attachment [40]. These studies also did not exclude the possibility of non-specific or serum dependent adhesion. These results also highlight the potential for SMC species differences, lot-to-lot differences in cell response, or changes in cell receptor library due to slightly different expansion culture techniques, although we found similar results for VAPG and YIGSR surfaces with two additional lots of human SMCs (not shown), and other groups have reported difficulty in establishing
SMC attachment to YIGSR-bearing hydrogels [28]. Our results, obtained by seeding in serum-free medium on protein adsorption resistant PEGDA hydrogels, suggests that there is variable cell response in SMC-peptide interactions and that, among the peptides tested, only the GRGDSP or its derivatives can provide reliable, peptide-specific cell attachment to these gels. We have also confirmed that SMC attachment to RGD-gels was inhibited by soluble peptide and did not occur on non-RGD bearing hydrogels. These results indicate that SMC attachment to these scaffolds, at least initially, is highly specific to the
RGD ligand.
Due to the well-defined chemistry of the PEGDA system, the overall properties of the gel and the concentration of this ligand can be controlled quantitatively and independently (Chapter 4, [41]). Here, we have demonstrated that the RGD-gels can
171 support differentiation of cultured SMCs toward a contractile phenotype. Consequently, this scaffold material will be an excellent candidate for further development as a bioactive scaffold for smooth muscle tissue engineering.
Furthermore, our results suggest that cultured SMCs may respond differently to the extracellular matrix than primary cells, which is an important design consideration for scaffold systems intended for ex vivo expanded cells. Since the population of SMCs that would likely populate a regeneration-inducing TEBV in vivo would also adopt a synthetic phenotype (in order to migrate and proliferate in the scaffold), scaffold-cell interactions might be best designed to accommodate a synthetic-to-contractile switch, rather than inhibit de-differentiation to the synthetic phenotype. Our results suggest that a fibronectin or fibronectin-mimicking scaffold, such as the RGD-gels employed here, may be superior for this purpose.
5.5. Conclusions
In this report, we demonstrated that FN, LN, and RGD-gels support equivalent levels of expression of the contractile markers, SMαA, calponin, and SM-22α over a 6-d culture period in low serum medium at the transcriptional and translational level and that these markers are organized into filaments within the cell. These results indicate that
SMCs cultured on RGD-bearing hydrogels can re-differentiate toward a contractile phenotype suggesting this material is an excellent candidate for further development as a bioactive scaffold that regulates SMC phenotype in TEBV designs.
5.6. Acknowledgements
The project described was supported by Grant Number 5R01EB002067 for the
National Institute of Biomedical Imaging and Bioengineering and Grant Number
172 1R01HL087843 for the National Heart, Lung, and Blood Institute. JAB also was supported by NIH T32GM07250 and American Heart Association Predoctoral
Fellowship 0715422B. We would like to acknowledge the technical assistance of Dr.
Faina Kligman in the synthesis and purification of peptides, Dr. Meghan Pennini in developing RT-PCR assays, and Domenick Prosdocimo in developing Western blot assays.
173 5.7. References
1. Brewster LP, Bufallino D, Ucuzian A, Greisler HP. Growing a living blood vessel: insights for the second hundred years. Biomaterials 2007;28(34):5028-5032.
2. Weinberg CB, Bell E. A blood vessel model constructed from collagen and cultured vascular cells. Science 1986;231(4736):397-400.
3. Niklason LE, Gao J, Abbott WM, Hirschi KK, Houser S, Marini R, Langer R. Functional arteries grown in vitro. Science 1999;284(5413):489-493.
4. L'Heureux N, Dusserre N, Konig G, Victor B, Keire P, Wight TN, Chronos NA, Kyles AE, Gregory CR, Hoyt G, Robbins RC, McAllister TN. Human tissue- engineered blood vessels for adult arterial revascularization. Nat Med 2006;12(3):361-365.
5. Reidy MA. Regulation of Arterial Smooth Muscle Growth. In: Schwartz SM, Mecham RP, editors. The vascular smooth muscle cell : molecular and biological responses to the extracellular matrix. San Diego: Academic Press, 1995. p. xvi, 410 p.
6. Chamley-Campbell J, Campbell GR, Ross R. The smooth muscle cell in culture. Physiol Rev 1979;59(1):1-61.
7. Stegemann JP, Hong H, Nerem RM. Mechanical, biochemical, and extracellular matrix effects on vascular smooth muscle cell phenotype. J Appl Physiol 2005;98(6):2321-2327.
8. Hedin U, Daum G, Clowes AW. Heparin inhibits thrombin-induced mitogen- activated protein kinase signaling in arterial smooth muscle cells. J Vasc Surg 1998;27(3):512-520.
9. Pukac L, Huangpu J, Karnovsky MJ. Platelet-derived growth factor-BB, insulin-like growth factor-I, and phorbol ester activate different signaling pathways for stimulation of vascular smooth muscle cell migration. Exp Cell Res 1998;242(2):548-560.
10. Pukac LA, Carter JE, Ottlinger ME, Karnovsky MJ. Mechanisms of inhibition by heparin of PDGF stimulated MAP kinase activation in vascular smooth muscle cells. J Cell Physiol 1997;172(1):69-78.
11. Daum G, Hedin U, Wang Y, Wang T, Clowes AW. Diverse effects of heparin on mitogen-activated protein kinase-dependent signal transduction in vascular smooth muscle cells. Circ Res 1997;81(1):17-23.
12. Rauch BH, Millette E, Kenagy RD, Daum G, Clowes AW. Thrombin- and factor Xa- induced DNA synthesis is mediated by transactivation of fibroblast growth factor receptor-1 in human vascular smooth muscle cells. Circ Res 2004;94(3):340-345.
174 13. Hayashi K, Saga H, Chimori Y, Kimura K, Yamanaka Y, Sobue K. Differentiated phenotype of smooth muscle cells depends on signaling pathways through insulin- like growth factors and phosphatidylinositol 3-kinase. J Biol Chem 1998;273(44):28860-28867.
14. Bjorkerud S. Effects of transforming growth factor-beta 1 on human arterial smooth muscle cells in vitro. Arterioscler Thromb 1991;11(4):892-902.
15. Kim BS, Nikolovski J, Bonadio J, Mooney DJ. Cyclic mechanical strain regulates the development of engineered smooth muscle tissue. Nat Biotechnol 1999;17(10):979-983.
16. Williams B. Mechanical influences on vascular smooth muscle cell function. J Hypertens 1998;16(12 Pt 2):1921-1929.
17. Hedin U, Bottger BA, Forsberg E, Johansson S, Thyberg J. Diverse effects of fibronectin and laminin on phenotypic properties of cultured arterial smooth muscle cells. J Cell Biol 1988;107(1):307-319.
18. Hayward IP, Bridle KR, Campbell GR, Underwood PA, Campbell JH. Effect of extracellular matrix proteins on vascular smooth muscle cell phenotype. Cell Biol Int 1995;19(10):839-846.
19. Thyberg J, Hultgardh-Nilsson A. Fibronectin and the basement membrane components laminin and collagen type IV influence the phenotypic properties of subcultured rat aortic smooth muscle cells differently. Cell Tissue Res 1994;276(2):263-271.
20. Yamamoto M, Yamamoto K, Noumura T. Type I collagen promotes modulation of cultured rabbit arterial smooth muscle cells from a contractile to a synthetic phenotype. Exp Cell Res 1993;204(1):121-129.
21. Hedin U, Bottger BA, Luthman J, Johansson S, Thyberg J. A substrate of the cell- attachment sequence of fibronectin (Arg-Gly-Asp-Ser) is sufficient to promote transition of arterial smooth muscle cells from a contractile to a synthetic phenotype. Dev Biol 1989;133(2):489-501.
22. Burdick JA, Anseth KS. Photoencapsulation of osteoblasts in injectable RGD- modified PEG hydrogels for bone tissue engineering. Biomaterials 2002;23(22):4315-4323.
23. Hern DL, Hubbell JA. Incorporation of adhesion peptides into nonadhesive hydrogels useful for tissue resurfacing. J Biomed Mater Res 1998;39(2):266-276.
24. Mann BK, Gobin AS, Tsai AT, Schmedlen RH, West JL. Smooth muscle cell growth in photopolymerized hydrogels with cell adhesive and proteolytically degradable domains: synthetic ECM analogs for tissue engineering. Biomaterials 2001;22(22):3045-3051.
175 25. Nuttelman CR, Tripodi MC, Anseth KS. In vitro osteogenic differentiation of human mesenchymal stem cells photoencapsulated in PEG hydrogels. J Biomed Mater Res A 2004;68(4):773-782.
26. Hwang NS, Varghese S, Zhang Z, Elisseeff J. Chondrogenic differentiation of human embryonic stem cell-derived cells in arginine-glycine-aspartate-modified hydrogels. Tissue Eng 2006;12(9):2695-2706.
27. Peyton SR, Raub CB, Keschrumrus VP, Putnam AJ. The use of poly(ethylene glycol) hydrogels to investigate the impact of ECM chemistry and mechanics on smooth muscle cells. Biomaterials 2006;27(28):4881-4893.
28. Fittkau MH, Zilla P, Bezuidenhout D, Lutolf MP, Human P, Hubbell JA, Davies N. The selective modulation of endothelial cell mobility on RGD peptide containing surfaces by YIGSR peptides. Biomaterials 2005;26(2):167-174.
29. Mann BK, West JL. Cell adhesion peptides alter smooth muscle cell adhesion, proliferation, migration, and matrix protein synthesis on modified surfaces and in polymer scaffolds. J Biomed Mater Res 2002;60(1):86-93.
30. Hahn MS, McHale MK, Wang E, Schmedlen RH, West JL. Physiologic pulsatile flow bioreactor conditioning of poly(ethylene glycol)-based tissue engineered vascular grafts. Ann Biomed Eng 2007;35(2):190-200.
31. Pierschbacher MD, Ruoslahti E. Cell attachment activity of fibronectin can be duplicated by small synthetic fragments of the molecule. Nature 1984;309(5963):30- 33.
32. Mann BK, Tsai AT, Scott-Burden T, West JL. Modification of surfaces with cell adhesion peptides alters extracellular matrix deposition. Biomaterials 1999;20(23- 24):2281-2286.
33. Sagnella S, Anderson E, Sanabria N, Marchant RE, Kottke-Marchant K. Human endothelial cell interaction with biomimetic surfactant polymers containing Peptide ligands from the heparin binding domain of fibronectin. Tissue Eng 2005;11(1- 2):226-236.
34. Staatz WD, Fok KF, Zutter MM, Adams SP, Rodriguez BA, Santoro SA. Identification of a tetrapeptide recognition sequence for the alpha 2 beta 1 integrin in collagen. J Biol Chem 1991;266(12):7363-7367.
35. Mecham RP, Hinek A, Entwistle R, Wrenn DS, Griffin GL, Senior RM. Elastin binds to a multifunctional 67-kilodalton peripheral membrane protein. Biochemistry 1989;28(9):3716-3722.
36. Graf J, Ogle RC, Robey FA, Sasaki M, Martin GR, Yamada Y, Kleinman HK. A pentapeptide from the laminin B1 chain mediates cell adhesion and binds the 67,000 laminin receptor. Biochemistry 1987;26(22):6896-6900.
176 37. Hayashi K, Shibata K, Morita T, Iwasaki K, Watanabe M, Sobue K. Insulin receptor substrate-1/SHP-2 interaction, a phenotype-dependent switching machinery of insulin-like growth factor-I signaling in vascular smooth muscle cells. J Biol Chem 2004;279(39):40807-40818.
38. Hedin UL, Daum G, Clowes AW. Disruption of integrin alpha 5 beta 1 signaling does not impair PDGF-BB-mediated stimulation of the extracellular signal-regulated kinase pathway in smooth muscle cells. J Cell Physiol 1997;172(1):109-116.
39. Moiseeva EP. Adhesion receptors of vascular smooth muscle cells and their functions. Cardiovasc Res 2001;52(3):372-386.
40. Gobin AS, West JL. Val-ala-pro-gly, an elastin-derived non-integrin ligand: smooth muscle cell adhesion and specificity. J Biomed Mater Res A 2003;67(1):255-259.
41. Beamish JA, Zhu J, Kottke-Marchant K, Marchant RE. The effects of monoacrylated poly(ethylene glycol) on the properties of poly(ethylene glycol) diacrylate hydrogels used for tissue engineering. J Biomed Mater Res 2009;in press.
177 CHAPTER 6: The Effects of Heparin Releasing Hydrogels on Vascular Smooth Muscle Cell Phenotype
6.1. Introduction
Anastomotic intimal hyperplasia (IH) is a significant cause of long-term failure in synthetic vascular grafts. In this pathology, normal smooth muscle cells (SMCs), which exhibit a quiescent, contractile phenotype, de-differentiate into a phenotype characterized by proliferation and excessive synthesis of extracellular matrix (ECM) [1]. These
“synthetic” SMCs are implicated in the stenosis of the vascular reconstruction and ultimately contribute to failure. Many stimuli induce this pathology, including mechanical mismatch, flow disturbances, and injury [1, 2]. Vascular tissue engineering has the potential to mitigate response to these stimuli by employing cells and tissues to provide the complex set of responses necessary to maintain long-term patency [3].
Scaffold materials designed to regulate SMC phenotype are required to realize this potential.
Regulation of SMC phenotype is complex and related to soluble signaling factors, mechanical stimuli, and cell substrate [4]. A variety of signaling factors have been implicated that affect the balance between contractile and synthetic phenotype including platelet derived growth factor, basic fibroblast growth factor (bFGF), heparin, and transforming growth factor β1, among many others [5-11]. It also is well known that mechanical stress can affect the phenotype of SMCs [4, 12, 13]. Furthermore, primary
SMCs seeded on fibronectin (FN) de-differentiate more quickly in culture than cells seeded on laminin (LN) [10, 14-17]. In general, these studies have focused on factors that mediate de-differentiation of SMCs in culture. Strategies to re-differentiate cultured
SMCs toward a contractile phenotype have been investigated less extensively.
178 In addition to its well-known anti-coagulation function, it has been known for several decades that heparin is capable of reducing SMC proliferation in vitro and in vivo
[7, 8, 18, 19]. Heparin’s antiproliferative effect is unrelated to its anticoagulant activity
[20]. The most important structural determinant of anti-proliferative activity appears to be the overall level of sulfation, although changes in size, structure, and specific O- or N- linked sulfation patterns all contribute to activity [21]. One prominent mechanism of heparin’s antiproliferative activity may be related to disruption of exogenous or autocrine bFGF signaling [9, 22]. However, heparin may also directly stimulate extracellular receptors [23] and can be internalized [24] allowing it to modulate cytoplasmic signaling pathways [25]. Heparin also can induce expression of smooth muscle α-actin (SMαA)
[26-28] and other markers of contractile smooth muscle phenotype [26], in addition to inhibiting SMC proliferation. These results suggest that heparin may be a useful approach to drive SMC differentiation toward a contractile phenotype.
Photopolymerized poly(ethylene glycol) diacrylate (PEGDA)-based hydrogels have been employed widely as tissue engineering scaffolds for a variety of tissues including bone, cartilage, and blood vessels [29-33]. Synthetic peptides can be copolymerized with the network to provide sites for specific cell adhesion to the hydrogel
[30]. The network structure of these hydrogels can be controlled quantitatively and independently of peptide incorporation (Chapter 4, [34]) allowing these gels to be tailored to serve as depots for the controlled delivery of bioactive agents, such as heparin.
PEGDA and other PEG-based hydrogels systems have been used as scaffolds to study SMC biology by several groups [31, 35-39]. This work has established the ability of these scaffolds to support basic cell functions, such as attachment, migration, and
179 growth [36, 37]. Recently, SMC phenotype in and on these hydrogel constructs has been characterized in greater detail and several mechanisms to modulate phenotype have been explored [35, 39]. Our group has recently shown that SMCs cultured on RGD-bearing
PEGDA hydrogels can rapidly and robustly shift toward a contractile phenotype (Chapter
5, [38]). However, these studies utilized exogenous stimuli to drive SMC differentiation.
In this report, our goal was to explore the potential of heparin releasing, RGD-bearing
PEGDA hydrogels scaffolds to drive SMCs toward a contractile phenotype without extensive exogenous stimulation.
6.2. Materials and Methods
6.2.1. Materials
All reagents were obtained from Sigma-Aldrich (St. Louis, MO) and used as received unless otherwise stated.
6.2.2. Preparation of PEGDA
PEG (MW 1000, 3000, or 6000) was dried by azeotropic distillation with toluene and stored in vacuo for 1 d at 60-80 °C. The dry PEG was dissolved in anhydrous dichloromethane (DCM, 0.2 g/ml), purged with argon, and placed on ice. Triethylamine
(dried under molecular sieves, 1.1 molar excess per OH), then acryloyl chloride (1.1 molar excess per OH) were added drop-wise. The reaction was stirred overnight at 4 °C.
Excess DCM was evaporated to concentrate the acrylated PEG product, which then was filtered to remove triethylamine hydrochloride salt, precipitated in diethyl ether, collected by filtration, reprecipitated, and dried in vacuo. The resulting products are denoted
PEGDA1k, PEGDA3k, and PEGDA6k, respectively. Substitution was approximately
70%, as determined by 1H NMR (Chapter 5, [38]).
180 6.2.3. Preparation of GRGDSP-PEG-Acrylate Derivatives
GRGDSP-PEG-acrylate was prepared as described previously (Chapter 5, [38]).
Briefly, GRGDSP peptide was synthesized using a solid phase peptide synthesizer
(Applied Biosystems, Model 433A, Foster City, CA) using standard Fmoc chemistry on an amide (Knorr) resin. Peptides were cleaved and deprotected using trifluoroacetic acid and purified using reverse phase high performance liquid chromatography (HPLC).
Peptide synthesis was confirmed by matrix assisted laser desorption/ionization mass spectroscopy (MALDI-MS).
GRGDSP (added dropwise, 0-15% molar excess) was reacted with acrylate-PEG-
N-hydroxysuccinimide (ACRL-PEG-NHS, 40 mg/ml, MW ~3400, Laysan Bio,
Huntsville, AL) in aqueous sodium bicarbonate (pH 8.4) under argon for 2 h. Salts and unreacted peptide were removed by dialysis against water for 1 day (1:400 volume ratio,
3 exchanges minimum, 1000 MWCO). The purified product was lyophilized and stored at -20 °C. Conjugation of the peptide was confirmed by MALDI-MS.
6.2.4. Cell Culture
Human coronary artery SMCs (HCASMCs, Lonza, Walkersville, MD) were routinely cultured in SmGM-2 (Lonza) growth medium which contains 5% fetal bovine serum (FBS) and proprietary amounts of basic fibroblast growth factor, epidermal growth factor, and insulin. Media used for hydrogel experiments were supplemented with gentamycin and amphotericin (1X, Lonza). All cell culture was performed at 37 °C, 5%
CO2. HCASMCs were used between passage 6 and 9. Unless otherwise noted, all materials used for cell culture were received sterile or were steam sterilized prior to use.
181 6.2.5. Effects of Heparin on HCASMC Proliferation
HCASMCs were cultured to confluence, growth arrested in serum-free medium
(SFM: insulin-transferrin-selenium supplement [ITS-X, 1X, Invitrogen, Carlsbad, CA], taurine [5 mM], bovine serum albumin [BSA, 1 mg/ml] in Dulbecco's Modified Eagle
Medium [DMEM, Invitrogen]) for 3 d, and seeded on fibronectin (FN, 1 μg/cm2) coated
24-well culture plates (5,000 cells/cm2) overnight. Medium was then changed to DMEM containing FBS (2% or 10% v/v) or SmGM-2 supplemented with heparin (400 μg/ml, from porcine intestinal mucosa, approximate MW 20 kD, H9399, Sigma), chondroitin sulfate (400 μg/ml), or empty vehicle. Medium was changed every 2 d. After 6 d, relative cell populations were determined using the CellTiter 96 AQueous One Solution
Cell Proliferation Assay (MTS Assay, Promega, Madison, WI).
6.2.6. Effects of Heparin on HCASMC Differentiation
HCASMCs were cultured in 6-well plates. After the HCASMCs attained confluence, the medium was changed to low serum medium (LSM: 2% v/v FBS in
DMEM) supplemented with heparin (0-1000 μg/ml). LSM containing chondroitin sulfate
(400 μg/ml) and SmGM were used as controls. Medium was changed every 2 d. After 6 d, protein was collected in radioimmunoprecipitation assay (RIPA) lysis buffer (Santa
Cruz Biotechnology, Santa Cruz, CA) on ice. The soluble protein content was determined using the bicinchoninic acid (BCA) protein assay (Pierce Biotechnologies,
Rockford, IL). Expression of contractile marker proteins was determined by western blotting.
182 6.2.7. Western Blotting
Samples were suspended in sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer (final composition: 62.5 mM Tris [pH 6.8],
2% w/v SDS, 10% v/v glycerol, 5% v/v β-mercaptoethanol, 25 μg/ml bromophenol blue).
Protein samples (5 μg per lane) were separated on a Tris-HCl Ready Gel (12% resolving,
4% stacking, Bio-rad) and transferred to a nitrocellulose membrane. Membranes were blocked in 4% (w/v) dried milk and probed for SMαA (200 ng/ml IgG2a, clone 1A4,
Santa Cruz), calponin (400 ng/ml IgG1, clone CALP, Santa Cruz), and SM-22α (800 ng/ml, rabbit polyclonal, H-75, Santa Cruz). The blot was visualized using horseradish peroxidase conjugated secondary antibodies (goat-anti-mouse IgG2a: 1:20,000, goat-anti- mouse IgG1: 1:2000, goat-anti-rabbit: 1:2000, Santa Cruz) and the SuperSignal West Pico
Chemiluminescent detection system (Pierce) using CL-XPosure film (Pierce).
Membranes were then stripped in stripping buffer (62.5 mM Tris, 2% SDS, 100 mM β- mercaptoethanol) for 20 min at 50 °C and reprobed for GAPDH (400 ng/ml, clone 0411,
Santa Cruz) using the same procedure. Films were scanned using a Hewlett-Packard
ScanJet IIcx and densitometry was performed on the digitized images using ImageJ
(NIH).
6.2.8. Heparin Release from PEGDA Hydrogels
Hydrogel precursor solutions were prepared containing PEGDA (10-30% w/w,
MW 1-6 kD), Irgacure 2959 (0.1% w/v, 1-[4-[2-Hydroxyethoxy]-phenyl]-2-hydroxy-2- methyl-1-propane-1-one, Ciba Specialty Chemicals, Tarrytown, NY), and heparin (2 mg/ml) dissolved in phosphate buffered saline (PBS, pH 7.4). Hydrogel disks were polymerized in a stainless steel mold (D = 10 mm, H = 1.2 mm) for 10 min under
183 ultraviolet irradiation (365 nm, 0.4-0.5 mW/cm2). The resulting hydrogels were transferred to a 24-well plate containing loading buffer (2 ml/well, 2 mg/ml heparin,
0.1% w/v sodium azide in PBS) to swell and load overnight (12-16 h). A set of control hydrogels (20% w/w PEGDA, MW 1-6 kD) were formed and swollen in the same manner but without heparin. At the start of the release experiments, the loading buffer was removed, the gels were rinsed with PBS, and fresh release buffer (1 ml/well, 0.1% w/v sodium azide in PBS) was added to each hydrogel. The plates were tightly sealed with parafilm and incubated at 37 °C and 80 rpm. At predetermined intervals, the release buffer was completely removed and replaced.
The heparin concentration in the recovered samples was determined using dimethylmethylene blue (DMMB) [40] on the same day the samples were collected.
Release buffer recovered from heparin-free control hydrogels was used as a blank to form a standard curve. Heparin release was obtained by multiplying the sample concentration by the volume of release buffer recovered, which was measured gravimetrically. The overall heparin release was calculated by summation of released heparin over the 34-d experiment.
6.2.9. Characterization of Hydrogel Networks
PEGDA hydrogels were formed as described in the release study, swollen in release buffer for 2 d and the swollen mass determined. Buffer salts were leached in excess distilled water, the gels lyophilized, and the polymer network mass determined.
The mass swelling ratio, q, was calculated as the ratio of the swollen hydrogel and polymer network masses. Since the hydrogels studied here were formed in solution, the method of Bray and Merrill was adapted to calculate ve, the effective cross-linked chains
184 per gram of polymer, using the following equation [41] as previously described (Chapter
4, [34]):
2 ln( 1−++vvps,,) psχ v ps , v = (7) e ⎛⎞1 ⎡ ⎜⎟ ⎤ ⎢⎛⎞vv⎝⎠3 2 ⎛⎞⎥ −−Vvρ ⎜⎟ps,, ⎜⎟ ps sppr, ⎢⎜⎟ ⎜⎟⎥ ⎢⎝⎠vFvpr,, ⎝⎠ pr ⎥ ⎣ ⎦
where:
1 v = (8) ps, ρ p ()q −11+ ρs
1 vpr, = (9) ρ p ⎛⎞1 ⎜⎟−11+ ρs ⎝⎠f
and χ is the Flory-Huggins interaction parameter for PEG and water (0.426) [42], Vs is
3 -1 the molar volume of the solvent, water (18.0 cm mol ), ρ p is the bulk density of PEG
-1 (1.18 g ml )[43], F is the junction functionality (arbitrarily set to 4, see Discussion), ρs is the density of PBS (1.01 g ml-1), and f is the mass fraction of polymer in solution at the time of cross-linking. Because dangling ends and other defects cannot be calculated explicitly in PEGDA systems (as is the case for networks formed from cross-linking of long chains), we have proposed using the number of effective chains, estimated from
* swelling data, per PEGDA molecule at the time of polymerization, denoted ve , to characterize these systems (Chapter 4, [34]).
* vvMWe = e ⋅ PEGDA (10)
185 where MWPEGDA is the number averaged molecular weight of the PEGDA. Ignoring defects, the molecular weight between cross-links, Mc, can be estimated as:
* ve −1 M ce≈=v (11) MWPEGDA
From Mc, the mesh size, ε, of the hydrogel can be estimated using the method of Canal and Peppas [44]:
1 ⎛⎞2 1 M c 3 ε = lC⋅⋅⋅⎜⎟2nps ⋅ v, (12) ⎝⎠M r where l is the weighted average bond length in PEG (1.5 Å) [45], Mr is the PEG repeat
-1 unit (44 g mol ) and Cn is the characteristic ratio for PEG (3.8) [42].
6.2.10. Effects of Heparin Release on SMC Phenotype
PEGDA3k (30% w/w, 0.2 μm filtered) hydrogel discs (D=19 mm, H=1.6 mm) containing heparin (2 mg/ml) were formed between glass plates separated by a silicone sheet with punched circles and prepared as in release studies except the gels were swollen and loaded in a cell compatible loading buffer (DMEM, 2% FBS v/v, 1X gentamycin/amphotericin, 2 mg/ml heparin). Control gels without heparin were prepared in parallel. Hydrogel discs were rinsed in PBS and transferred to transwell culture inserts
(3 µm pore, Corning, Lowell, MA) over confluent, synthetic cultured HCASMCs in a 6- well plate (Fig. 1A). Fresh LSM was added to both chambers. Cultures in LSM supplemented with heparin (400 µg/ml, denoted LSM+H) and SmGM-2 were also maintained as controls for the contractile and synthetic phenotype, respectively. Media were removed and replaced every 2 d. RNA was collected 0, 2, 4, and 6 d after first exposure to the heparin releasing gels. The expression of contractile marker mRNA was determined by real-time RT-PCR. Samples of the media also were collected just before
186 RNA isolation and heparin concentrations were determined using DMMB with LSM as a blank.
6.2.11. Modulation of SMC Phenotype on Heparin Releasing PEGDA Scaffolds
PEGDA3k (30% w/w, 0.2 μm filtered) hydrogel discs (D=25 mm, H=1.6 mm) containing heparin (1 mg/ml) were formed between glass plates. After initial polymerization, PEGDA supplemented with GRGDSP-PEG-acrylate (PEGDA3k: 30 % w/w, GRGDSP-PEG-acrylate: 5 mM, 0.2 μm filtered, 40 μl) was spread evenly on the surface of the hydrogel, covered with a glass plate and polymerized for 10 min to form a thin cell-adhesive hydrogel film linked to the bulk hydrogel (Fig. 1B). Control gels containing the negative control glycosaminoglycan (GAG), chondroitin sulfate (1 mg/ml) were prepared in parallel. Composite constructs were transferred to a 6-well plate, gently secured with a piece of silicone tubing (ID = 2.5 cm), and swollen in DMEM containing heparin or chondroitin sulfate (matching loaded GAG, 1 mg/ml, 3 exchanges) overnight at 37 ºC. Just before seeding, the hydrogels were quickly rinsed with PBS and
HCASMCs were seeded on the constructs in LSM (60,000 cell/cm2). After 4 h, non- adherent cells were rinsed from the construct by replacing the medium with fresh LSM.
RNA samples were collected from the constructs after 3 d.
6.2.12. Real-time Reverse Transcription Polymerase Chain Reaction (RT-PCR)
RNA purification, reverse transcription, and PCR were performed as previously reported (Chapter 5, [38]). Briefly, RNA was purified using RNeasy spin columns
(Qiagen, Valencia, CA). Total RNA yield was determined spectroscopically and RNA was reverse transcribed using the Superscript First Strand Synthesis System (Invitrogen).
The resulting cDNA template was diluted in ultra pure water (final template: 11-25
187
A. Effects of Released Heparin on SMCs (Sect. 6.2.10)
Heparin Releasing Gel Heparin Transwell Insert SMCs
B. Heparin Releasing Scaffolds (Sect. 6.2.11)
SMCs Heparin RGD-containing Gel Heparin Releasing Hydrogel
Figure 6.1-Schematic diagram of heparin release experiments with SMCs. A.) Heparin was released from PEGDA hydrogels through a transwell culture insert. SMCs were seeded below the insert (described in section 6.2.10). B.) Heparin releasing hydrogel scaffolds were constructed from a bulk heparin releasing hydrogel depot that was covered with a thin PEGDA hydrogel film containing GRGDSP-PEG-acrylate (5 mM) to provide cell attachment. SMCs were seeded on the surface of the scaffold (described in section 6.2.11).
188 ng/PCR well assuming 1:1 RNA to cDNA conversion) and was used for real-time PCR analysis using the iQ SYBR Green Supermix (Bio-rad, Hercules, CA) and an iCycler fluorescence detection system (Bio-rad). Gene specific primers for SMαA, calponin,
SM-22α, smooth muscle myosin heavy chain (SM-MHC) and glyceraldehyde-3- phosphate dehydrogenase (GAPDH) were previously reported (Chapter 5, [38]).
Confirmatory western blots were not performed. Previous work has demonstrated that mRNA and protein levels for these markers follow similar expression patterns (Chapter
5, [38]). Primers for myocardin expression were ATGAAGATGCCGTAAAGCAG and
CTTCGGGAAGATCTGGGTAT. Relative transcript abundance was normalized to
GAPDH expression assuming 100% PCR efficiency.
6.2.13. Statistics
Statistical analysis was done using Microsoft Excel and Minitab 15. Data are represented as mean ± standard deviation. Single comparisons were made using an un- paired student’s t-test. Analysis of variance (ANOVA) followed by Tukey’s post hoc test was used for data sets with multiple comparisons. Statistical analysis of real time PCR data was performed in the logarithmic domain. A value of α < 0.05 was considered significant.
6.3. Results
6.3.1. Effects of Heparin on HCASMC Proliferation
Soluble heparin (400 μg/ml) inhibited proliferation of SMCs (Fig. 6.2). After 6 d of culture in 10% v/v FBS in DMEM or SmGM-2, the overall cell population was reduced by 33% or 40%, respectively (p < 0.05), in the presence of heparin. Using growth in LSM as a control for limited proliferation during the culture period, heparin
189
Figure 6.2-Inhibition of cell proliferation by heparin (400 µg/ml, solid bar) but not the negative control, chondroitin sulfate (400 µg/ml, hashed bar), determined by relative MTS absorbance after 6 d of culture in 10% v/v FBS in DMEM (10% FBS), SmGM-2, or 2% v/v FBS in DMEM (2% FBS). *: p < 0.05 compared with no treatment; †: p < 0.05 compared with chondroitin sulfate.
190 inhibited proliferation in 10% FBS and SmGM by 63% and 88%, respectively. The negative control GAG, chondroitin sulfate, did not affect growth in 10% FBS or SmGM, but did slightly increase proliferation in LSM (p < 0.05).
6.3.2. Effects of Heparin on Contractile Phenotype Marker Expression
Soluble heparin (0-1000 μg/ml) increased expression of the contractile phenotype marker proteins SMαA, calponin, and SM-22α (Fig. 6.3). After 6 d of culture, 25 μg/ml heparin (the lowest studied) was sufficient to induce noticeable increases in the level of
SMαA, calponin, and SM-22α protein. The dose response from 25-400 μg/ml heparin was subtle, but at 1000 μg/ml heparin protein expression was increase substantially for all three marker proteins. Marker protein expression was also substantially greater for
SMCs cultured in LSM (all heparin concentrations) than SMCs in SmGM-2. Expression in LSM + chondroitin sulfate was similar to SmGM-2 (not shown).
6.3.3. Heparin Release from PEGDA Gels
PEGDA concentration (10-30% w/w) at the time of polymerization or PEGDA molecular weight (1-6 kD) was use to alter the release kinetics of heparin from PEGDA gels (Fig. 6.4). Increasing PEGDA concentration from 10-30% w/w resulted in extended kinetics of heparin release (Fig. 6.4A) and a longer time required for 90% of heparin to be released from the gels (t90%, Table 6.1) for all PEGDA molecular weights explored.
Decreasing PEGDA molecular weight from 6 to 1 kD resulted in extended kinetics of heparin release (Fig. 6.4B) and a longer t90% (Table 6.1), although these trends were only clear for 30% w/w gels. The time scale of release kinetics, t90%, ranged from 0.7±0.3 to
20.4±4.2 d. Heparin loading in the hydrogels (determined by cumulative heparin release
191
Figure 6.3-Western blot of SMC contractile marker proteins showing dose-dependent up-regulation with increasing heparin concentration in DMEM + 2% v/v FBS. SmGM-2 was used as a control for the synthetic phenotype. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as a loading control.
192
Table 6.1-Summary of heparin release from PEGDA hydrogels
Cumulative t ‡ Composition† Release 90% (d) (μg) 30% PEGDA1k 90.3 ± 21.8 20.4 ± 4.2 20% PEGDA1k 86.9 ± 5.4 6.9 ± 1.1 10% PEGDA1k 26.2 ± 7.1 2.5 ± 1.1 30% PEGDA3k 105.9 ± 6.5 13.4 ± 0.3 20% PEGDA3k 68.4 ± 18.1 3.9 ± 2.2 10% PEGDA3k 70.5 ± 8.5 0.7 ± 0.3 30% PEGDA6k 120.8 ± 20.4 6.8 ± 0.8 20% PEGDA6k 68.5 ± 17.0 4.9 ± 3.8 10% PEGDA6k 33.0 ± 7.4 2.9 ± 2.0
†composition provided as % w/w at the time of polymerization ‡ t90%: time at 90% of cumulative release, determined by linear interpolation
193
Figure 6.4-Heparin release profiles for PEGDA hydrogels of various formulations showing that increasing PEGDA concentration (A: PEGDA3k shown, but PEGDA1k and PEGDA6k followed similar patterns) or decreasing the PEGDA molecular weight (B: 30% w/w PEGDA shown) retards heparin release. The effect of molecular weight was weak for 10 and 20% w/w PEGDA (see Table 6.1). Cumulative heparin release was determined after 34 d for each sample and used to calculate fractional release (see Table 6.1 for cumulative release measurements for each formulation).
194 after 34 d) also varied with PEGDA concentration and molecular weight (Table 6.1).
Loading generally was greatest in gels with high PEGDA concentration and molecular weight.
The mass swelling ratio, q, of the gels was determined and used to characterize the hydrogel networks (Table 6.2). The value of q increased with decreasing PEGDA concentration and increasing molecular weight (Table 6.2). Hydrogels formed from 10% w/w PEGDA1k or PEGDA3k formed flimsy hydrogels that were noticeably weaker than the other hydrogels studied. Heparin loading was negatively correlated with the mass swelling ratio of the hydrogels. The presence of heparin during polymerization did not
* affect the swelling ratio of the hydrogels (p = 0.52). Analysis of ve suggested that the
30% PEGDA hydrogels formed highly entangled networks, whereas 10% PEGDA hydrogels formed weak networks with few entanglements (especially for the PEGDA3k and PEGDA1k) which correlates with the qualitatively weak mechanical properties of these hydrogels (Table 6.2). The estimated mesh sizes of the hydrogels spanned a broad range from 16-185 Å (Table 6.2). Based on these estimates, a mesh size of approximately 50 Å was required to effectively impede heparin release from the hydrogels.
6.3.4. Effect of Heparin Release on HCASMC Phenotype
Heparin released from 30% w/w PEGDA3k gels (Fig. 6.1A) resulted in increased expression of contractile phenotype marker genes in SMCs over a 6 d culture period (Fig.
6.5). Over the first 2 d of the experiment, expression of SMαA, calponin, SM-22α, and smooth muscle myosin heavy chain (SM-MHC) increased in SMCs exposed to heparin
195
Table 6.2-Network properties of PEGDA hydrogels
† ** § *‡ -1 Composition Heparin q ve Mc (g mol ) Mesh Size (Å)
30% PEGDA1k + 3.72 ± 0.03 3.8 ± 0.1 260 ± 10 16.0 ±0.3 20% PEGDA1k + 5.02 ± 0.06 2.6 ± 0.1 380 ± 10 21.4 ±0.5 20% PEGDA1k - 5.14 ± 0.01 2.5 ± 0.0 400 ± 0 22.3 ±0.1 10% PEGDA1k + 11.67 ± 0.17 0.7 ± 0.0 1540 ± 60 57.3 ±1.4 30% PEGDA3k + 7.13 ± 0.06 2.0 ± 0.0 1510 ± 30 48.0 ±0.6 20% PEGDA3k + 9.62 ± 0.18 1.5 ± 0.1 2060 ± 90 62.1 ±1.7 20% PEGDA3k - 9.81 ± 0.09 1.4 ± 0.0 2160 ± 50 63.9 ±0.9 10% PEGDA3k + 26.11 ± 0.77 0.3 ± 0.0 9430 ± 570 185.6 ±7.4 30% PEGDA6k + 8.02 ± 0.08 3.0 ± 0.1 2000 ± 40 57.5 ±0.8 20% PEGDA6k + 10.61 ± 0.12 2.3 ± 0.1 2590 ± 70 71.9 ±1.2 20% PEGDA6k - 10.66 ± 0.09 2.3 ± 0.0 2620 ± 50 72.4 ±0.9 10% PEGDA6k + 19.58 ± 0.29 1.2 ± 0.0 5130 ± 170 124.4 ±2.6
†composition provided as % w/w at the time of polymerization ** “+” indicates presence of 2 mg/ml heparin at the time of polymerization §q calculated as ratio of swollen mass to polymer network mass ‡ * ve is the ratio of effect network chains to PEGDA molecules at polymerization (F = 4)
196 releasing hydrogels (p < 0.05 for calponin, SM-22α, and SM-MHC) and approached the same level as the SMCs in differentiation control medium, LSM+H (p > 0.49 for SMαA, calponin, and SM-22α compared with LSM+H). SMCs cultured with empty gels also increased expression of marker genes, but to a lesser degree than SMCs cultured with heparin releasing gels (p < 0.025 for calponin, SM-22α, and SM-MHC). During the remainder of the experiment (2-6 d), SMCs cultured with heparin releasing hydrogels maintained 2.0-, 2.6-, 1.5-, and 8.1-fold greater expression of SMαA, calponin, SM-22α, and SM-MHC, respectively, than SMCs cultured with empty gels (p < 0.0026). From 2-6 d, the expression level generally plateaued for SMCs cultured with either hydrogel type, while expression continued to increase slightly in the LSM+H control cultures. SMCs cultured in SmGM-2 as controls for the synthetic phenotype, maintained a relatively constant expression that was significantly less than the other conditions (p < 0.022). The heparin concentration in the lower culture chamber, which contained the SMCs, decreased from 48±3 μg/ml after 2 d to 7±1 μg/ml after 6 d (Table 6.3).
The expression of the transcription factor myocardin was distinctly different from the other contractile marker genes (Fig. 6.6). Over the 6 d culture, myocardin mRNA remained constant in SMCs cultured with heparin releasing hydrogels. Myocardin mRNA increased slightly in the synthetic SMCs in SmGM-2 and in the SMCs cultured with empty hydrogels (p < 0.046 vs. heparin hydrogels, 4-6 d). Myocardin mRNA dramatically decreased over the 6 d culture in SMCs cultured in LSM+H and, by 6 d, was at least 3.9-fold less than in any of the other conditions (p < 0.0001).
197
Figure 6.5-Effect of heparin released from PEGDA gels on the expression of contractile phenotype marker genes. SMCs cultured in LSM with heparin releasing hydrogels (Gel + Heparin) showed increased mRNA expression of the markers smooth muscle α-actin (A), calponin (B), SM-22α (C), and smooth muscle myosin heavy chain (SM-MHC, D) compared with unloaded control gels (Empty Gel). LSM + 400 μg/ml heparin (LSM+H) was used as a control for differentiation and SmGM-2 was used as control for the synthetic phenotype. Expression of mRNA was determined using real time RT-PCR. Expression levels relative to 0 d are shown. *: p < 0.05, heparin releasing hydrogel vs. unloaded hydrogel control.
198
Table 6.3-Heparin concentration in medium during transwell insert study
Heparin Concentration Day Culture Condition (µg/ml) † Heparin Releasing Gel 48 ± 3 2 Empty Gel 2 ± 2 Heparin Releasing Gel 11 ± 1 4 Empty Gel 1 ± 0 Heparin Releasing Gel 7 ± 1 6 Empty Gel 1 ± 0 2-6 LSM+H‡ 362 ± 66
†Heparin concentration determined using DMMB with standards made in LSM ‡ LSM+H: DMEM + FBS (2% v/v) + Heparin (400 µg/ml); average for all samples (2, 4, & 6 d) shown
199
Figure 6.6-Effect of heparin released from PEGDA gels on the transcription factor myocardin. LSM + 400 μg/ml heparin (LSM+H) was used as a control for differentiation and SmGM-2 was used as control for the synthetic phenotype. Expression of mRNA was determined using real time RT-PCR. Expression levels relative to 0 d are shown. *: p < 0.05, heparin releasing hydrogel vs. unloaded hydrogel control (same day), †: p < 0.0001, LSM+H vs. all other conditions (same day)
200 6.3.5. Modulation of SMC Phenotype on Heparin Releasing PEGDA Scaffolds
SMCs cultured on GRGDSP-bearing, heparin releasing PEGDA composite scaffolds (Fig. 1B) up-regulated a panel of contractile marker genes (Fig. 6.7).
Expression of SMαA, calponin, SM-22α, and SM-MHC increased by 1.5-, 1.6-, 1.4-, and
1.5-fold, respectively, after 3 d of culture in LSM compared with SMCs cultured on scaffolds loaded with the negative control GAG, chondroitin sulfate (p = 0.061, 0.046,
0.028, and 0.151, respectively).
6.4. Discussion
The goal of this study was to investigate the ability of heparin releasing scaffold systems to actively drive SMC differentiation toward a contractile phenotype. We have previously shown that GRGDSP-bearing PEGDA hydrogels can support rapid and robust expression of markers of contractile phenotype given appropriate external stimuli
(Chapter 5, [38]). By altering the network structure of these hydrogels we sought to expand their utility to also include controlled release of heparin as a mechanism to modulate SMC phenotype.
To assess the potential of heparin to promote contractile SMC phenotype, its effect on SMC proliferation and differentiation was characterized. The effect of soluble heparin on serum stimulated proliferation has been well-established [8, 46]. Inhibition of serum stimulated SMC proliferation (Fig. 6.2) confirmed the activity of our heparin with the cultured HCASMCs used here. The use of heparin to promote expression of contractile marker proteins has been studied less extensively [26], especially in human vascular SMCs. Therefore, we also explored the ability of heparin to promote the re- expression of the contractile marker proteins SMαA, calponin, and SM-22α in cultured
201
Figure 6.7-Expression of markers of contractile SMC phenotype on heparin releasing PEGDA scaffold constructs compared with constructs loaded with the negative control chondroitin sulfate. Expression of mRNA was determined using real time RT-PCR after 3 d of culture. Expression levels relative to chondroitin sulfate controls for each gene are shown. *: p < 0.05 compared with chondroitin sulfate loaded constructs.
202 HCASMCs (Fig. 6.3). The dose-response relationship observed suggested that heparin concentrations near 25 μg/ml were sufficient to induce noticeable changes in cell phenotype, although higher doses (~ 1 mg/ml) could greatly augment this response without any observed cytotoxicity. These results were encouraging since they suggested that a diffusion controlled release system with heparin would be a feasible strategy to modulate SMC phenotype. Using such a system, a high initial heparin concentration can be achieved, but this concentration decreases rapidly over time. Our results suggest that a high initial heparin concentration could shift SMCs toward the contractile phenotype very effectively. Later, lower heparin doses also would promote the contractile phenotype, though to a lesser degree.
PEGDA6k (20% w/w) hydrogels used as scaffold materials in our previous work
(Chapter 5, [38]) and the work of others [37, 39] provided minimal resistance to heparin release (Table 6.1). Altering PEGDA concentration during polymerization and molecular weight were explored as strategies to engineer the hydrogel scaffold system to provide better control of heparin release. By increasing the PEGDA concentration or decreasing the molecular weight, we obtained hydrogels with decreased estimated mesh size (Table
6.2) that extended heparin release by several weeks (Fig. 6.4, Table 6.1).
The library of formulations employed resulted in a range of mesh sizes from 16-
185 Å. Because of the complex network structure of photopolymerized PEGDA hydrogels, it is difficult to estimate the network characteristics accurately. We utilized a model that did not consider all network defects, since it is not possible to explicitly account for these with PEGDA systems, as it is for networks formed from cross-linking long polymer chains. However, we anticipate that our networks contained varying
203 amounts of unreacted acrylate ends and cycles. Furthermore, because the cross-linking nodes of these networks are formed by polyacrylate kinetic chains the junction functionality, F, is generally larger than 4, which was used here to provide consistency with previous studies of PEGDA networks [45]. These issues suggest that the mesh sizes presented here may not estimate the actual network mesh size accurately, but the direction of the error is difficult to predict, since accounting for network defects will tend to decrease and using F > 4 will tend to increase mesh size estimates. Despite these limitations, we found a mesh size of approximately 50 Å was required to effectively impede heparin release from the hydrogels, which correlated well with the 55 Å radius of gyration (as a rough estimate of heparin size), determined by small angle x-ray scattering, for ~20 kD heparin reported by others [47].
However, changes in the network structure also altered the swelling of the hydrogels. Since we chose to swell the hydrogels in heparin containing medium prior to use to facilitate cell culture experiments, the amount of heparin loaded in each formulation also varied as a function of composition (Table 6.1). To optimize the overall heparin delivery, we employed 30% PEGDA3k hydrogels for our studies with SMCs, which balanced extended release kinetics with high heparin loading.
Indirect heparin release studies were utilized to assess both the bioactivity of released heparin and the effect of release-dependent changes in heparin concentration on the expression of contractile marker mRNA. Heparin released from 30% PEGDA3k gels resulted in a significant up-regulation of markers of contractile phenotype compared with unloaded controls (Fig. 6.5). Despite a rapid decrease in heparin concentration after 2 d
(Table 6.3), marker gene expression remained elevated in SMCs exposed to heparin
204 releasing hydrogels, suggesting that the SMCs may be latching into a contractile phenotype expression pattern due to the initial heparin exposure that requires little or no heparin to maintain. These results suggest that long term changes in gene expression may be accomplished using a high initial dose followed by a lower maintenance dose, such as is achieved by diffusion controlled release. However, the fact that SMCs treated with constant high concentration heparin (400 μg/ml) continued to induce marker gene expression suggests that the diffusion-based release profile explored here was sub- optimal.
The expression of the transcription factor myocardin followed a pattern distinct from the other contractile markers (Fig. 6.6) Myocardin interacts with another transcription factor, serum response factor, in smooth muscle cells to induce expression of many SMC marker genes including SMαA, calponin, SM-22α, and SM-MHC, [48, 49] and appears to be regulated, at least in part, at the transcriptional level [50, 51].
Therefore, our observation that myocardin mRNA levels were inversely related to contractile marker expression was unexpected. An alternate activation pathway for
SMαA expression has been described for myofibroblasts that is myocardin-independent
[52]. Since SM-22α expression also has been reported in myofibroblasts [53], it is possible that the SMCs in these studies were trans-differentiating toward this related, but distinct phenotype. However, it is also possible that heparin was driving this or another myocardin-independent pathway in a yet undescribed fashion in cultured human SMCs.
The significance of this altered transcriptional control on the functional phenotype of the
SMCs is unclear, although cells cultured for 6 d in LSM+H did not exhibit contraction in
205 response to carbachol (Chapter 5, [38]), suggesting that this effect results in incomplete modulation toward a truly contractile phenotype.
To form a hydrogel scaffold system, heparin loaded hydrogels were modified with a thin hydrogel film containing the ubiquitous cell binding peptide GRGDSP to provide cell adhesion. Since this film only contributed to ~4% of the construct volume, had the same PEGDA composition as the bulk hydrogel, and was applied before swelling and loading, it is unlikely that this layer altered the release behavior of heparin from the hydrogels. SMCs on these films attached and spread normally and within 3 d showed increased expression of contractile marker proteins. The results suggest this is an effective scaffold system for modulation of cultured SMCs toward a contractile phenotype. Other studies using mechanical and/or biochemical stimulation have reported increases in SMαA [54] and calponin [39] on the order of ~2 fold, but require longer culture durations and complicated bioreactor set-ups. In contrast, the composite scaffold system employed here can provide similar or greater up-regulation of contractile marker genes in 2-3 d without an extensive apparatus. The approach and scaffold design outlined here may substantially enhance the effects of additional mechanical or biochemical stimulation used by others and may act as an effective stand-alone cell- instructive scaffold system for regeneration of functional contractile smooth muscle tissue. If employed as a component of a vascular graft construct, released heparin also may act on adjacent tissues to minimize IH by inhibiting SMC proliferation and promoting contractile phenotype.
206 6.5. Conclusions
Heparin was found to be an effective modulator of SMC phenotype, with the ability to induce expression of markers of contractile SMC phenotype. Manipulation of
PEGDA concentration and molecular weight were effective strategies to engineer
PEGDA scaffold structures with heparin dose and release kinetics sufficient to induce a significant up-regulation of SMC contractile markers. SMCs seeded on cell-instructive
GRGDSP-modified, heparin releasing hydrogels significantly up-regulated contractile marker expression suggesting these scaffold systems may facilitate the formation of contractile smooth muscle tissues in vitro from cultured SMCs.
6.6. Acknowledgments
The project described was supported by Grant Number 5R01EB002067 for the
National Institute of Biomedical Imaging and Bioengineering and Grant Number
1R01HL087843 for the National Heart, Lung, and Blood Institute. JAB also was supported by NIH T32GM07250 and American Heart Association Predoctoral
Fellowship 0715422B.
207 6.7. References
1. Reidy MA. Regulation of Arterial Smooth Muscle Growth. In: Schwartz SM, Mecham RP, editors. The vascular smooth muscle cell : molecular and biological responses to the extracellular matrix. San Diego: Academic Press, 1995. p. xvi, 410 p.
2. Haruguchi H, Teraoka S. Intimal hyperplasia and hemodynamic factors in arterial bypass and arteriovenous grafts: a review. J Artif Organs 2003;6(4):227-235.
3. Brewster LP, Bufallino D, Ucuzian A, Greisler HP. Growing a living blood vessel: insights for the second hundred years. Biomaterials 2007;28(34):5028-5032.
4. Stegemann JP, Hong H, Nerem RM. Mechanical, biochemical, and extracellular matrix effects on vascular smooth muscle cell phenotype. J Appl Physiol 2005;98(6):2321-2327.
5. Hedin U, Daum G, Clowes AW. Heparin inhibits thrombin-induced mitogen- activated protein kinase signaling in arterial smooth muscle cells. J Vasc Surg 1998;27(3):512-520.
6. Pukac L, Huangpu J, Karnovsky MJ. Platelet-derived growth factor-BB, insulin-like growth factor-I, and phorbol ester activate different signaling pathways for stimulation of vascular smooth muscle cell migration. Exp Cell Res 1998;242(2):548-560.
7. Pukac LA, Carter JE, Ottlinger ME, Karnovsky MJ. Mechanisms of inhibition by heparin of PDGF stimulated MAP kinase activation in vascular smooth muscle cells. J Cell Physiol 1997;172(1):69-78.
8. Daum G, Hedin U, Wang Y, Wang T, Clowes AW. Diverse effects of heparin on mitogen-activated protein kinase-dependent signal transduction in vascular smooth muscle cells. Circ Res 1997;81(1):17-23.
9. Rauch BH, Millette E, Kenagy RD, Daum G, Clowes AW. Thrombin- and factor Xa- induced DNA synthesis is mediated by transactivation of fibroblast growth factor receptor-1 in human vascular smooth muscle cells. Circ Res 2004;94(3):340-345.
10. Hayashi K, Saga H, Chimori Y, Kimura K, Yamanaka Y, Sobue K. Differentiated phenotype of smooth muscle cells depends on signaling pathways through insulin- like growth factors and phosphatidylinositol 3-kinase. J Biol Chem 1998;273(44):28860-28867.
11. Bjorkerud S. Effects of transforming growth factor-beta 1 on human arterial smooth muscle cells in vitro. Arterioscler Thromb 1991;11(4):892-902.
208 12. Kim BS, Nikolovski J, Bonadio J, Mooney DJ. Cyclic mechanical strain regulates the development of engineered smooth muscle tissue. Nat Biotechnol 1999;17(10):979-983.
13. Williams B. Mechanical influences on vascular smooth muscle cell function. J Hypertens 1998;16(12 Pt 2):1921-1929.
14. Hedin U, Bottger BA, Forsberg E, Johansson S, Thyberg J. Diverse effects of fibronectin and laminin on phenotypic properties of cultured arterial smooth muscle cells. J Cell Biol 1988;107(1):307-319.
15. Hayward IP, Bridle KR, Campbell GR, Underwood PA, Campbell JH. Effect of extracellular matrix proteins on vascular smooth muscle cell phenotype. Cell Biol Int 1995;19(10):839-846.
16. Thyberg J, Hultgardh-Nilsson A. Fibronectin and the basement membrane components laminin and collagen type IV influence the phenotypic properties of subcultured rat aortic smooth muscle cells differently. Cell Tissue Res 1994;276(2):263-271.
17. Yamamoto M, Yamamoto K, Noumura T. Type I collagen promotes modulation of cultured rabbit arterial smooth muscle cells from a contractile to a synthetic phenotype. Exp Cell Res 1993;204(1):121-129.
18. Bingley JA, Hayward IP, Campbell JH, Campbell GR. Arterial heparan sulfate proteoglycans inhibit vascular smooth muscle cell proliferation and phenotype change in vitro and neointimal formation in vivo. J Vasc Surg 1998;28(2):308-318.
19. Lindner V, Olson NE, Clowes AW, Reidy MA. Inhibition of smooth muscle cell proliferation in injured rat arteries. Interaction of heparin with basic fibroblast growth factor. J Clin Invest 1992;90(5):2044-2049.
20. Castellot JJ, Choay J, Lormeau JC, Petitou M, Sache E, Karnovsky MJ. Structural Determinants of the Capacity of Heparin to Inhibit the Proliferation of Vascular Smooth-Muscle Cells .2. Evidence for a Pentasaccharide Sequence That Contains a 3-O-Sulfate Group. J Cell Biol 1986;102(5):1979-1984.
21. Garg HG, Thompson BT, Hales CA. Structural determinants of antiproliferative activity of heparin on pulmonary artery smooth muscle cells. Am J Physiol Lung Cell Mol Physiol 2000;279(5):L779-789.
22. Millette E, Rauch BH, Defawe O, Kenagy RD, Daum G, Clowes AW. Platelet- derived growth factor-BB-induced human smooth muscle cell proliferation depends on basic FGF release and FGFR-1 activation. Circ Res 2005;96(2):172-179.
23. Savage JM, Gilotti AC, Granzow CA, Molina F, Lowe-Krentz LJ. Antibodies against a putative heparin receptor slow cell proliferation and decrease MAPK activation in vascular smooth muscle cells. J Cell Physiol 2001;187(3):283-293.
209 24. Castellot JJ, Wong K, Herman B, Hoover RL, Albertini DF, Wright TC, Caleb BL, Karnovsky MJ. Binding and Internalization of Heparin by Vascular Smooth-Muscle Cells. J Cell Physiol 1985;124(1):13-20.
25. Patel RC, Handy I, Patel CV. Contribution of double-stranded RNA-activated protein kinase toward antiproliferative actions of heparin on vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 2002;22(9):1439-1444.
26. Hashimoto T, Kihara M, Sato K, Imai N, Tanaka Y, Sakai M, Tamura K, Hirawa N, Toya Y, Kitamura H, Umemura S. Heparin recovers AT1 receptor and its intracellular signal transduction in cultured vascular smooth muscle cells. FEBS Lett 2005;579(1):281-284.
27. Orlandi A, Ropraz P, Gabbiani G. Proliferative Activity and Alpha-Smooth Muscle Actin Expression in Cultured Rat Aortic Smooth-Muscle Cells Are Differently Modulated by Transforming Growth-Factor-Beta-1 and Heparin. Exp Cell Res 1994;214(2):528-536.
28. Stegemann JP, Nerem RM. Altered response of vascular smooth muscle cells to exogenous biochemical stimulation in two- and three-dimensional culture. Exp Cell Res 2003;283(2):146-155.
29. Burdick JA, Anseth KS. Photoencapsulation of osteoblasts in injectable RGD- modified PEG hydrogels for bone tissue engineering. Biomaterials 2002;23(22):4315-4323.
30. Hern DL, Hubbell JA. Incorporation of adhesion peptides into nonadhesive hydrogels useful for tissue resurfacing. J Biomed Mater Res 1998;39(2):266-276.
31. Mann BK, Gobin AS, Tsai AT, Schmedlen RH, West JL. Smooth muscle cell growth in photopolymerized hydrogels with cell adhesive and proteolytically degradable domains: synthetic ECM analogs for tissue engineering. Biomaterials 2001;22(22):3045-3051.
32. Nuttelman CR, Tripodi MC, Anseth KS. In vitro osteogenic differentiation of human mesenchymal stem cells photoencapsulated in PEG hydrogels. J Biomed Mater Res A 2004;68(4):773-782.
33. Hwang NS, Varghese S, Zhang Z, Elisseeff J. Chondrogenic differentiation of human embryonic stem cell-derived cells in arginine-glycine-aspartate-modified hydrogels. Tissue Eng 2006;12(9):2695-2706.
34. Beamish JA, Zhu J, Kottke-Marchant K, Marchant RE. The effects of monoacrylated poly(ethylene glycol) on the properties of poly(ethylene glycol) diacrylate hydrogels used for tissue engineering. J Biomed Mater Res 2009;in press.
210 35. Peyton SR, Raub CB, Keschrumrus VP, Putnam AJ. The use of poly(ethylene glycol) hydrogels to investigate the impact of ECM chemistry and mechanics on smooth muscle cells. Biomaterials 2006;27(28):4881-4893.
36. Fittkau MH, Zilla P, Bezuidenhout D, Lutolf MP, Human P, Hubbell JA, Davies N. The selective modulation of endothelial cell mobility on RGD peptide containing surfaces by YIGSR peptides. Biomaterials 2005;26(2):167-174.
37. Mann BK, West JL. Cell adhesion peptides alter smooth muscle cell adhesion, proliferation, migration, and matrix protein synthesis on modified surfaces and in polymer scaffolds. J Biomed Mater Res 2002;60(1):86-93.
38. Beamish JA, Fu A, Choi A, Haq N, Kottke-Marchant K, Marchant RE. The influence of RGD-bearing hydrogels on the re-expression of contractile vascular smooth muscle cell phenotype. Biomaterials 2009;in press.
39. Bulick AS, Munoz-Pinto DJ, Qu X, Mani M, Cristancho D, Urban M, Hahn MS. Impact of endothelial cells and mechanical conditioning on smooth muscle cell extracellular matrix production and differentiation. Tissue Eng Part A 2009;15(4):815-825.
40. Farndale RW, Buttle DJ, Barrett AJ. Improved quantitation and discrimination of sulphated glycosaminoglycans by use of dimethylmethylene blue. Biochimica et Biophysica Acta (BBA) - General Subjects 1986;883(2):173-177.
41. Bray JC, Merrill EW. Poly(vinyl alcohol) hydrogels. Formation by electron beam irradiation of aqueous solutions and subsequent crystallization. Journal of Applied Polymer Science 1973;17:3779-3794.
42. Merrill EW, Dennison KA, Sung C. Partitioning and diffusion of solutes in hydrogels of poly(ethylene oxide). Biomaterials 1993;14(15):1117-1126.
43. Lin HQ, Kai T, Freeman BD, Kalakkunnath S, Kalika DS. The effect of cross- linking on gas permeability in cross-linked poly(ethylene glycol diacrylate). Macromolecules 2005;38(20):8381-8393.
44. Canal T, Peppas NA. Correlation between Mesh Size and Equilibrium Degree of Swelling of Polymeric Networks. Journal of Biomedical Materials Research 1989;23(10):1183-1193.
45. Cruise GM, Scharp DS, Hubbell JA. Characterization of permeability and network structure of interfacially photopolymerized poly(ethylene glycol) diacrylate hydrogels. Biomaterials 1998;19(14):1287-1294.
46. Garg HG, Mrabat H, Yu L, Freeman C, Li B, Zhang F, Linhardt RJ, Hales CA. Significance of the 2-O-sulfo group of L-iduronic acid residues in heparin on the growth inhibition of bovine pulmonary artery smooth muscle cells. Carbohydr Res 2008;343(14):2406-2410.
211 47. Pavlov G, Finet S, Tatarenko K, Korneeva E, Ebel C. Conformation of heparin studied with macromolecular hydrodynamic methods and X-ray scattering. Eur Biophys J Biophys Lett 2003;32(5):437-449.
48. Wang Z, Wang DZ, Pipes GC, Olson EN. Myocardin is a master regulator of smooth muscle gene expression. Proc Natl Acad Sci U S A 2003;100(12):7129-7134.
49. Miano JM. Serum response factor: toggling between disparate programs of gene expression. J Mol Cell Cardiol 2003;35(6):577-593.
50. Yoshida T, Kawai-Kowase K, Owens GK. Forced expression of myocardin is not sufficient for induction of smooth muscle differentiation in multipotential embryonic cells. Arterioscler Thromb Vasc Biol 2004;24(9):1596-1601.
51. Mack CP, Hinson JS. Regulation of smooth muscle differentiation by the myocardin family of serum response factor co-factors. J Thromb Haemost 2005;3(9):1976-1984.
52. Gan Q, Yoshida T, Li J, Owens GK. Smooth muscle cells and myofibroblasts use distinct transcriptional mechanisms for smooth muscle alpha-actin expression. Circ Res 2007;101(9):883-892.
53. Faggin E, Puato M, Zardo L, Franch R, Millino C, Sarinella F, Pauletto P, Sartore S, Chiavegato A. Smooth muscle-specific SM22 protein is expressed in the adventitial cells of balloon-injured rabbit carotid artery. Arterioscler Thromb Vasc Biol 1999;19(6):1393-1404.
54. Stegemann JP, Nerem RM. Phenotype modulation in vascular tissue engineering using biochemical and mechanical stimulation. Ann Biomed Eng 2003;31(4):391- 402.
212 CHAPTER 7: Conclusions and Future Directions
7.1. Summary and Conclusions of Completed Work
This work investigated the hypothesis that smooth muscle cell (SMC) phenotype can be modulated by a cell-instructive hydrogel scaffold. A photopolymerizable poly(ethylene glycol) (PEG)-based hydrogel system was employed as the scaffold material. The properties of this system can be controlled across a wide range of length scales, from the nm-sized network mesh to the cm-sized bulk shape of the gels. PEG also resists protein adsorption and non-specific cell attachment. The inclusion of ligands in the network can mediate specific cell interactions that can be controlled quantitatively.
In Chapter 4, networks formed from copolymerization of PEG diacrylate
(PEGDA) with PEG monoacrylate (PEGMA, models for ligand-network tethers) were characterized. These experiments showed that PEGDA and PEGMA can be copolymerized with independent control of PEGMA-tethered ligand concentration and bulk hydrogel properties. Analysis of these networks also provided new insight into their structure, suggesting a large contribution of chain entanglements to the hydrogel properties. The quantitative control of PEGDA hydrogel physical properties demonstrated suggests these systems have potential as a versatile scaffold material, at least from a non-biologic perspective.
In Chapter 5, biological characteristics of PEGDA scaffolds were explored, as they relate to smooth muscle tissue engineering. The ability of GRGDSP-bearing
PEGDA hydrogels (RGD-gels) to support the differentiation of cultured SMCs toward a contractile phenotype was quantified. Cell interactions with these scaffolds were highly
RGD-specific, and this biomimetic cell-scaffold interaction did not affect the ability of
213 cultured SMCs to re-express markers of contractile phenotype, driven by external stimulation. Marker proteins were rapidly up-regulated at the level of mRNA and protein and were organized into filaments within the cells on RGD-gels. The ability of other extracellular matrix-derived, cell-adhesive peptides, including YIGSR, VAPG,
KQAGDV, GKDGEA, and GSWSGSPPRRARVT, to modulate phenotype marker expression was explored, but these studies were precluded by a lack of specific SMC attachment to hydrogels bearing these peptides. These studies indicated that PEGDA scaffolds with pendant RGD ligands can support the differentiation of cultured SMCs toward a contractile phenotype, given the appropriate external stimuli.
In Chapter 6, RGD-bearing PEGDA hydrogel scaffolds were engineered to provide controlled release of heparin, a factor that can drive SMC differentiation.
Expression of markers of contractile phenotype were rapidly up-regulated on these bioactive scaffolds, driven by released heparin with limited external stimulation. These results directly support the hypothesis that SMC phenotype can be modulated by a cell- instructive hydrogel scaffold, and suggest that the use of soluble signaling factors, such as heparin, is an effective approach to induce rapid changes in SMC phenotype.
The development of a bioactive, heparin releasing PEGDA hydrogel construct represents a step forward in the evolution of cell-instructive scaffold systems for smooth muscle tissue engineering. However, significant work remains to develop a system that can be used for vascular tissue engineering applications in vivo. Elements of this system that require additional engineering to meet design requirements for this application will be discussed below. Furthermore, this work has also raised questions about the fundamental biology of SMC phenotype modulation, especially as it relates to phenotype
214 switching from synthetic, cultured SMCs toward a contractile phenotype. Potential studies exploring this topic also are proposed below.
7.2. Engineering of Improved Scaffold Systems
The heparin-releasing hydrogel scaffold system has several limitations that warrant continued development. Improvements can be made in the following areas: 1) improved control of heparin release, 2) investigation of other contractile phenotype inducing factors, 3) engineering of cell-responsive degradation into the hydrogel network, and 4) developing approaches for 3D fabrication of scaffold systems for vascular prostheses (Fig. 7.1).
7.2.1. Improved Control of Heparin Release
Adjusting the PEGDA hydrogel mesh size by using molecular weight and concentration was effective for extending heparin release. However, this strategy sacrifices independent control of heparin release kinetics, heparin loading, and hydrogel mechanical properties. Hydrogel formulations that extended heparin release also were stiff, had low swelling, and had low heparin loading. Stiff, low swelling gels may not be optimal for cell infiltration. Degradable systems with similar mesh sizes may be resorbed more slowly. Also, low heparin loading limited heparin delivery. Heparin concentrations of approximately 50 μg/ml were obtained, which were still effective, but well below doses where significantly larger contractile marker up-regulation was observed (~1000 μg/ml) (Chapter 6). There are several approaches that could be explored to enhance heparin delivery in these systems:
1) Increase heparin loading during polymerization. Hydrogel loading of 1-2 mg/ml of heparin was chosen to minimize concentration dependent cytotoxicity at the
215
Current Design
A. Enhance/extend D. Integrate with existing heparin release graft materials B. Investigate other C. Engineer cell- bioactive factors responsive degradation
Tissue Engineered Vascular Graft
LEGEND: PEGDA Microparticle Graft material Drug carrier (e.g. ePTFE) + endothelium Tethered ligand Heparin “Synthetic” SMC Degradable Other bioactive PEGDA “Contractile” factor SMC
Figure 7.1-Potential future design improvements for SMC phenotype modulating scaffolds. The current design consists of an RGD-bearing hydrogel with bulk diffusion controlled release of heparin to induce changes in SMC phenotype. Areas requiring additional development include A) enhanced and extended heparin release, B) investigation of additional bioactive factors to improve phenotype modulation, C) engineering of degradable PEGDA derivatives to allow SMC in-growth and remodeling, and D) integrating scaffold systems with existing graft materials (such as ePTFE). Ultimately future iterations of this design can be incorporated into a tissue-engineered graft construct suitable for in vivo experiments.
216 cell-hydrogel interface, where heparin would initially approach the loaded concentration.
The results of recent studies (not shown) have indicated that heparin concentrations up to
3.2 mg/ml are cytocompatible during long-term culture suggesting that initial heparin incorporation (during polymerization and loading) can be increased several times higher than described here. Low heparin loading concentrations (1-2 mg/ml) also do not interfere with hydrogel polymerization (Chapter 6). Additional heparin may affect the gel network properties, introducing another co-dependent variable in the system, which would further limit independent control of release, loading, and hydrogel properties.
2) Introduce a heparin carrier for controlled release of heparin. In the studies presented here, gel formulations explored with mesh sizes greater than ~50 Å displayed minimal resistance to heparin release. These hydrogel formulations would be excellent carriers for heparin containing microparticles. The microparticles would provide the rate limiting step for heparin release since the hydrogel imparts little additional resistance.
This strategy has been employed for TGF-β1 release from microparticles entrapped within PEG-based hydrogels [1, 2] and, by incorporating a variety of particle types, could facilitate the release of multiple bioactive factors with independent release patterns [3].
This approach would permit better independent control of hydrogel properties and release kinetics, provided microparticles have a minimal effect on hydrogel cross-linking and mechanical properties. Furthermore, an increase in the hydrogel mesh size would improve transport of other critical nutrients through the gels and would be consistent with
PEGDA materials that others have utilized for 3D SMC cultures [4, 5].
This strategy would also allow independent control of heparin loading. However, since the effective dose of heparin is relatively high (for example, 1-2 orders of
217 magnitude larger than TGF-β1), it may be difficult to achieve a degree of loading higher than the current system. For example, to achieve 2 mg/ml heparin in the final formulation the composition of heparin in the particles may need to be 5% or higher
(assumptions: microparticles are incorporated into 20% PEGDA hydrogels at a ratio of
1:5 w:w particles to PEGDA, similar to [2]). This degree of loading may be difficult to achieve without compromising control of release kinetics. There are several strategies to generate heparin loaded carrier particles. Poly(lactide-co-glycolide) microspheres have been utilized for a variety of drug delivery applications [1]. Emulsification followed by photopolymerization of low molecular weight PEGDA could also be used to form highly cross-linked hydrogel microparticles, in a fashion similar to that employed for other photopolymerizable hydrogel systems [6, 7], provided effective emulsification solvents can be found.
3) Covalently link heparin to the network for continuous signaling. Several groups have reported covalent modification of polysaccharides that allows for copolymerization with PEGDA [8-10]. This would immobilize heparin to the network.
For other cell types, such as valvular interstitial cells, heparin remains active and able to modulate cell behavior [9]. Since the mechanism of heparin’s action in SMCs is not well understood and may involve heparin internalization and intracellular signaling [11, 12], it is unclear if this approach would be effective in modulating SMC behavior. However, because it cannot be internalized, scaffolds with immobilized heparin may help to elucidate the signaling pathways that are most important for heparin-induced modulation of SMC phenotype.
218 7.2.2. Exploration of Additional Bioactive Factors
As described in Chapter 5, while dramatically bringing their marker profile closer to a contractile phenotype, SMCs cultured for 6 d in LSM+H (DMEM, 2% v/v FBS, 400
μg/ml heparin) did not show evidence of contraction in response to carbachol.
Furthermore, the overall level of expression of smooth muscle myosin heavy chain (SM-
MHC) mRNA was significantly lower than the other markers and the SM-MHC protein, a critical component of the contractile machinery, was not detectable by western blotting or immunofluorescence. While an import next step is to screen additional contraction- inducing compounds (KCl, endothelin-1, phenylephrine, etc), these observations suggest the contractile apparatus was not reformed completely. More work is needed to restore a truly contractile phenotype to cultured SMCs. One approach to accomplish this goal is co-stimulation with additional factors to enhance SMC modulation toward a contractile phenotype. For example, transforming growth factor β1 (TGF-β1) has also been shown to enhance the expression and organization of smooth muscle α-actin (SMαA) and other more rigorous differentiation markers such as SM-MHC and SM-22α in SMC lines as well as primary rat and human SMC cultures [13-17]. We also have conducted preliminary studies with TGF-β1 suggesting it can up-regulate expression of SMαA in cultured human SMCs using a cell-based ELISA method (Fig. 7.2). The use of other soluble signals, such as angiotensin II, or mechanical stimulation may further enhance development of contractile phenotype in cultured SMCs.
7.2.3. Cell-Mediated Hydrogel Degradation
The PEGDA scaffold system described in this work is essentially non-degradable
(networks are stable in neutral aqueous buffers for several months, unpublished
219
Figure 7.2-Expression of smooth muscle α-actin (SMαA) as a function of TGF-β1 concentration. HCASMCs were cultured for 6 d in DMEM + 2% FBS + TGF-β1. Expression of SMαA was assessed using a cell-based ELISA method and normalized by DNA content determined by CyQUANT. Cell-based ELISA yields relative results only, so units are arbitrary. Results were normalized to 0 ng/ml TGF-β1.
220 observations). Since the mesh size of these networks is several orders of magnitude less than the size of a cell (ε < ~100 Å, cells ~ 10 μm), cell penetration through non- degradable PEGDA networks is minimal. Encapsulated cells take on a rounded morphology which is not characteristic of native SMCs [4]. Therefore, to allow 3D
SMC in-growth and remodeling, degradation must be engineered into the PEGDA.
PEGDA hydrogels have been created that are hydrolytically [18, 19] or proteolytically degradable [20-22]. Proteolytically degradable gels inherently respond to local cellular action, limiting degradation to sites of cellular invasion and allowing the unpopulated matrix to remain intact for as long as is necessary for cells to invade it. Such gels typically contain a peptide that is sensitive to cleavage by selected matrix metalloproteinases (MMP) along the PEG backbone of the PEGDA molecule (Fig. 7.3).
After cleavage, the PEGDA molecules eventually cease to form a polymer network.
MMP-1 (Collagenase-1) has been well characterized and is expressed by SMCs [23, 24].
Therefore, peptide sequences sensitive to this enzyme are an excellent choice for incorporation into degradable PEGDA.
Initial studies using a collagenase sensitive PEGDA derivative (CS-PEGDA) with a centrally located, MMP sensitive peptide (GPQGIAGQ from Collagen Type I, α1 chain) have been conducted showing time-dependent degradation of these hydrogels by
Clostridial collagenase (1 mg/ml, Fig. 7.3B). However, this approach has several important limitations. First, the CS-PEGDA macromer initially tested in these studies formed networks that could not be completely degraded at concentrations higher than
10% (not shown). This suggests this material also contained non-degradable PEGDA, either as a result of impure peptide or side reactions during synthesis. Second, further
221
H H O N N O A O GPQGIAGQK O O n O O n O
B 125 100 75 50
% Initial Mass Initial % 25 0 0 5 10 15 20 Time (h)
Figure 7.3-A) One possible collagenase sensitive-PEGDA (CS-PEGDA) molecule with a central peptide sensitive to cleavage by MMP-1 linked to the PEG chains via its N- terminus and a C-terminal lysine (as shown) or diaminopropionic acid. B) Degradation of 10% w/v CS-PEGDA hydrogels using Clostridial collagenase (which is also capable of cleaving the peptide sequence GPQGIAGQ). Legend: =10% CS-PEGDA hydrogel + 2 μg/ml collagenase, =10% CS-PEGDA hydrogel (no collagenase added). (Acknowledgement: J. Zhu synthesized this degradable material).
222 attempts to synthesize this material from HPLC purified GPQGIAGQK peptide, using N- hydroxysuccinimide functionalized PEGMA (as is used to form pendant peptide conjugates in Chapters 5 and 6), failed to show di-functional conjugation to both the N- terminal amine and the pendant amine in the terminal lysine residue. Because the pKa of these amines differ, adjustments in the pH of the conjugation reaction (currently set at pH
= 8.4) may facilitate conjugation at both sites. The peptide also could be re-synthesized with a c-terminal diaminopropionic acid residue instead of lysine, which will bring the pKa of the terminal amines into closer alignment. Furthermore, any trifluoroacetic acid- amine complexes (that may block subsequent conjugation) can be dissociated with acetic acid.
However, even with these improvements, CS-PEGDA-based hydrogels may have limitations for tissue engineering applications. Although some studies suggest migration through CS-PEGDA materials may occur [20, 21], cell morphology and histology in these scaffolds used for smooth muscle applications show poorly spread, rounded cells without formation of smooth muscle-like tissue architectures [21, 25]. Degradation of
CS-PEGDA materials has often been tested using high concentrations of bacterial collagenase [21]. It is unclear if degradation results from these studies provide relevant data regarding the local, cell-mediated degradation of these networks.
Poor tissue development in CS-PEGDA materials contrasts with results obtained using PEG-based hydrogels, cross-linked by Michael-type addition of cysteine containing peptides to vinyl sulfone-terminated multi-arm PEG, developed by Lutolf, Hubbell, and co-workers. These hydrogels allow extensive cell spreading and tissue organization using similar enzymatically degradable peptide sequences [26, 27]. There are several
223 possible explanations for these differences. First, the multi-arm PEG gels form weaker networks than PEGDA systems, with typical gels having approximately 2-fold greater swelling and nearly an order of magnitude weaker elastic modulus [28, 29]. These loose networks are more easily degraded, allowing for cell infiltration. These networks also employed several collagenase sensitive sequences, with varying degrees of collagenase sensitivity [27]. The best invasion rates were obtained with weak gels and a highly labile degradable peptide (GPQGIWGQ) [27]. Since the CS-PEGDA system utilizes intermediately labile peptides (such as GPQGIAGQ) and forms highly cross-linked networks, it is not surprising that results have been poor. Incorporation of more highly labile collagenase sensitive peptides, such as those recently reported [30], may improve the cell-mediated degradation of CS-PEGDA systems. Investigation of more loosely cross-linked CS-PEGDA systems or other hydrogel systems will be necessary to obtain useful, 3D scaffold systems for smooth muscle tissue engineering. Also, if such weak gel systems are required for SMC infiltration and tissue development, delivery of bioactive molecules, such as heparin, from the scaffold bulk likely would not be feasible, necessitating the development of microparticle-based drug delivery carriers.
7.2.4. Scaffold Integration with Existing Graft Materials
In preparation for 3D in vitro bioreactor studies and in vivo testing of hydrogel scaffolds, these materials must be incorporated into existing vascular grafts. The calculated compliance [31] of hydrogel tubes (20% w/w PEGDA, MW = 6k, D = 4 mm, wall = 0.5 mm) is several fold greater than native vasculature [32] suggesting this material is too weak to serve as a stand-alone scaffold for a vascular prosthesis (Fig. 7.4).
Furthermore, the ultimate strength of PEGDA, though not measured, is qualitatively far
224
Figure 7.4-Calculated compliance of vascular graft materials. Compliance was calculated as described [31] based on measured elastic moduli of ePTFE (for materials with a range of intermodal distances, IND) and PEGDA (MW = 6k, 20% w/w) assuming a diameter of 4 mm and a wall thickness of 0.5 mm and a Poisson’s ratio of 0.5. †Measurements for saphenous vein and muscular arteries were obtained from the literature [32].
225 less than expanded poly(tetrafluoroethylene) (ePTFE), a common graft material, suggesting these tubes also would have inadequate burst strength.
To address this issue, PEGDA scaffolding can be copolymerized in and around a more mechanically robust material to reinforce the gel (Fig. 7.5). Since compliance matching is thought to be an important criteria for minimizing intimal hyperplasia (IH)
[33], ePTFE materials with large internodal distances (IND), which have better compliance matched mechanical properties (Fig. 7.4) could be utilized for reinforcement.
Normally, high IND ePTFE leads to plasma leakage. The pores are eventually filled with fibrin, but this repair matrix may not stimulate appropriate healing responses, also contributing to IH. PEGDA scaffolds can fill these pores to prevent leakage and exclude fibrin, as well as act as instructive scaffolds for development of contractile, non- hyperplastic tissue. An important next step in this work is the development of a robust protocol for incorporation of bioactive PEGDA scaffold materials in high IND ePTFE grafts.
7.3. Modulation of Cultured SMCs Toward a Contractile Phenotype
The modulation of cultured SMCs toward a contractile phenotype has been studied in less detail than SMC de-differentiation. A better understanding of this process will facilitate the development of tissue engineering approaches that result in functional contractile smooth muscle tissue. Several observations in the course of the work presented here have raised important questions about the biology of this process.
7.3.1. Correlation of Focal Adhesions and Contractile Marker Expression
In Chapter 5, the role of the extracellular matrix proteins fibronectin (FN) and laminin (LN) in the re-differentiation of cultured SMCs toward a contractile phenotype
226
AB C
Figure 7.5-Coating of the adventitial surface of an ePTFE graft material with PEGDA hydrogel. The graft material is coated with surfactant (which may be a biomimetic surfactant polymer) to facilitate infiltration of PEGDA within the graft pores. The graft is placed on a mandrel and transferred to a solution of PEGDA in a second tube with the desired outer diameter. The PEGDA solution is photopolymerized and the hydrogel graft material is removed from the tube and the mandrel. (Acknowledgement: J. Zhu developed this procedure and prepared these photographs).
227 was explored. It has been well established that primary SMCs seeded on FN lose their contractile phenotype more quickly than cells seeded on LN [34-38]. However, in our study, we noted that after a 14-17 h seeding period in serum-free medium, cultured SMCs plated on LN expressed a lower level of SMαA and calponin than SMCs plated on FN.
After the addition of low serum medium, the differences disappeared. Under these conditions with cultured SMCs, FN appeared to promote SMC modulation toward a contractile phenotype whereas LN appeared to inhibit it, the opposite as observed for primary cells. We also observed that during the seeding process SMCs spread poorly on
LN, and after 4 h, SMC attachment to LN was not strong enough to withstand a change in medium. The observed differences in SMC gene expression on LN may be due to changes in LN-cell signal transduction in cultured SMCs compared with primary SMCs, as has been observed with other signals such as IGF-1 [39]. However, we hypothesize that increased contractile marker gene expression on FN compared with LN is due to differences in the initial degree of attachment to the substrates, which may be abrogated as the SMCs produce their own endogenous FN matrix.
During phenotypic modulation from a contractile to synthetic phenotype such as occurs in culture, SMCs down-regulate LN and LN receptors, such as α7β1 integrin, while up-regulating the expression of FN and FN receptors, such as α5β1 and αvβ3 [40,
41]. Whereas primary cells may be competent to interact with LN, cultured SMCs may have limited interactions with LN. This change may explain the poor initial attachment to LN in our experiments. Eventually, these SMCs spread on LN and, by 14 h, are indistinguishable from SMCs plated on FN. Others have shown that cultured baboon
SMCs plated on LN developed a well-formed provisional FN matrix after 4 h of culture,
228 effectively masking the LN substrate [42]. Although we have not measured FN production yet, our morphological observations were consistent with our human SMCs also organizing a FN matrix within the first few hours of culture. To test the hypothesis that attachment to LN was initially poor but improved with extended culture duration, we quantified the development of focal adhesions on FN, LN, and RGD-bearing hydrogels
(RGD-gels) in serum-free medium over the first 48 h of culture. Preliminary results indicate that focal adhesion number and area are initially low on LN, but increase to levels indistinguishable from FN after 2 d (Fig. 7.6). Focal adhesions develop rapidly on
FN and RGD-gels then remain essentially unchanged.
In addition to replicating these results, an important next step is to characterize the expression and organization of the underlying FN matrix to confirm that the expression of endogenous FN on LN and RGD-gels is correlated with the development of focal adhesions. The degree of intracellular activation of focal adhesion kinase (FAK) should also be measured to confirm that focal adhesion development, determined by counting methods, is correlated with differences in intracellular signaling. Gene expression of contractile marker proteins and endogenous FN should also be correlated with focal adhesion development. It has been widely recognized that intracellular tension, afforded by focal adhesion development, is an important regulator of cell behavior [43] and F- actin dependent rho-A signaling has been shown to increase the expression of SMαA and
SM-22α [44]. Recently it has been shown that intracellular tension, modulated by focal adhesion (FA) size, among other methods, also regulates the incorporation of SMαA into stress fibers in myofibroblasts [45]. This work may provide new insights into the influence of ECM on synthetic-to-contractile phenotype modulation in cultured SMCs.
229
Figure 7.6-Time-dependent focal adhesion (FA) development on fibronectin (FN), laminin (LN), and RGD (5 mM) containing PEGDA (20% w/w) hydrogels (RGD-Gel) showing A) number of focal adhesions per cell and B) focal adhesion area per cell area. FA formation on LN lags behind FN and RGD-gels. Briefly, SMCs were seeded in serum-free medium onto FN (1 μg/cm2), LN (2 μg/cm2), or RGD-Gel coated glass coverslips and cultured in serum-free medium for 2 d. At 4, 12, 24, and 48 h after seeding SMCs were fixed with 4% paraformaldehyde and stained for vinculin. Images were filtered for low spatial frequency staining, thresholded, and FA number and area were determined. N = 25 cells per condition and timepoint. (Acknowledgement: AJ Choi performed this data collection and analysis).
230 The results of this work may have important implications for the design of scaffolds for smooth muscle tissue engineering. SMCs that would likely populate a regeneration-inducing TEBV in vivo (or one prepared ex vivo with cultured SMCs) would also adopt a synthetic phenotype, in order to migrate and proliferate in the scaffold.
Scaffold-cell interactions might be best designed to accommodate a synthetic-to- contractile switch, rather than inhibit initial de-differentiation to the synthetic phenotype.
Our results suggest that fibronectin or fibronectin-mimicking scaffolds, such as the RGD- gels described here, may be superior for this purpose. Continued work on this subject will better elucidate the role of scaffold-cell interactions in re-differentiation toward the contractile phenotype.
7.3.2. Myocardin in Contractile Marker Re-expression in Cultured SMCs
In experiments described in Chapter 6, we observed that myocardin mRNA levels were inversely related to contractile marker expression. This observation was unexpected because myocardin is thought to be an important activator of expression of many SMC marker genes including SMαA, calponin, SM-22α, and SM-MHC, [46, 47] and appears to be regulated, at least in part, at the transcriptional level [48, 49]. An alternate activation pathway for SMαA expression has been described for myofibroblasts that is myocardin-independent [50]. The myocardin family of transcription factors also contains additional members with close homology to myocardin [49]. In particular, myocardin related transcription factor A (MRTF-A, also known as MAL and MKL-1) is thought to be capable of modulating SMC marker gene expression [49]. Unlike myocardin, which is confined to the nucleus, MRTF-A exists in both the nuclear and cytoplasmic compartments. Its partitioning may be related, in part, to its interaction with cytoplasmic
231 G-actin [49]. MRTF-A’s relationship with the cytoskeleton, our observation that poorly adhered SMCs on LN have reduced marker gene expression, and our observation that myocardin does not appear to be the major factor driving gene expression in our cultured
SMCs, suggest MRTF-A may play an important role in marker re-expression. Analysis of MRTF-A localization and activity may improve the understanding of transcriptional control of contractile marker re-expression in cultured SMCs.
232 7.4. References
1. DeFail AJ, Chu CR, Izzo N, Marra KG. Controlled release of bioactive TGF-beta(1) from microspheres embedded within biodegradable hydrogels. Biomaterials 2006;27(8):1579-1585.
2. Holland TA, Tabata Y, Mikos AG. In vitro release of transforming growth factor- beta 1 from gelatin microparticles encapsulated in biodegradable, injectable oligo(poly(ethylene glycol) fumarate) hydrogels. J Control Release 2003;91(3):299- 313.
3. Holland TA, Tabata Y, Mikos AG. Dual growth factor delivery from degradable oligo(poly(ethylene glycol) fumarate) hydrogel scaffolds for cartilage tissue engineering. J Control Release 2005;101(1-3):111-125.
4. Bulick AS, Munoz-Pinto DJ, Qu X, Mani M, Cristancho D, Urban M, Hahn MS. Impact of endothelial cells and mechanical conditioning on smooth muscle cell extracellular matrix production and differentiation. Tissue Eng Part A 2009;15(4):815-825.
5. Mann BK, West JL. Cell adhesion peptides alter smooth muscle cell adhesion, proliferation, migration, and matrix protein synthesis on modified surfaces and in polymer scaffolds. J Biomed Mater Res 2002;60(1):86-93.
6. Van Tomme SR, van Nostrum CF, Dijkstra M, De Smedt SC, Hennink WE. Effect of particle size and charge on the network properties of microsphere-based hydrogels. Eur J Pharm Biopharm 2008;70(2):522-530.
7. Torres-Lugo M, Peppas NA. Preparation and Characterization of P(MAA-g-EG) Nanospheres for Protein Delivery Applications. Journal of Nanoparticle Research 2002;4(1):73-81.
8. Benoit DS, Anseth KS. Heparin functionalized PEG gels that modulate protein adsorption for hMSC adhesion and differentiation. Acta Biomater 2005;1(4):461- 470.
9. Cushing MC, Liao JT, Jaeggli MP, Anseth KS. Material-based regulation of the myofibroblast phenotype. Biomaterials 2007;28(23):3378-3387.
10. Smeds KA, Pfister-Serres A, Miki D, Dastgheib K, Inoue M, Hatchell DL, Grinstaff MW. Photocrosslinkable polysaccharides for in situ hydrogel formation. J Biomed Mater Res 2001;54(1):115-121.
11. Castellot JJ, Wong K, Herman B, Hoover RL, Albertini DF, Wright TC, Caleb BL, Karnovsky MJ. Binding and Internalization of Heparin by Vascular Smooth-Muscle Cells. J Cell Physiol 1985;124(1):13-20.
233 12. Patel RC, Handy I, Patel CV. Contribution of double-stranded RNA-activated protein kinase toward antiproliferative actions of heparin on vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 2002;22(9):1439-1444.
13. Bjorkerud S. Effects of transforming growth factor-beta 1 on human arterial smooth muscle cells in vitro. Arterioscler Thromb 1991;11(4):892-902.
14. Deaton RA, Su C, Valencia TG, Grant SR. Transforming growth factor-beta 1- induced expression of smooth muscle marker genes involves activation of PKN and p38 MAPK. Journal of Biological Chemistry 2005;280(35):31172-31181.
15. Hautmann MB, Madsen CS, Owens GK. A transforming growth factor beta (TGFbeta) control element drives TGFbeta-induced stimulation of smooth muscle alpha-actin gene expression in concert with two CArG elements. J Biol Chem 1997;272(16):10948-10956.
16. Kawai-Kowase K, Sato H, Oyama Y, Kanai H, Sato M, Doi H, Kurabayashi M. Basic fibroblast growth factor antagonizes transforming growth factor-beta1-induced smooth muscle gene expression through extracellular signal-regulated kinase 1/2 signaling pathway activation. Arterioscler Thromb Vasc Biol 2004;24(8):1384-1390.
17. Papetti M, Shujath J, Riley KN, Herman IM. FGF-2 antagonizes the TGF-beta1- mediated induction of pericyte alpha-smooth muscle actin expression: a role for myf- 5 and Smad-mediated signaling pathways. Invest Ophthalmol Vis Sci 2003;44(11):4994-5005.
18. Sawhney AS, Pathak CP, Hubbell JA. Bioerodible Hydrogels Based on Photopolymerized Poly(Ethylene Glycol)-Co-Poly(Alpha-Hydroxy Acid) Diacrylate Macromers. Macromolecules 1993;26(4):581-587.
19. Mahoney MJ, Anseth KS. Three-dimensional growth and function of neural tissue in degradable polyethylene glycol hydrogels. Biomaterials 2006;27(10):2265-2274.
20. Gobin AS, West JL. Cell migration through defined, synthetic ECM analogs. Faseb J 2002;16(7):751-753.
21. Mann BK, Gobin AS, Tsai AT, Schmedlen RH, West JL. Smooth muscle cell growth in photopolymerized hydrogels with cell adhesive and proteolytically degradable domains: synthetic ECM analogs for tissue engineering. Biomaterials 2001;22(22):3045-3051.
22. West JL, Hubbell JA. Polymeric biomaterials with degradation sites for proteases involved in cell migration. Macromolecules 1999;32(1):241-244.
23. Newby AC. Matrix metalloproteinases regulate migration, proliferation, and death of vascular smooth muscle cells by degrading matrix and non-matrix substrates. Cardiovascular Research 2006;69(3):614-624.
234 24. Parks WC, Mecham RP. Matrix metalloproteinases. San Diego: Academic Press, 1998.
25. Hahn MS, McHale MK, Wang E, Schmedlen RH, West JL. Physiologic pulsatile flow bioreactor conditioning of poly(ethylene glycol)-based tissue engineered vascular grafts. Ann Biomed Eng 2007;35(2):190-200.
26. Adelow C, Segura T, Hubbell JA, Frey P. The effect of enzymatically degradable poly(ethylene glycol) hydrogels on smooth muscle cell phenotype. Biomaterials 2008;29(3):314-326.
27. Lutolf MP, Lauer-Fields JL, Schmoekel HG, Metters AT, Weber FE, Fields GB, Hubbell JA. Synthetic matrix metalloproteinase-sensitive hydrogels for the conduction of tissue regeneration: engineering cell-invasion characteristics. Proc Natl Acad Sci U S A 2003;100(9):5413-5418.
28. Beamish JA, Zhu J, Kottke-Marchant K, Marchant RE. The effects of monoacrylated poly(ethylene glycol) on the properties of poly(ethylene glycol) diacrylate hydrogels used for tissue engineering. J Biomed Mater Res 2009;in press.
29. Lutolf MP, Hubbell JA. Synthesis and physicochemical characterization of end- linked poly(ethylene glycol)-co-peptide hydrogels formed by Michael-type addition. Biomacromolecules 2003;4(3):713-722.
30. Patterson J, Hubbell JA. Molecularly Engineered PEG Hydrogels with Enhanced Proteolysis. Society for Biomaterials Annual Meeting; 2009 April 22-25; San Antonio, TX; 2009. p. ABSTRACT.
31. Salacinski HJ, Goldner S, Giudiceandrea A, Hamilton G, Seifalian AM, Edwards A, Carson RJ. The mechanical behavior of vascular grafts: a review. J Biomater Appl 2001;15(3):241-278.
32. Walden R, L'Italien GJ, Megerman J, Abbott WM. Matched elastic properties and successful arterial grafting. Arch Surg 1980;115(10):1166-1169.
33. Haruguchi H, Teraoka S. Intimal hyperplasia and hemodynamic factors in arterial bypass and arteriovenous grafts: a review. J Artif Organs 2003;6(4):227-235.
34. Hedin U, Bottger BA, Forsberg E, Johansson S, Thyberg J. Diverse effects of fibronectin and laminin on phenotypic properties of cultured arterial smooth muscle cells. J Cell Biol 1988;107(1):307-319.
35. Hayashi K, Saga H, Chimori Y, Kimura K, Yamanaka Y, Sobue K. Differentiated phenotype of smooth muscle cells depends on signaling pathways through insulin- like growth factors and phosphatidylinositol 3-kinase. J Biol Chem 1998;273(44):28860-28867.
235 36. Hayward IP, Bridle KR, Campbell GR, Underwood PA, Campbell JH. Effect of extracellular matrix proteins on vascular smooth muscle cell phenotype. Cell Biol Int 1995;19(10):839-846.
37. Thyberg J, Hultgardh-Nilsson A. Fibronectin and the basement membrane components laminin and collagen type IV influence the phenotypic properties of subcultured rat aortic smooth muscle cells differently. Cell Tissue Res 1994;276(2):263-271.
38. Yamamoto M, Yamamoto K, Noumura T. Type I collagen promotes modulation of cultured rabbit arterial smooth muscle cells from a contractile to a synthetic phenotype. Exp Cell Res 1993;204(1):121-129.
39. Hayashi K, Shibata K, Morita T, Iwasaki K, Watanabe M, Sobue K. Insulin receptor substrate-1/SHP-2 interaction, a phenotype-dependent switching machinery of insulin-like growth factor-I signaling in vascular smooth muscle cells. J Biol Chem 2004;279(39):40807-40818.
40. Moiseeva EP. Adhesion receptors of vascular smooth muscle cells and their functions. Cardiovasc Res 2001;52(3):372-386.
41. Raines EW, Koyama H, Carragher NO. The extracellular matrix dynamically regulates smooth muscle cell responsiveness to PDGF. Ann N Y Acad Sci 2000;902:39-51; discussion 51-32.
42. Hedin UL, Daum G, Clowes AW. Disruption of integrin alpha 5 beta 1 signaling does not impair PDGF-BB-mediated stimulation of the extracellular signal-regulated kinase pathway in smooth muscle cells. J Cell Physiol 1997;172(1):109-116.
43. Peyton SR, Ghajar CM, Khatiwala CB, Putnam AJ. The emergence of ECM mechanics and cytoskeletal tension as important regulators of cell function. Cell Biochem Biophys 2007;47(2):300-320.
44. Mack CP, Somlyo AV, Hautmann M, Somlyo AP, Owens GK. Smooth muscle differentiation marker gene expression is regulated by RhoA-mediated actin polymerization. Journal of Biological Chemistry 2001;276(1):341-347.
45. Goffin JM, Pittet P, Csucs G, Lussi JW, Meister JJ, Hinz B. Focal adhesion size controls tension-dependent recruitment of alpha-smooth muscle actin to stress fibers. J Cell Biol 2006;172(2):259-268.
46. Wang Z, Wang DZ, Pipes GC, Olson EN. Myocardin is a master regulator of smooth muscle gene expression. Proc Natl Acad Sci U S A 2003;100(12):7129-7134.
47. Miano JM. Serum response factor: toggling between disparate programs of gene expression. J Mol Cell Cardiol 2003;35(6):577-593.
236 48. Yoshida T, Kawai-Kowase K, Owens GK. Forced expression of myocardin is not sufficient for induction of smooth muscle differentiation in multipotential embryonic cells. Arterioscler Thromb Vasc Biol 2004;24(9):1596-1601.
49. Mack CP, Hinson JS. Regulation of smooth muscle differentiation by the myocardin family of serum response factor co-factors. J Thromb Haemost 2005;3(9):1976-1984.
50. Gan Q, Yoshida T, Li J, Owens GK. Smooth muscle cells and myofibroblasts use distinct transcriptional mechanisms for smooth muscle alpha-actin expression. Circ Res 2007;101(9):883-892.
237 Bibliography
Abbott WM, Megerman J, Hasson JE, L'Italien G, Warnock DF. Effect of compliance mismatch on vascular graft patency. J Vasc Surg 1987;5(2):376-382.
Abbott WM, Callow A, Moore W, Rutherford R, Veith F, Weinberg S. Evaluation and performance standards for arterial prostheses. J Vasc Surg 1993;17(4):746-756.
Adam PJ, Regan CP, Hautmann MB, Owens GK. Positive- and negative-acting Kruppel- like transcription factors bind a transforming growth factor beta control element required for expression of the smooth muscle cell differentiation marker SM22alpha in vivo. J Biol Chem 2000;275(48):37798-37806.
Adelow C, Segura T, Hubbell JA, Frey P. The effect of enzymatically degradable poly(ethylene glycol) hydrogels on smooth muscle cell phenotype. Biomaterials 2008;29(3):314-326.
Alberts B. Molecular biology of the cell. 4th ed. New York: Garland Science, 2002.
Almany L, Seliktar D. Biosynthetic hydrogel scaffolds made from fibrinogen and polyethylene glycol for 3D cell cultures. Biomaterials 2005;26(15):2467-2477.
Anseth KS, Kline LM, Walker TA, Anderson KJ, Bowman CN. Reaction-Kinetics and Volume Relaxation during Polymerizations of Multiethylene Glycol Dimethacrylates. MACROMOLECULES 1995;28(7):2491-2499.
Arciniegas E, Sutton AB, Allen TD, Schor AM. Transforming growth factor beta 1 promotes the differentiation of endothelial cells into smooth muscle-like cells in vitro. J Cell Sci 1992;103 ( Pt 2):521-529.
Assinder SJ, Stanton JA, Prasad PD. Transgelin: an actin-binding protein and tumour suppressor. Int J Biochem Cell Biol 2009;41(3):482-486.
Babij P, Kawamoto S, White S, Adelstein RS, Periasamy M. Differential expression of SM1 and SM2 myosin isoforms in cultured vascular smooth muscle. Am J Physiol 1992;262(3 Pt 1):C607-613.
Babu GJ, Warshaw DM, Periasamy M. Smooth muscle myosin heavy chain isoforms and their role in muscle physiology. Microsc Res Tech 2000;50(6):532-540.
Babu GJ, Pyne GJ, Zhou Y, Okwuchukuasanya C, Brayden JE, Osol G, Paul RJ, Low RB, Periasamy M. Isoform switching from SM-B to SM-A myosin results in decreased contractility and altered expression of thin filament regulatory proteins. Am J Physiol Cell Physiol 2004;287(3):C723-729.
Baker SC, Southgate J. Towards control of smooth muscle cell differentiation in synthetic 3D scaffolds. Biomaterials 2008;29(23):3357-3366.
238 Ballyk PD, Walsh C, Butany J, Ojha M. Compliance mismatch may promote graft-artery intimal hyperplasia by altering suture-line stresses. J Biomech 1998;31(3):229-237.
Bassiouny HS, White S, Glagov S, Choi E, Giddens DP, Zarins CK. Anastomotic intimal hyperplasia: mechanical injury or flow induced. J Vasc Surg 1992;15(4):708-716; discussion 716-707.
Beamish JA, Fu A, Choi A, Haq N, Kottke-Marchant K, Marchant RE. The influence of RGD-bearing hydrogels on the re-expression of contractile vascular smooth muscle cell phenotype. Biomaterials 2009;in press.
Beamish JA, Zhu J, Kottke-Marchant K, Marchant RE. The effects of monoacrylated poly(ethylene glycol) on the properties of poly(ethylene glycol) diacrylate hydrogels used for tissue engineering. J Biomed Mater Res 2009;in press.
Benoit DS, Anseth KS. Heparin functionalized PEG gels that modulate protein adsorption for hMSC adhesion and differentiation. Acta Biomater 2005;1(4):461-470.
Benoit DS, Anseth KS. The effect on osteoblast function of colocalized RGD and PHSRN epitopes on PEG surfaces. Biomaterials 2005;26(25):5209-5220.
Benoit DS, Nuttelman CR, Collins SD, Anseth KS. Synthesis and characterization of a fluvastatin-releasing hydrogel delivery system to modulate hMSC differentiation and function for bone regeneration. Biomaterials 2006;27(36):6102-6110.
Bingley JA, Hayward IP, Campbell JH, Campbell GR. Arterial heparan sulfate proteoglycans inhibit vascular smooth muscle cell proliferation and phenotype change in vitro and neointimal formation in vivo. J Vasc Surg 1998;28(2):308-318.
Birukov KG, Shirinsky VP, Stepanova OV, Tkachuk VA, Hahn AW, Resink TJ, Smirnov VN. Stretch affects phenotype and proliferation of vascular smooth muscle cells. Mol Cell Biochem 1995;144(2):131-139.
Bjorkerud S. Effects of transforming growth factor-beta 1 on human arterial smooth muscle cells in vitro. Arterioscler Thromb 1991;11(4):892-902.
Boerth NJ, Dey NB, Cornwell TL, Lincoln TM. Cyclic GMP-dependent protein kinase regulates vascular smooth muscle cell phenotype. J Vasc Res 1997;34(4):245-259.
Bono F, Rigon P, Lamarche I, Savi P, Salel V, Herbert JM. Heparin inhibits the binding of basic fibroblast growth factor to cultured human aortic smooth-muscle cells. Biochem J 1997;326 ( Pt 3):661-668.
Boron WF, Boulpaep EL. Medical physiology : a cellular and molecular approach. Updated ed. Philadelphia, Pa.: Elsevier Saunders, 2005.
239 Borschel GH, Huang YC, Calve S, Arruda EM, Lynch JB, Dow DE, Kuzon WM, Dennis RG, Brown DL. Tissue engineering of recellularized small-diameter vascular grafts. Tissue Engineering 2005;11(5-6):778-786.
Boxer LM, Prywes R, Roeder RG, Kedes L. The sarcomeric actin CArG-binding factor is indistinguishable from the c-fos serum response factor. Mol Cell Biol 1989;9(2):515- 522.
Bray JC, Merrill EW. Poly(vinyl alcohol) hydrogels. Formation by electron beam irradiation of aqueous solutions and subsequent crystallization. J Appl Polym Sci 1973;17:3779-3794.
Brewster LP, Bufallino D, Ucuzian A, Greisler HP. Growing a living blood vessel: insights for the second hundred years. Biomaterials 2007;28(34):5028-5032.
Brophy CM, Woodrum DA, Pollock J, Dickinson M, Komalavilas P, Cornwell TL, Lincoln TM. cGMP-dependent protein kinase expression restores contractile function in cultured vascular smooth muscle cells. J Vasc Res 2002;39(2):95-103.
Bryant SJ, Nuttelman CR, Anseth KS. The effects of crosslinking density on cartilage formation in photocrosslinkable hydrogels. Biomed Sci Instrum 1999;35:309-314.
Bryant SJ, Anseth KS. Hydrogel properties influence ECM production by chondrocytes photoencapsulated in poly(ethylene glycol) hydrogels. J Biomed Mater Res 2002;59(1):63-72.
Bryant SJ, Bender RJ, Durand KL, Anseth KS. Encapsulating chondrocytes in degrading PEG hydrogels with high modulus: engineering gel structural changes to facilitate cartilaginous tissue production. Biotechnol Bioeng 2004;86(7):747-755.
Bulick AS, Munoz-Pinto DJ, Qu X, Mani M, Cristancho D, Urban M, Hahn MS. Impact of endothelial cells and mechanical conditioning on smooth muscle cell extracellular matrix production and differentiation. Tissue Eng Part A 2009;15(4):815-825.
Burdick JA, Anseth KS. Photoencapsulation of osteoblasts in injectable RGD-modified PEG hydrogels for bone tissue engineering. Biomaterials 2002;23(22):4315-4323.
Burdick JA, Lovestead TM, Anseth KS. Kinetic chain lengths in highly cross-linked networks formed by the photoinitiated polymerization of divinyl monomers: A gel permeation chromatography investigation. Biomacromolecules 2003;4(1):149-156.
Burgstaller G, Kranewitter WJ, Gimona M. The molecular basis for the autoregulation of calponin by isoform-specific C-terminal tail sequences. J Cell Sci 2002;115(Pt 10):2021-2029.
Buxton AN, Zhu J, Marchant R, West JL, Yoo JU, Johnstone B. Design and characterization of poly(ethylene glycol) photopolymerizable semi-interpenetrating
240 networks for chondrogenesis of human mesenchymal stem cells. Tissue Eng 2007;13(10):2549-2560.
Cameron A, Davis KB, Green G, Schaff HV. Coronary Bypass Surgery with Internal- Thoracic-Artery Grafts -- Effects on Survival over a 15-Year Period. N Engl J Med 1996;334(4):216-220.
Campbell JH, Campbell GR. Vascular smooth muscle in culture. Boca Raton, Fla.: CRC Press, 1987.
Canal T, Peppas NA. Correlation between Mesh Size and Equilibrium Degree of Swelling of Polymeric Networks. J Biomed Mater Res 1989;23(10):1183-1193.
Cappadona C, Redmond EM, Theodorakis NG, McKillop IH, Hendrickson R, Chhabra A, Sitzmann JV, Cahill PA. Phenotype dictates the growth response of vascular smooth muscle cells to pulse pressure in vitro. Exp Cell Res 1999;250(1):174-186.
Castellot JJ, Wong K, Herman B, Hoover RL, Albertini DF, Wright TC, Caleb BL, Karnovsky MJ. Binding and Internalization of Heparin by Vascular Smooth-Muscle Cells. J Cell Physiol 1985;124(1):13-20.
Castellot JJ, Choay J, Lormeau JC, Petitou M, Sache E, Karnovsky MJ. Structural Determinants of the Capacity of Heparin to Inhibit the Proliferation of Vascular Smooth-Muscle Cells .2. Evidence for a Pentasaccharide Sequence That Contains a 3- O-Sulfate Group. J Cell Biol 1986;102(5):1979-1984.
Cavigelli M, Dolfi F, Claret FX, Karin M. Induction of c-fos expression through JNK- mediated TCF/Elk-1 phosphorylation. Embo J 1995;14(23):5957-5964.
Chamley-Campbell J, Campbell GR, Ross R. The smooth muscle cell in culture. Physiol Rev 1979;59(1):1-61.
Chamley-Campbell JH, Campbell GR, Ross R. Phenotype-dependent response of cultured aortic smooth muscle to serum mitogens. J Cell Biol 1981;89(2):379-383.
Chan-Park MB, Shen JY, Cao Y, Xiong Y, Liu Y, Rayatpisheh S, Kang GC, Greisler HP. Biomimetic control of vascular smooth muscle cell morphology and phenotype for functional tissue-engineered small-diameter blood vessels. J Biomed Mater Res A 2009;88(4):1104-1121.
Chapman GB, Durante W, Hellums JD, Schafer AI. Physiological cyclic stretch causes cell cycle arrest in cultured vascular smooth muscle cells. Am J Physiol Heart Circ Physiol 2000;278(3):H748-754.
Chard RB, Johnson DC, Nunn GR, Cartmill TB. Aorta-coronary bypass grafting with polytetrafluoroethylene conduits. Early and late outcome in eight patients. J Thorac Cardiovasc Surg 1987;94(1):132-134.
241 Chen Q, Li W, Quan Z, Sumpio BE. Modulation of vascular smooth muscle cell alignment by cyclic strain is dependent on reactive oxygen species and P38 mitogen- activated protein kinase. J Vasc Surg 2003;37(3):660-668.
Chi M, Zhou Y, Vedamoorthyrao S, Babu GJ, Periasamy M. Ablation of smooth muscle myosin heavy chain SM2 increases smooth muscle contraction and results in postnatal death in mice. Proc Natl Acad Sci U S A 2008;105(47):18614-18618.
Cho SW, Lim SH, Kim IK, Hong YS, Kim SS, Yoo KJ, Park HY, Jang Y, Chang BC, Choi CY, Hwang KC, Kim BS. Small-diameter blood vessels engineered with bone marrow-derived cells. Annals of Surgery 2005;241(3):506-515.
Cho SW, Kim IK, Kang JM, Song KW, Kim HS, Park CH, Yoo KJ, Kim BS. Evidence for in vivo growth potential and vascular remodeling of tissue-engineered artery. Tissue Eng Part A 2009;15(4):901-912.
Christen T, Bochaton-Piallat ML, Neuville P, Rensen S, Redard M, van Eys G, Gabbiani G. Cultured porcine coronary artery smooth muscle cells. A new model with advanced differentiation. Circ Res 1999;85(1):99-107.
Christen T, Verin V, Bochaton-Piallat M, Popowski Y, Ramaekers F, Debruyne P, Camenzind E, van Eys G, Gabbiani G. Mechanisms of neointima formation and remodeling in the porcine coronary artery. Circulation 2001;103(6):882-888.
Cirillo P, Golino P, Calabro P, Cali G, Ragni M, De Rosa S, Cimmino G, Pacileo M, De Palma R, Forte L, Gargiulo A, Corigliano FG, Angri V, Spagnuolo R, Nitsch L, Chiariello M. C-reactive protein induces tissue factor expression and promotes smooth muscle and endothelial cell proliferation. Cardiovasc Res 2005;68(1):47-55.
Clowes AW, Kirkman TR, Reidy MA. Mechanisms of arterial graft healing. Rapid transmural capillary ingrowth provides a source of intimal endothelium and smooth muscle in porous PTFE prostheses. Am J Pathol 1986;123(2):220-230.
Conklin BS, Richter ER, Kreutziger KL, Zhong DS, Chen C. Development and evaluation of a novel decellularized vascular xenograft. Medical Engineering & Physics 2002;24(3):173-183.
Cruise GM, Scharp DS, Hubbell JA. Characterization of permeability and network structure of interfacially photopolymerized poly(ethylene glycol) diacrylate hydrogels. Biomaterials 1998;19(14):1287-1294.
Cushing MC, Liao JT, Jaeggli MP, Anseth KS. Material-based regulation of the myofibroblast phenotype. Biomaterials 2007;28(23):3378-3387.
Dandre F, Owens GK. Platelet-derived growth factor-BB and Ets-1 transcription factor negatively regulate transcription of multiple smooth muscle cell differentiation marker genes. Am J Physiol Heart Circ Physiol 2004;286(6):H2042-2051.
242 Daniel J, Abe K, McFetridge PS. Development of the human umbilical vein scaffold for cardiovascular tissue engineering applications. Asaio Journal 2005;51(3):252-261.
Darouiche RO. Current concepts - Treatment of infections associated with surgical implants. N Engl J Med 2004;350(14):1422-1429.
Darouiche RO, Mansouri MD. In vitro activity and in vivo efficacy of antimicrobial- coated vascular grafts. Annals of Vascular Surgery 2004;18(4):497-501.
Dartsch PC, Hammerle H, Betz E. Orientation of cultured arterial smooth muscle cells growing on cyclically stretched substrates. Acta Anat (Basel) 1986;125(2):108-113.
Daum G, Hedin U, Wang Y, Wang T, Clowes AW. Diverse effects of heparin on mitogen-activated protein kinase-dependent signal transduction in vascular smooth muscle cells. Circ Res 1997;81(1):17-23.
Davies PF, Olesen SP, Clapham DE, Morrel EM, Schoen FJ. Endothelial Communication - State of the Art Lecture. Hypertension 1988;11(6):563-572.
Deaton RA, Su C, Valencia TG, Grant SR. Transforming growth factor-beta 1-induced expression of smooth muscle marker genes involves activation of PKN and p38 MAPK. J Biol Chem 2005;280(35):31172-31181.
DeFail AJ, Chu CR, Izzo N, Marra KG. Controlled release of bioactive TGF-beta(1) from microspheres embedded within biodegradable hydrogels. Biomaterials 2006;27(8):1579-1585.
Deissler H, Deissler H, Lang GK, Lang GE. TGFbeta induces transdifferentiation of iBREC to alphaSMA-expressing cells. Int J Mol Med 2006;18(4):577-582.
DeLong SA, Moon JJ, West JL. Covalently immobilized gradients of bFGF on hydrogel scaffolds for directed cell migration. Biomaterials 2005;26(16):3227-3234.
Deruiter MC, Rensen SS, Coolen GP, Hierck BP, Bergwerff M, Debie WM, Gittenberger-De Groot AC, Van Eys GJ. Smoothelin expression during chicken embryogenesis: detection of an embryonic isoform. Dev Dyn 2001;221(4):460-463.
Desmouliere A, Geinoz A, Gabbiani F, Gabbiani G. Transforming growth factor-beta 1 induces alpha-smooth muscle actin expression in granulation tissue myofibroblasts and in quiescent and growing cultured fibroblasts. J Cell Biol 1993;122(1):103-111.
Dobrin PB, Littooy FN, Endean ED. Mechanical factors predisposing to intimal hyperplasia and medial thickening in autogenous vein grafts. Surgery 1989;105(3):393-400.
Doi H, Iso T, Yamazaki M, Akiyama H, Kanai H, Sato H, Kawai-Kowase K, Tanaka T, Maeno T, Okamoto E, Arai M, Kedes L, Kurabayashi M. HERP1 inhibits myocardin-
243 induced vascular smooth muscle cell differentiation by interfering with SRF binding to CArG box. Arterioscler Thromb Vasc Biol 2005;25(11):2328-2334.
Donaldson MC, Mannick JA, Whittemore AD. Causes of primary graft failure after in situ saphenous vein bypass grafting. J Vasc Surg 1992;15(1):113-118; discussion 118-120.
Duband JL, Gimona M, Scatena M, Sartore S, Small JV. Calponin and SM 22 as differentiation markers of smooth muscle: spatiotemporal distribution during avian embryonic development. Differentiation 1993;55(1):1-11.
Faggin E, Puato M, Zardo L, Franch R, Millino C, Sarinella F, Pauletto P, Sartore S, Chiavegato A. Smooth muscle-specific SM22 protein is expressed in the adventitial cells of balloon-injured rabbit carotid artery. Arterioscler Thromb Vasc Biol 1999;19(6):1393-1404.
Fannon M, Forsten KE, Nugent MA. Potentiation and inhibition of bFGF binding by heparin: a model for regulation of cellular response. Biochemistry 2000;39(6):1434- 1445.
Faries PL, Logerfo FW, Arora S, Hook S, Pulling MC, Akbari CM, Campbell DR, Pomposelli FB, Jr. A comparative study of alternative conduits for lower extremity revascularization: all-autogenous conduit versus prosthetic grafts. J Vasc Surg 2000;32(6):1080-1090.
Farndale RW, Buttle DJ, Barrett AJ. Improved quantitation and discrimination of sulphated glycosaminoglycans by use of dimethylmethylene blue. Biochimica et Biophysica Acta (BBA) - General Subjects 1986;883(2):173-177.
Fasciano S, Patel RC, Handy I, Patel CV. Regulation of vascular smooth muscle proliferation by heparin: inhibition of cyclin-dependent kinase 2 activity by p27(kip1). J Biol Chem 2005;280(16):15682-15689.
Fillinger MF, Sampson LN, Cronenwett JL, Powell RJ, Wagner RJ. Coculture of endothelial cells and smooth muscle cells in bilayer and conditioned media models. J Surg Res 1997;67(2):169-178.
Fittkau MH, Zilla P, Bezuidenhout D, Lutolf MP, Human P, Hubbell JA, Davies N. The selective modulation of endothelial cell mobility on RGD peptide containing surfaces by YIGSR peptides. Biomaterials 2005;26(2):167-174.
Flory PJ. Principles of polymer chemistry. Ithaca,: Cornell University Press, 1953.
Frid MG, Printesva OY, Chiavegato A, Faggin E, Scatena M, Koteliansky VE, Pauletto P, Glukhova MA, Sartore S. Myosin heavy-chain isoform composition and distribution in developing and adult human aortic smooth muscle. J Vasc Res 1993;30(5):279-292.
244 Fu Y, Liu HW, Forsythe SM, Kogut P, McConville JF, Halayko AJ, Camoretti-Mercado B, Solway J. Mutagenesis analysis of human SM22: characterization of actin binding. J Appl Physiol 2000;89(5):1985-1990.
Gan Q, Yoshida T, Li J, Owens GK. Smooth muscle cells and myofibroblasts use distinct transcriptional mechanisms for smooth muscle alpha-actin expression. Circ Res 2007;101(9):883-892.
Garg HG, Joseph PAM, Yoshida K, Thompson BT, Hales CA. Antiproliferative role of 3-O-sulfate glucosamine in heparin on cultured pulmonary artery smooth muscle cells. Biochem Biophys Res Commun 1996;224(2):468-473.
Garg HG, Joseph PAM, Thompson BT, Hales CA, Toida T, Imanari T, Capila I, Linhardt RJ. Effect of fully sulfated glycosaminoglycans on pulmonary artery smooth muscle cell proliferation. Arch Biochem Biophys 1999;371(2):228-233.
Garg HG, Thompson BT, Hales CA. Structural determinants of antiproliferative activity of heparin on pulmonary artery smooth muscle cells. Am J Physiol Lung Cell Mol Physiol 2000;279(5):L779-789.
Garg HG, Cindhuchao N, Quinn DA, Hales CA, Thanawiroon C, Capila I, Linhardt RJ. Heparin oligosaccharide sequence and size essential for inhibition of pulmonary artery smooth muscle cell proliferation. Carbohydr Res 2002;337(21-23):2359-2364.
Garg HG, Yu L, Hales CA, Toida T, Islam T, Linhardt RJ. Sulfation patterns in heparin and heparan sulfate: effects on the proliferation of bovine pulmonary artery smooth muscle cells. Biochim Biophys Acta 2003;1639(3):225-231.
Garg HG, Mrabat H, Yu L, Freeman C, Li B, Zhang F, Linhardt RJ, Hales CA. Significance of the 2-O-sulfo group of L-iduronic acid residues in heparin on the growth inhibition of bovine pulmonary artery smooth muscle cells. Carbohydr Res 2008;343(14):2406-2410.
Gartner LP, Hiatt JL. Color atlas of histology. 4th ed. Philadelphia: Lippincott Williams & Wilkins, 2006.
Geisterfer AAT, Peach MJ, Owens GK. Angiotensin-Ii Induces Hypertrophy, Not Hyperplasia, of Cultured Rat Aortic Smooth-Muscle Cells. CircRes 1988;62(4):749- 756.
Girton TS, Oegema TR, Tranquillo RT. Exploiting glycation to stiffen and strengthen tissue equivalents for tissue engineering. J Biomed Mater Res 1999;46(1):87-92.
Gobin AS, West JL. Cell migration through defined, synthetic ECM analogs. Faseb J 2002;16(7):751-753.
Gobin AS, West JL. Val-ala-pro-gly, an elastin-derived non-integrin ligand: smooth muscle cell adhesion and specificity. J Biomed Mater Res A 2003;67(1):255-259.
245 Gobin AS, West JL. Effects of epidermal growth factor on fibroblast migration through biomimetic hydrogels. Biotechnol Prog 2003;19(6):1781-1785.
Goffin JM, Pittet P, Csucs G, Lussi JW, Meister JJ, Hinz B. Focal adhesion size controls tension-dependent recruitment of alpha-smooth muscle actin to stress fibers. J Cell Biol 2006;172(2):259-268.
Graf J, Ogle RC, Robey FA, Sasaki M, Martin GR, Yamada Y, Kleinman HK. A pentapeptide from the laminin B1 chain mediates cell adhesion and binds the 67,000 laminin receptor. Biochemistry 1987;26(22):6896-6900.
Grainger DJ, Metcalfe JC, Grace AA, Mosedale DE. Transforming growth factor-beta dynamically regulates vascular smooth muscle differentiation in vivo. J Cell Sci 1998;111 ( Pt 19):2977-2988.
Grassl ED, Oegema TR, Tranquillo RT. A fibrin-based arterial media equivalent. J Biomed Mater Res Part A 2003;66A(3):550-561.
Greenwald SE, Berry CL. Improving vascular grafts: the importance of mechanical and haemodynamic properties. J Pathol 2000;190(3):292-299.
Gunn JW, Turner SD, Mann BK. Adhesive and mechanical properties of hydrogels influence neurite extension. J Biomed Mater Res A 2005;72(1):91-97.
Guyton JR, Rosenberg RD, Clowes AW, Karnovsky MJ. Inhibition of Rat Arterial Smooth-Muscle Cell-Proliferation by Heparin - Invivo Studies with Anti-Coagulant and Non-Anticoagulant Heparin. CircRes 1980;46(5):625-634.
Hahn MS, Taite LJ, Moon JJ, Rowland MC, Ruffino KA, West JL. Photolithographic patterning of polyethylene glycol hydrogels. Biomaterials 2006;27(12):2519-2524.
Hahn MS, McHale MK, Wang E, Schmedlen RH, West JL. Physiologic pulsatile flow bioreactor conditioning of poly(ethylene glycol)-based tissue engineered vascular grafts. Ann Biomed Eng 2007;35(2):190-200.
Halstenberg S, Panitch A, Rizzi S, Hall H, Hubbell JA. Biologically engineered protein- graft-poly(ethylene glycol) hydrogels: A cell adhesive and plasm in-degradable biosynthetic material for tissue repair. Biomacromolecules 2002;3(4):710-723.
Hancock WW, Adams DH, Wyner LR, Sayegh MH, Karnovsky MJ. CD4+ mononuclear cells induce cytokine expression, vascular smooth muscle cell proliferation, and arterial occlusion after endothelial injury. Am J Pathol 1994;145(5):1008-1014.
Hannan EL, Wu C, Walford G, Culliford AT, Gold JP, Smith CR, Higgins RS, Carlson RE, Jones RH. Drug-eluting stents vs. coronary-artery bypass grafting in multivessel coronary disease. N Engl J Med 2008;358(4):331-341.
246 Haruguchi H, Teraoka S. Intimal hyperplasia and hemodynamic factors in arterial bypass and arteriovenous grafts: a review. J Artif Organs 2003;6(4):227-235.
Hashi CK, Zhu YQ, Yang GY, Young WL, Hsiao BS, Wang K, Chu B, Li S. Antithrombogenic property of bone marrow mesenchymal stem cells in nanofibrous vascular grafts. Proceedings of the National Academy of Sciences of the United States of America 2007;104(29):11915-11920.
Hashimoto T, Kihara M, Sato K, Imai N, Tanaka Y, Sakai M, Tamura K, Hirawa N, Toya Y, Kitamura H, Umemura S. Heparin recovers AT1 receptor and its intracellular signal transduction in cultured vascular smooth muscle cells. FEBS Lett 2005;579(1):281-284.
Hautmann MB, Madsen CS, Owens GK. A transforming growth factor beta (TGFbeta) control element drives TGFbeta-induced stimulation of smooth muscle alpha-actin gene expression in concert with two CArG elements. J Biol Chem 1997;272(16):10948-10956.
Hayashi K, Saga H, Chimori Y, Kimura K, Yamanaka Y, Sobue K. Differentiated phenotype of smooth muscle cells depends on signaling pathways through insulin-like growth factors and phosphatidylinositol 3-kinase. J Biol Chem 1998;273(44):28860- 28867.
Hayashi K, Takahashi M, Kimura K, Nishida W, Saga H, Sobue K. Changes in the balance of phosphoinositide 3-kinase/protein kinase B (Akt) and the mitogen- activated protein kinases (ERK/p38MAPK) determine a phenotype of visceral and vascular smooth muscle cells. J Cell Biol 1999;145(4):727-740.
Hayashi K, Shibata K, Morita T, Iwasaki K, Watanabe M, Sobue K. Insulin receptor substrate-1/SHP-2 interaction, a phenotype-dependent switching machinery of insulin-like growth factor-I signaling in vascular smooth muscle cells. J Biol Chem 2004;279(39):40807-40818.
Hayward IP, Bridle KR, Campbell GR, Underwood PA, Campbell JH. Effect of extracellular matrix proteins on vascular smooth muscle cell phenotype. Cell Biol Int 1995;19(10):839-846.
Hedin U, Thyberg J. Plasma fibronectin promotes modulation of arterial smooth-muscle cells from contractile to synthetic phenotype. Differentiation 1987;33(3):239-246.
Hedin U, Bottger BA, Forsberg E, Johansson S, Thyberg J. Diverse effects of fibronectin and laminin on phenotypic properties of cultured arterial smooth muscle cells. J Cell Biol 1988;107(1):307-319.
Hedin U, Bottger BA, Luthman J, Johansson S, Thyberg J. A substrate of the cell- attachment sequence of fibronectin (Arg-Gly-Asp-Ser) is sufficient to promote transition of arterial smooth muscle cells from a contractile to a synthetic phenotype. Dev Biol 1989;133(2):489-501.
247 Hedin U, Thyberg J, Roy J, Dumitrescu A, Tran PK. Role of tyrosine kinases in extracellular matrix-mediated modulation of arterial smooth muscle cell phenotype. Arterioscler Thromb Vasc Biol 1997;17(10):1977-1984.
Hedin UL, Daum G, Clowes AW. Disruption of integrin alpha 5 beta 1 signaling does not impair PDGF-BB-mediated stimulation of the extracellular signal-regulated kinase pathway in smooth muscle cells. J Cell Physiol 1997;172(1):109-116.
Hedin U, Daum G, Clowes AW. Heparin inhibits thrombin-induced mitogen-activated protein kinase signaling in arterial smooth muscle cells. J Vasc Surg 1998;27(3):512- 520.
Hehrlein FW, Schlepper M, Loskot F, Scheld HH, Walter P, Mulch J. The use of expanded polytetrafluoroethylene (PTFE) grafts for myocardial revascularization. J Cardiovasc Surg (Torino) 1984;25(6):549-553.
Hendrix JA, Wamhoff BR, McDonald OG, Sinha S, Yoshida T, Owens GK. 5' CArG degeneracy in smooth muscle alpha-actin is required for injury-induced gene suppression in vivo. J Clin Invest 2005;115(2):418-427.
Heo SK, Yun HJ, Park WH, Park SD. Emodin inhibits TNF-alpha-induced human aortic smooth-muscle cell proliferation via caspase- and mitochondrial-dependent apoptosis. J Cell Biochem 2008;105(1):70-80.
Herman IM. Actin isoforms. Curr Opin Cell Biol 1993;5(1):48-55.
Hern DL, Hubbell JA. Incorporation of adhesion peptides into nonadhesive hydrogels useful for tissue resurfacing. J Biomed Mater Res 1998;39(2):266-276.
Heydarkhan-Hagvall S, Helenius G, Johansson BR, Li JY, Mattsson E, Risberg B. Co- culture of endothelial cells and smooth muscle cells affects gene expression of angiogenic factors. J Cell Biochem 2003;89(6):1250-1259.
Higgins SP, Solan AK, Niklason LE. Effects of polyglycolic acid on porcine smooth muscle cell growth and differentiation. J Biomed Mater Res A 2003;67(1):295-302.
Hinck AP, Archer SJ, Qian SW, Roberts AB, Sporn MB, Weatherbee JA, Tsang ML, Lucas R, Zhang BL, Wenker J, Torchia DA. Transforming growth factor beta 1: three-dimensional structure in solution and comparison with the X-ray structure of transforming growth factor beta 2. Biochemistry 1996;35(26):8517-8534.
Hipper A, Isenberg G. Cyclic mechanical strain decreases the DNA synthesis of vascular smooth muscle cells. Pflugers Arch 2000;440(1):19-27.
Hirst SJ, Twort CH, Lee TH. Differential effects of extracellular matrix proteins on human airway smooth muscle cell proliferation and phenotype. Am J Respir Cell Mol Biol 2000;23(3):335-344.
248 Holland TA, Tabata Y, Mikos AG. In vitro release of transforming growth factor-beta 1 from gelatin microparticles encapsulated in biodegradable, injectable oligo(poly(ethylene glycol) fumarate) hydrogels. J Control Release 2003;91(3):299- 313.
Holland TA, Tabata Y, Mikos AG. Dual growth factor delivery from degradable oligo(poly(ethylene glycol) fumarate) hydrogel scaffolds for cartilage tissue engineering. J Control Release 2005;101(1-3):111-125.
Hong H, McCullough CM, Stegemann JP. The role of ERK signaling in protein hydrogel remodeling by vascular smooth muscle cells. Biomaterials 2007;28(26):3824-3833.
Huynh T, Abraham G, Murray J, Brockbank K, Hagen PO, Sullivan S. Remodeling of an acellular collagen graft into a physiologically responsive neovessel. Nature Biotechnology 1999;17(11):1083-1086.
Hwang NS, Varghese S, Zhang Z, Elisseeff J. Chondrogenic differentiation of human embryonic stem cell-derived cells in arginine-glycine-aspartate-modified hydrogels. Tissue Eng 2006;12(9):2695-2706.
Janakidevi K, Fisher MA, Del Vecchio PJ, Tiruppathi C, Figge J, Malik AB. Endothelin- 1 stimulates DNA synthesis and proliferation of pulmonary artery smooth muscle cells. Am J Physiol 1992;263(6 Pt 1):C1295-1301.
Je HD, Sohn UD. SM22alpha is required for agonist-induced regulation of contractility: evidence from SM22alpha knockout mice. Mol Cells 2007;23(2):175-181.
Jeong SI, Kwon JH, Lim JI, Cho SW, Jung Y, Sung WJ, Kim SH, Kim YH, Lee YM, Kim BS, Choi CY, Kim SJ. Mechano-active tissue engineering of vascular smooth muscle using pulsatile perfusion bioreactors and elastic PLCL scaffolds. Biomaterials 2005;26(12):1405-1411.
Joddar B, Ibrahim S, Ramamurthi A. Impact of delivery mode of hyaluronan oligomers on elastogenic responses of adult vascular smooth muscle cells. Biomaterials 2007;28(27):3918-3927.
Joki N, Kaname S, Hirakata M, Hori Y, Yamaguchi T, Fujita T, Katoh T, Kurokawa K. Tyrosine-kinase dependent TGF-beta and extracellular matrix expression by mechanical stretch in vascular smooth muscle cells. Hypertens Res 2000;23(2):91-99.
Kalmes A, Vesti BR, Daum G, Abraham JA, Clowes AW. Heparin blockade of thrombin-induced smooth muscle cell migration involves inhibition of epidermal growth factor (EGF) receptor transactivation by heparin-binding EGF-like growth factor. Circ Res 2000;87(2):92-98.
Karnik SK, Brooke BS, Bayes-Genis A, Sorensen L, Wythe JD, Schwartz RS, Keating MT, Li DY. A critical role for elastin signaling in vascular morphogenesis and disease. Development 2003;130(2):411-423.
249 Karnik SK, Wythe JD, Sorensen L, Brooke BS, Urness LD, Li DY. Elastin induces myofibrillogenesis via a specific domain, VGVAPG. Matrix Biol 2003;22(5):409- 425.
Kaushal S, Amiel GE, Guleserian KJ, Shapira OM, Perry T, Sutherland FW, Rabkin E, Moran AM, Schoen FJ, Atala A, Soker S, Bischoff J, Mayer JE. Functional small- diameter neovessels created using endothelial progenitor cells expanded ex vivo. Nature Medicine 2001;7(9):1035-1040.
Kawai-Kowase K, Sato H, Oyama Y, Kanai H, Sato M, Doi H, Kurabayashi M. Basic fibroblast growth factor antagonizes transforming growth factor-beta1-induced smooth muscle gene expression through extracellular signal-regulated kinase 1/2 signaling pathway activation. Arterioscler Thromb Vasc Biol 2004;24(8):1384-1390.
Kawai-Kowase K, Kumar MS, Hoofnagle MH, Yoshida T, Owens GK. PIAS1 activates the expression of smooth muscle cell differentiation marker genes by interacting with serum response factor and class I basic helix-loop-helix proteins. Mol Cell Biol 2005;25(18):8009-8023.
Kawai-Kowase K, Owens GK. Multiple repressor pathways contribute to phenotypic switching of vascular smooth muscle cells. Am J Physiol Cell Physiol 2007;292(1):C59-69.
Kawai-Kowase K, Ohshima T, Matsui H, Tanaka T, Shimizu T, Iso T, Arai M, Owens GK, Kurabayashi M. PIAS1 mediates TGFbeta-induced SM alpha-actin gene expression through inhibition of KLF4 function-expression by protein sumoylation. Arterioscler Thromb Vasc Biol 2009;29(1):99-106.
Kazi M, Lundmark K, Religa P, Gouda I, Larm O, Ray A, Swedenborg J, Hedin U. Inhibition of rat smooth muscle cell adhesion and proliferation by non-anticoagulant heparins. J Cell Physiol 2002;193(3):365-372.
Kenagy RD, Clowes AW. Regulation of baboon arterial smooth muscle cell plasminogen activators by heparin and growth factors. Thromb Res 1995;77(1):55-61.
Keynton RS, Rittgers SE, Shu MC. The effect of angle and flow rate upon hemodynamics in distal vascular graft anastomoses: an in vitro model study. J Biomech Eng 1991;113(4):458-463.
Keynton RS, Evancho MM, Sims RL, Rodway NV, Gobin A, Rittgers SE. Intimal hyperplasia and wall shear in arterial bypass graft distal anastomoses: an in vivo model study. J Biomech Eng 2001;123(5):464-473.
Keys KB, Andreopoulos FM, Peppas NA. Poly(ethylene glycol) star polymer hydrogels. MACROMOLECULES 1998;31(23):8149-8156.
Khaitlina SY. Functional specificity of actin isoforms. Int Rev Cytol 2001;202:35-98.
250 Kidd KR, Patula VB, Williams SK. Accelerated endothelialization of interpositional 1- mm vascular grafts. J Surg Res 2003;113(2):234-242.
Kim BS, Nikolovski J, Bonadio J, Mooney DJ. Cyclic mechanical strain regulates the development of engineered smooth muscle tissue. Nat Biotechnol 1999;17(10):979- 983.
Kim BS, Nikolovski J, Bonadio J, Smiley E, Mooney DJ. Engineered smooth muscle tissues: regulating cell phenotype with the scaffold. Exp Cell Res 1999;251(2):318- 328.
Kim JH, Kim JG, Kim D, Kim YH. Preparation and properties of PEG hydrogel from PEG macromonomer with sulfonate end group. J Appl Polym Sci 2005;96(1):56-61.
Kim HR, Gallant C, Leavis PC, Gunst SJ, Morgan KG. Cytoskeletal remodeling in differentiated vascular smooth muscle is actin isoform dependent and stimulus dependent. Am J Physiol Cell Physiol 2008;295(3):C768-778.
Kohler TR. Intimal Hyperplasia. In: White RA, Hollier LH, editors. Vascular surgery : basic science and clinical correlations. 2nd ed. Elmsford, N.Y.: Futura, 2004. p. p.
Kraiss LW, Kirkman TR, Kohler TR, Zierler B, Clowes AW. Shear stress regulates smooth muscle proliferation and neointimal thickening in porous polytetrafluoroethylene grafts. Arterioscler Thromb 1991;11(6):1844-1852.
Kumar V, Abbas AK, Fausto N, Robbins SL, Cotran RS. Robbins and Cotran pathologic basis of disease. 7th ed. Philadelphia: Elsevier Saunders, 2005.
Kurdikar DL, Peppas NA. A Kinetic-Study of Diacrylate Photopolymerizations. Polymer 1994;35(5):1004-1011.
Kuro-o M, Nagai R, Nakahara K, Katoh H, Tsai RC, Tsuchimochi H, Yazaki Y, Ohkubo A, Takaku F. cDNA cloning of a myosin heavy chain isoform in embryonic smooth muscle and its expression during vascular development and in arteriosclerosis. J Biol Chem 1991;266(6):3768-3773.
Langer R, Vacanti JP. Tissue Engineering. Science 1993;260(5110):920-926.
Larsen CC, Kligman F, Tang C, Kottke-Marchant K, Marchant RE. A biomimetic peptide fluorosurfactant polymer for endothelialization of ePTFE with limited platelet adhesion. Biomaterials 2007;28(24):3537-3548.
Laube HR, Duwe J, Rutsch W, Konertz W. Clinical experience with autologous endothelial cell-seeded polytetrafluoroethylene coronary artery bypass grafts. J Thorac Cardiovasc Surg 2000;120(1):134-141.
251 Lauzon AM, Tyska MJ, Rovner AS, Freyzon Y, Warshaw DM, Trybus KM. A 7-amino- acid insert in the heavy chain nucleotide binding loop alters the kinetics of smooth muscle myosin in the laser trap. J Muscle Res Cell Motil 1998;19(8):825-837.
Layne MD, Endege WO, Jain MK, Yet SF, Hsieh CM, Chin MT, Perrella MA, Blanar MA, Haber E, Lee ME. Aortic carboxypeptidase-like protein, a novel protein with discoidin and carboxypeptidase-like domains, is up-regulated during vascular smooth muscle cell differentiation. J Biol Chem 1998;273(25):15654-15660.
Layne MD, Yet SF, Maemura K, Hsieh CM, Liu X, Ith B, Lee ME, Perrella MA. Characterization of the mouse aortic carboxypeptidase-like protein promoter reveals activity in differentiated and dedifferentiated vascular smooth muscle cells. Circ Res 2002;90(6):728-736.
Lee SJ, Liu J, Oh SH, Soker S, Atala A, Yoo JJ. Development of a composite vascular scaffolding system that withstands physiological vascular conditions. Biomaterials 2008;29(19):2891-2898.
Leinweber BD, Leavis PC, Grabarek Z, Wang CL, Morgan KG. Extracellular regulated kinase (ERK) interaction with actin and the calponin homology (CH) domain of actin-binding proteins. Biochem J 1999;344 Pt 1:117-123.
Leinweber B, Parissenti AM, Gallant C, Gangopadhyay SS, Kirwan-Rhude A, Leavis PC, Morgan KG. Regulation of protein kinase C by the cytoskeletal protein calponin. J Biol Chem 2000;275(51):40329-40336.
Lepore JJ, Cheng L, Min Lu M, Mericko PA, Morrisey EE, Parmacek MS. High- efficiency somatic mutagenesis in smooth muscle cells and cardiac myocytes in SM22alpha-Cre transgenic mice. Genesis 2005;41(4):179-184.
L'Heureux N, Paquet S, Labbe R, Germain L, Auger FA. A completely biological tissue- engineered human blood vessel. Faseb Journal 1998;12(1):47-56.
L'Heureux N, Dusserre N, Konig G, Victor B, Keire P, Wight TN, Chronos NA, Kyles AE, Gregory CR, Hoyt G, Robbins RC, McAllister TN. Human tissue-engineered blood vessels for adult arterial revascularization. Nat Med 2006;12(3):361-365.
Li L, Miano JM, Cserjesi P, Olson EN. SM22 alpha, a marker of adult smooth muscle, is expressed in multiple myogenic lineages during embryogenesis. Circ Res 1996;78(2):188-195.
Li Q, Muragaki Y, Hatamura I, Ueno H, Ooshima A. Stretch-induced collagen synthesis in cultured smooth muscle cells from rabbit aortic media and a possible involvement of angiotensin II and transforming growth factor-beta. J Vasc Res 1998;35(2):93-103.
Li G, Chen SJ, Oparil S, Chen YF, Thompson JA. Direct in vivo evidence demonstrating neointimal migration of adventitial fibroblasts after balloon injury of rat carotid arteries. Circulation 2000;101(12):1362-1365.
252 Li S, Lao J, Chen BP, Li YS, Zhao Y, Chu J, Chen KD, Tsou TC, Peck K, Chien S. Genomic analysis of smooth muscle cells in 3-dimensional collagen matrix. Faseb J 2003;17(1):97-99.
Li S, Wang DZ, Wang Z, Richardson JA, Olson EN. The serum response factor coactivator myocardin is required for vascular smooth muscle development. Proc Natl Acad Sci U S A 2003;100(16):9366-9370.
Li W, Chen Q, Mills I, Sumpio BE. Involvement of S6 kinase and p38 mitogen activated protein kinase pathways in strain-induced alignment and proliferation of bovine aortic smooth muscle cells. J Cell Physiol 2003;195(2):202-209.
Lin HQ, Kai T, Freeman BD, Kalakkunnath S, Kalika DS. The effect of cross-linking on gas permeability in cross-linked poly(ethylene glycol diacrylate). MACROMOLECULES 2005;38(20):8381-8393.
Lindner V, Olson NE, Clowes AW, Reidy MA. Inhibition of smooth muscle cell proliferation in injured rat arteries. Interaction of heparin with basic fibroblast growth factor. J Clin Invest 1992;90(5):2044-2049.
Lin-Gibson S, Bencherif S, Cooper JA, Wetzel SJ, Antonucci JM, Vogel BM, Horkay F, Washburn NR. Synthesis and characterization of PEG dimethacrylates and their hydrogels. Biomacromolecules 2004;5(4):1280-1287.
Lin-Gibson S, Jones RL, Washburn NR, Horkay F. Structure-property relationships of photopolymerizable poly(ethylene glycol) dimethacrylate hydrogels. MACROMOLECULES 2005;38(7):2897-2902.
Liu Y, Sinha S, Owens G. A transforming growth factor-beta control element required for SM alpha-actin expression in vivo also partially mediates GKLF-dependent transcriptional repression. J Biol Chem 2003;278(48):48004-48011.
Liu Y, Sinha S, McDonald OG, Shang Y, Hoofnagle MH, Owens GK. Kruppel-like factor 4 abrogates myocardin-induced activation of smooth muscle gene expression. J Biol Chem 2005;280(10):9719-9727.
Liu JY, Swartz DD, Peng HF, Gugino SF, Russell JA, Andreadis ST. Functional tissue- engineered blood vessels from bone marrow progenitor cells. Cardiovasc Res 2007;75(3):618-628.
Liu B, Qu MJ, Qin KR, Li H, Li ZK, Shen BR, Jiang ZL. Role of cyclic strain frequency in regulating the alignment of vascular smooth muscle cells in vitro. Biophys J 2008;94(4):1497-1507.
Liu JY, Peng HF, Andreadis ST. Contractile smooth muscle cells derived from hair- follicle stem cells. Cardiovasc Res 2008;79(1):24-33.
253 Lloyd-Jones D, Adams R, Carnethon M, De Simone G, Ferguson TB, Flegal K, Ford E, Furie K, Go A, Greenlund K, Haase N, Hailpern S, Ho M, Howard V, Kissela B, Kittner S, Lackland D, Lisabeth L, Marelli A, McDermott M, Meigs J, Mozaffarian D, Nichol G, O'Donnell C, Roger V, Rosamond W, Sacco R, Sorlie P, Stafford R, Steinberger J, Thom T, Wasserthiel-Smoller S, Wong N, Wylie-Rosett J, Hong Y, for the American Heart Association Statistics Committee and Stroke Statistics S. Heart Disease and Stroke Statistics--2009 Update: A Report From the American Heart Association Statistics Committee and Stroke Statistics Subcommittee. Circulation 2009;119(3):e21-181.
Lutolf MP, Hubbell JA. Synthesis and physicochemical characterization of end-linked poly(ethylene glycol)-co-peptide hydrogels formed by Michael-type addition. Biomacromolecules 2003;4(3):713-722.
Lutolf MP, Lauer-Fields JL, Schmoekel HG, Metters AT, Weber FE, Fields GB, Hubbell JA. Synthetic matrix metalloproteinase-sensitive hydrogels for the conduction of tissue regeneration: engineering cell-invasion characteristics. Proc Natl Acad Sci U S A 2003;100(9):5413-5418.
Mack CP, Somlyo AV, Hautmann M, Somlyo AP, Owens GK. Smooth muscle differentiation marker gene expression is regulated by RhoA-mediated actin polymerization. J Biol Chem 2001;276(1):341-347.
Mack CP, Hinson JS. Regulation of smooth muscle differentiation by the myocardin family of serum response factor co-factors. J Thromb Haemost 2005;3(9):1976-1984.
Mahoney MJ, Anseth KS. Three-dimensional growth and function of neural tissue in degradable polyethylene glycol hydrogels. Biomaterials 2006;27(10):2265-2274.
Manabe I, Owens GK. Recruitment of serum response factor and hyperacetylation of histones at smooth muscle-specific regulatory regions during differentiation of a novel P19-derived in vitro smooth muscle differentiation system. Circ Res 2001;88(11):1127-1134.
Mann BK, Tsai AT, Scott-Burden T, West JL. Modification of surfaces with cell adhesion peptides alters extracellular matrix deposition. Biomaterials 1999;20(23- 24):2281-2286.
Mann BK, Gobin AS, Tsai AT, Schmedlen RH, West JL. Smooth muscle cell growth in photopolymerized hydrogels with cell adhesive and proteolytically degradable domains: synthetic ECM analogs for tissue engineering. Biomaterials 2001;22(22):3045-3051.
Mann BK, Schmedlen RH, West JL. Tethered-TGF-beta increases extracellular matrix production of vascular smooth muscle cells. Biomaterials 2001;22(5):439-444.
254 Mann BK, West JL. Cell adhesion peptides alter smooth muscle cell adhesion, proliferation, migration, and matrix protein synthesis on modified surfaces and in polymer scaffolds. J Biomed Mater Res 2002;60(1):86-93.
Martens P, Anseth KS. Characterization of hydrogels formed from acrylate modified poly(vinyl alcohol) macromers. Polymer 2000;41(21):7715-7722.
Masters KS, Shah DN, Leinwand LA, Anseth KS. Crosslinked hyaluronan scaffolds as a biologically active carrier for valvular interstitial cells. Biomaterials 2005;26(15):2517-2525.
Masuki S, Takeoka M, Taniguchi S, Nose H. Enhanced baroreflex sensitivity in free- moving calponin knockout mice. Am J Physiol Heart Circ Physiol 2003;284(3):H939-946.
Masuki S, Takeoka M, Taniguchi S, Yokoyama M, Nose H. Impaired arterial pressure regulation during exercise due to enhanced muscular vasodilatation in calponin knockout mice. J Physiol 2003;553(Pt 1):203-212.
Matsumura G, Hibino N, Ikada Y, Kurosawa H, Shin'oka T. Successful application of tissue engineered vascular autografts: clinical experience. Biomaterials 2003;24(13):2303-2308.
Matthew JD, Khromov AS, McDuffie MJ, Somlyo AV, Somlyo AP, Taniguchi S, Takahashi K. Contractile properties and proteins of smooth muscles of a calponin knockout mouse. J Physiol 2000;529 Pt 3:811-824.
McAllister TN, Maruszewski M, Garrido SA, Wystrychowski W, Dusserre N, Marini A, Zagalski K, Fiorillo A, Avila H, Manglano X, Antonelli J, Kocher A, Zembala M, Cierpka L, de la Fuente LM, L'Heureux N. Effectiveness of haemodialysis access with an autologous tissue-engineered vascular graft: a multicentre cohort study. The Lancet 2009;373(9673):1440-1446.
Mecham RP, Hinek A, Entwistle R, Wrenn DS, Griffin GL, Senior RM. Elastin binds to a multifunctional 67-kilodalton peripheral membrane protein. Biochemistry 1989;28(9):3716-3722.
Meinhart JG, Deutsch M, Fischlein T, Howanietz N, Froschl A, Zilla P. Clinical autologous in vitro endothelialization of 153 infrainguinal ePTFE grafts. Ann Thorac Surg 2001;71(5 Suppl):S327-331.
Merrill EW, Dennison KA, Sung C. Partitioning and diffusion of solutes in hydrogels of poly(ethylene oxide). Biomaterials 1993;14(15):1117-1126.
Mezgueldi M, Mendre C, Calas B, Kassab R, Fattoum A. Characterization of the regulatory domain of gizzard calponin. Interactions of the 145-163 region with F- actin, calcium-binding proteins, and tropomyosin. J Biol Chem 1995;270(15):8867- 8876.
255 Miano JM, Cserjesi P, Ligon KL, Periasamy M, Olson EN. Smooth muscle myosin heavy chain exclusively marks the smooth muscle lineage during mouse embryogenesis. Circ Res 1994;75(5):803-812.
Miano JM, Olson EN. Expression of the smooth muscle cell calponin gene marks the early cardiac and smooth muscle cell lineages during mouse embryogenesis. J Biol Chem 1996;271(12):7095-7103.
Miano JM. Serum response factor: toggling between disparate programs of gene expression. J Mol Cell Cardiol 2003;35(6):577-593.
Miano JM, Long X, Fujiwara K. Serum response factor: master regulator of the actin cytoskeleton and contractile apparatus. Am J Physiol Cell Physiol 2007;292(1):C70- 81.
Miller JH, Foreman RK, Ferguson L, Faris I. Interposition vein cuff for anastomosis of prosthesis to small artery. Aust N Z J Surg 1984;54(3):283-285.
Millette E, Rauch BH, Defawe O, Kenagy RD, Daum G, Clowes AW. Platelet-derived growth factor-BB-induced human smooth muscle cell proliferation depends on basic FGF release and FGFR-1 activation. Circ Res 2005;96(2):172-179.
Mino T, Yuasa U, Nakamura F, Naka M, Tanaka T. Two distinct actin-binding sites of smooth muscle calponin. Eur J Biochem 1998;251(1-2):262-268.
Miralles F, Posern G, Zaromytidou AI, Treisman R. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 2003;113(3):329-342.
Moiseeva EP. Adhesion receptors of vascular smooth muscle cells and their functions. Cardiovasc Res 2001;52(3):372-386.
Morgan KG, Gangopadhyay SS. Invited review: cross-bridge regulation by thin filament- associated proteins. J Appl Physiol 2001;91(2):953-962.
Motwani JG, Topol EJ. Aortocoronary saphenous vein graft disease: pathogenesis, predisposition, and prevention. Circulation 1998;97(9):916-931.
Newby AC. Matrix metalloproteinases regulate migration, proliferation, and death of vascular smooth muscle cells by degrading matrix and non-matrix substrates. Cardiovascular Research 2006;69(3):614-624.
Niessen P, Clement S, Fontao L, Chaponnier C, Teunissen B, Rensen S, van Eys G, Gabbiani G. Biochemical evidence for interaction between smoothelin and filamentous actin. Exp Cell Res 2004;292(1):170-178.
Niessen P, Rensen S, van Deursen J, De Man J, De Laet A, Vanderwinden JM, Wedel T, Baker D, Doevendans P, Hofker M, Gijbels M, van Eys G. Smoothelin-a is essential
256 for functional intestinal smooth muscle contractility in mice. Gastroenterology 2005;129(5):1592-1601.
Niklason LE, Gao J, Abbott WM, Hirschi KK, Houser S, Marini R, Langer R. Functional arteries grown in vitro. Science 1999;284(5413):489-493.
Nikolovski J, Kim BS, Mooney DJ. Cyclic strain inhibits switching of smooth muscle cells to an osteoblast-like phenotype. Faseb J 2003;17(3):455-457.
Nishimura G, Manabe I, Tsushima K, Fujiu K, Oishi Y, Imai Y, Maemura K, Miyagishi M, Higashi Y, Kondoh H, Nagai R. DeltaEF1 mediates TGF-beta signaling in vascular smooth muscle cell differentiation. Dev Cell 2006;11(1):93-104.
Norman C, Runswick M, Pollock R, Treisman R. Isolation and properties of cDNA clones encoding SRF, a transcription factor that binds to the c-fos serum response element. Cell 1988;55(6):989-1003.
Nuttelman CR, Tripodi MC, Anseth KS. In vitro osteogenic differentiation of human mesenchymal stem cells photoencapsulated in PEG hydrogels. J Biomed Mater Res A 2004;68(4):773-782.
Nuttelman CR, Benoit DS, Tripodi MC, Anseth KS. The effect of ethylene glycol methacrylate phosphate in PEG hydrogels on mineralization and viability of encapsulated hMSCs. Biomaterials 2006;27(8):1377-1386.
Nwasokwa ON. Coronary-Artery Bypass Graft Disease. Ann Intern Med 1995;123(7):528-545.
O'Callaghan CJ, Williams B. Mechanical strain-induced extracellular matrix production by human vascular smooth muscle cells: role of TGF-beta(1). Hypertension 2000;36(3):319-324.
Odian GG. Principles of polymerization. 4th ed. Hoboken, N.J.: Wiley-Interscience, 2004.
Ojha M, Ethier CR, Johnston KW, Cobbold RS. Steady and pulsatile flow fields in an end-to-side arterial anastomosis model. J Vasc Surg 1990;12(6):747-753.
Okamoto E, Couse T, De Leon H, Vinten-Johansen J, Goodman RB, Scott NA, Wilcox JN. Perivascular inflammation after balloon angioplasty of porcine coronary arteries. Circulation 2001;104(18):2228-2235.
Orlandi A, Ropraz P, Gabbiani G. Proliferative Activity and Alpha-Smooth Muscle Actin Expression in Cultured Rat Aortic Smooth-Muscle Cells Are Differently Modulated by Transforming Growth-Factor-Beta-1 and Heparin. Exp Cell Res 1994;214(2):528- 536.
257 Owens GK, Loeb A, Gordon D, Thompson MM. Expression of smooth muscle-specific alpha-isoactin in cultured vascular smooth muscle cells: relationship between growth and cytodifferentiation. J Cell Biol 1986;102(2):343-352.
Owens GK, Geisterfer AA, Yang YW, Komoriya A. Transforming growth factor-beta- induced growth inhibition and cellular hypertrophy in cultured vascular smooth muscle cells. J Cell Biol 1988;107(2):771-780.
Owens GK. Regulation of differentiation of vascular smooth muscle cells. Physiol Rev 1995;75(3):487-517.
Owens GK, Kumar MS, Wamhoff BR. Molecular regulation of vascular smooth muscle cell differentiation in development and disease. Physiol Rev 2004;84(3):767-801.
Padmavathi NC, Chatterji PR. Structural characteristics and swelling behavior of poly(ethylene glycol) diacrylate hydrogels. MACROMOLECULES 1996;29(6):1976- 1979.
Papetti M, Shujath J, Riley KN, Herman IM. FGF-2 antagonizes the TGF-beta1-mediated induction of pericyte alpha-smooth muscle actin expression: a role for myf-5 and Smad-mediated signaling pathways. Invest Ophthalmol Vis Sci 2003;44(11):4994- 5005.
Parks WC, Mecham RP. Matrix metalloproteinases. San Diego: Academic Press, 1998.
Patel RC, Handy I, Patel CV. Contribution of double-stranded RNA-activated protein kinase toward antiproliferative actions of heparin on vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 2002;22(9):1439-1444.
Patterson J, Hubbell JA. Molecularly Engineered PEG Hydrogels with Enhanced Proteolysis. Society for Biomaterials Annual Meeting; 2009 April 22-25; San Antonio, TX; 2009. p. ABSTRACT.
Pavlov G, Finet S, Tatarenko K, Korneeva E, Ebel C. Conformation of heparin studied with macromolecular hydrodynamic methods and X-ray scattering. Eur Biophys J Biophys Lett 2003;32(5):437-449.
Pearson TA, Blair SN, Daniels SR, Eckel RH, Fair JM, Fortmann SP, Franklin BA, Goldstein LB, Greenland P, Grundy SM, Hong Y, Houston Miller N, Lauer RM, Ockene IS, Sacco RL, Sallis JF, Jr., Smith SC, Jr., Stone NJ, Taubert KA. AHA Guidelines for Primary Prevention of Cardiovascular Disease and Stroke: 2002 Update: Consensus Panel Guide to Comprehensive Risk Reduction for Adult Patients Without Coronary or Other Atherosclerotic Vascular Diseases. Circulation 2002;106(3):388-391.
Pektok E, Nottelet B, Tille JC, Gurny R, Kalangos A, Moeller M, Walpoth BH. Degradation and Healing Characteristics of Small-Diameter Poly(epsilon-
258 Caprolactone) Vascular Grafts in the Rat Systemic Arterial Circulation. Circulation 2008;118(24):2563-2570.
Peppas NA. Hydrogels in medicine and pharmacy. Boca Raton, Fla.: CRC Press, 1986.
Peyton SR, Raub CB, Keschrumrus VP, Putnam AJ. The use of poly(ethylene glycol) hydrogels to investigate the impact of ECM chemistry and mechanics on smooth muscle cells. Biomaterials 2006;27(28):4881-4893.
Peyton SR, Ghajar CM, Khatiwala CB, Putnam AJ. The emergence of ECM mechanics and cytoskeletal tension as important regulators of cell function. Cell Biochem Biophys 2007;47(2):300-320.
Peyton SR, Kim PD, Ghajar CM, Seliktar D, Putnam AJ. The effects of matrix stiffness and RhoA on the phenotypic plasticity of smooth muscle cells in a 3-D biosynthetic hydrogel system. Biomaterials 2008;29(17):2597-2607.
Pierschbacher MD, Ruoslahti E. Cell attachment activity of fibronectin can be duplicated by small synthetic fragments of the molecule. Nature 1984;309(5963):30-33.
Pipes GC, Sinha S, Qi X, Zhu CH, Gallardo TD, Shelton J, Creemers EE, Sutherland L, Richardson JA, Garry DJ, Wright WE, Owens GK, Olson EN. Stem cells and their derivatives can bypass the requirement of myocardin for smooth muscle gene expression. Dev Biol 2005;288(2):502-513.
Poh M, Boyer M, Solan A, Dahl SLM, Pedrotty D, Banik SSR, McKee JA, Klinger RY, Counter CM, Niklason LE. Blood vessels engineered from human cells. Lancet 2005;365(9477):2122-2124.
Prywes R, Roeder RG. Purification of the c-fos enhancer-binding protein. Mol Cell Biol 1987;7(10):3482-3489.
Pukac LA, Carter JE, Ottlinger ME, Karnovsky MJ. Mechanisms of inhibition by heparin of PDGF stimulated MAP kinase activation in vascular smooth muscle cells. J Cell Physiol 1997;172(1):69-78.
Pukac L, Huangpu J, Karnovsky MJ. Platelet-derived growth factor-BB, insulin-like growth factor-I, and phorbol ester activate different signaling pathways for stimulation of vascular smooth muscle cell migration. Exp Cell Res 1998;242(2):548- 560.
Qin H, Ishiwata T, Wang R, Kudo M, Yokoyama M, Naito Z, Asano G. Effects of extracellular matrix on phenotype modulation and MAPK transduction of rat aortic smooth muscle cells in vitro. Exp Mol Pathol 2000;69(2):79-90.
Qu MJ, Liu B, Wang HQ, Yan ZQ, Shen BR, Jiang ZL. Frequency-dependent phenotype modulation of vascular smooth muscle cells under cyclic mechanical strain. J Vasc Res 2007;44(5):345-353.
259 Quarto N, Amalric F. Heparan sulfate proteoglycans as transducers of FGF-2 signalling. J Cell Sci 1994;107 ( Pt 11):3201-3212.
Raines EW, Koyama H, Carragher NO. The extracellular matrix dynamically regulates smooth muscle cell responsiveness to PDGF. Ann N Y Acad Sci 2000;902:39-51; discussion 51-32.
Rapraeger AC, Krufka A, Olwin BB. Requirement of heparan sulfate for bFGF-mediated fibroblast growth and myoblast differentiation. Science 1991;252(5013):1705-1708.
Rauch BH, Millette E, Kenagy RD, Daum G, Clowes AW. Thrombin- and factor Xa- induced DNA synthesis is mediated by transactivation of fibroblast growth factor receptor-1 in human vascular smooth muscle cells. Circ Res 2004;94(3):340-345.
Rauch BH, Millette E, Kenagy RD, Daum G, Fischer JW, Clowes AW. Syndecan-4 is required for thrombin-induced migration and proliferation in human vascular smooth muscle cells. J Biol Chem 2005;280(17):17507-17511.
Ravi N, Mitra A, Hamilton P, Horkay F. Characterization of the network properties of poly(ethylene glycol)-acrylate hydrogels prepared by variations in the ethanol-water solvent composition during crosslinking copolymerization. Journal of Polymer Science Part B-Polymer Physics 2002;40(23):2677-2684.
Reidy MA. Regulation of Arterial Smooth Muscle Growth. In: Schwartz SM, Mecham RP, editors. The vascular smooth muscle cell : molecular and biological responses to the extracellular matrix. San Diego: Academic Press, 1995. p. xvi, 410 p.
Rensen SS, Thijssen VL, De Vries CJ, Doevendans PA, Detera-Wadleigh SD, Van Eys GJ. Expression of the smoothelin gene is mediated by alternative promoters. Cardiovasc Res 2002;55(4):850-863.
Rensen SS, Niessen PM, van Deursen JM, Janssen BJ, Heijman E, Hermeling E, Meens M, Lie N, Gijbels MJ, Strijkers GJ, Doevendans PA, Hofker MH, De Mey JG, van Eys GJ. Smoothelin-B deficiency results in reduced arterial contractility, hypertension, and cardiac hypertrophy in mice. Circulation 2008;118(8):828-836.
Reusch P, Wagdy H, Reusch R, Wilson E, Ives HE. Mechanical strain increases smooth muscle and decreases nonmuscle myosin expression in rat vascular smooth muscle cells. Circ Res 1996;79(5):1046-1053.
Reusch HP, Chan G, Ives HE, Nemenoff RA. Activation of JNK/SAPK and ERK by mechanical strain in vascular smooth muscle cells depends on extracellular matrix composition. Biochem Biophys Res Commun 1997;237(2):239-244.
Rose SL, Babensee JE. Complimentary endothelial cell/smooth muscle cell co-culture systems with alternate smooth muscle cell phenotypes. Ann Biomed Eng 2007;35(8):1382-1390.
260 Rose SL, Babensee JE. Smooth muscle cell phenotype alters cocultured endothelial cell response to biomaterial-pretreated leukocytes. J Biomed Mater Res A 2008;84(3):661-671.
Ross R. Atherosclerosis -- An Inflammatory Disease. N Engl J Med 1999;340(2):115- 126.
Rowlands AS, Cooper-White JJ. Directing phenotype of vascular smooth muscle cells using electrically stimulated conducting polymer. Biomaterials 2008;29(34):4510- 4520.
Rozenblum GT, Gimona M. Calponins: adaptable modular regulators of the actin cytoskeleton. Int J Biochem Cell Biol 2008;40(10):1990-1995.
Rzucidlo EM, Martin KA, Powell RJ. Regulation of vascular smooth muscle cell differentiation. J Vasc Surg 2007;45 Suppl A:A25-32.
Sagnella S, Anderson E, Sanabria N, Marchant RE, Kottke-Marchant K. Human endothelial cell interaction with biomimetic surfactant polymers containing Peptide ligands from the heparin binding domain of fibronectin. Tissue Eng 2005;11(1- 2):226-236.
Salacinski HJ, Goldner S, Giudiceandrea A, Hamilton G, Seifalian AM, Edwards A, Carson RJ. The mechanical behavior of vascular grafts: a review. J Biomater Appl 2001;15(3):241-278.
Sapsford RN, Oakley GD, Talbot S. Early and late patency of expanded polytetrafluoroethylene vascular grafts in aorta-coronary bypass. J Thorac Cardiovasc Surg 1981;81(6):860-864.
Savage JM, Gilotti AC, Granzow CA, Molina F, Lowe-Krentz LJ. Antibodies against a putative heparin receptor slow cell proliferation and decrease MAPK activation in vascular smooth muscle cells. J Cell Physiol 2001;187(3):283-293.
Sawhney AS, Pathak CP, Hubbell JA. Bioerodible Hydrogels Based on Photopolymerized Poly(Ethylene Glycol)-Co-Poly(Alpha-Hydroxy Acid) Diacrylate Macromers. MACROMOLECULES 1993;26(4):581-587.
Sayers RD, Raptis S, Berce M, Miller JH. Long-term results of femorotibial bypass with vein or polytetrafluoroethylene. Br J Surg 1998;85(7):934-938.
Schaner PJ, Martin ND, Tulenko TN, Shapiro IM, Tarola NA, Leichter RF, Carabasi RA, DiMuzio PJ. Decellularized vein as a potential scaffold for vascular tissue engineering. Journal of Vascular Surgery 2004;40(1):146-153.
Schneider F, O'Connor S, Becquemin JP. Efficacy of Collagen Silver-Coated Polyester and Rifampin-Soaked Vascular Grafts to Resist Infection from MRSA and Escherichia coli in a Dog Model. Annals of Vascular Surgery 2008;22(6):815-821.
261 Schultz K, Murthy V, Tatro JB, Beasley D. Endogenous interleukin-1 alpha promotes a proliferative and proinflammatory phenotype in human vascular smooth muscle cells. Am J Physiol Heart Circ Physiol 2007;292(6):H2927-2934.
Seliktar D, Black RA, Vito RP, Nerem RM. Dynamic mechanical conditioning of collagen-gel blood vessel constructs induces remodeling in vitro. Ann Biomed Eng 2000;28(4):351-362.
Seliktar D, Nerem RM, Galis ZS. Mechanical strain-stimulated remodeling of tissue- engineered blood vessel constructs. Tissue Engineering 2003;9(4):657-666.
Shen JY, Chan-Park MB, He B, Zhu AP, Zhu X, Beuerman RW, Yang EB, Chen W, Chan V. Three-dimensional microchannels in biodegradable polymeric films for control orientation and phenotype of vascular smooth muscle cells. Tissue Eng 2006;12(8):2229-2240.
Shin H, Jo S, Mikos AG. Modulation of marrow stromal osteoblast adhesion on biomimetic oligo[poly(ethylene glycol) fumarate] hydrogels modified with Arg-Gly- Asp peptides and a poly(ethyleneglycol) spacer. J Biomed Mater Res 2002;61(2):169- 179.
Shin'oka T, Matsumura G, Hibino N, Naito Y, Watanabe M, Konuma T, Sakamoto T, Nagatsu M, Kurosawa H. Midterm clinical result of tissue-engineered vascular autografts seeded with autologous bone marrow cells. J Thorac Cardiovasc Surg 2005;129(6):1330-1338.
Shum-Tim D, Stock U, Hrkach J, Shinoka T, Lien J, Moses MA, Stamp A, Taylor G, Moran AM, Landis W, Langer R, Vacanti JP, Mayer JE. Tissue engineering of autologous aorta using a new biodegradable polymer. Ann Thorac Surg 1999;68(6):2298-2304.
Sill TJ, von Recum HA. Electrospinning: applications in drug delivery and tissue engineering. Biomaterials 2008;29(13):1989-2006.
Smeds KA, Pfister-Serres A, Miki D, Dastgheib K, Inoue M, Hatchell DL, Grinstaff MW. Photocrosslinkable polysaccharides for in situ hydrogel formation. J Biomed Mater Res 2001;54(1):115-121.
Smith SC, Jr., Allen J, Blair SN, Bonow RO, Brass LM, Fonarow GC, Grundy SM, Hiratzka L, Jones D, Krumholz HM, Mosca L, Pasternak RC, Pearson T, Pfeffer MA, Taubert KA. AHA/ACC Guidelines for Secondary Prevention for Patients With Coronary and Other Atherosclerotic Vascular Disease: 2006 Update: Endorsed by the National Heart, Lung, and Blood Institute. Circulation 2006;113(19):2363-2372.
Sobue K, Sellers JR. Caldesmon, a novel regulatory protein in smooth muscle and nonmuscle actomyosin systems. J Biol Chem 1991;266(19):12115-12118.
262 Sottiurai VS, Yao JS, Batson RC, Sue SL, Jones R, Nakamura YA. Distal anastomotic intimal hyperplasia: histopathologic character and biogenesis. Ann Vasc Surg 1989;3(1):26-33.
Spofford CM, Chilian WM. The elastin-laminin receptor functions as a mechanotransducer in vascular smooth muscle. Am J Physiol Heart Circ Physiol 2001;280(3):H1354-1360.
Staatz WD, Fok KF, Zutter MM, Adams SP, Rodriguez BA, Santoro SA. Identification of a tetrapeptide recognition sequence for the alpha 2 beta 1 integrin in collagen. J Biol Chem 1991;266(12):7363-7367.
Standley PR, Obards TJ, Martina CL. Cyclic stretch regulates autocrine IGF-I in vascular smooth muscle cells: implications in vascular hyperplasia. Am J Physiol 1999;276(4 Pt 1):E697-705.
Standley PR, Cammarata A, Nolan BP, Purgason CT, Stanley MA. Cyclic stretch induces vascular smooth muscle cell alignment via NO signaling. Am J Physiol Heart Circ Physiol 2002;283(5):H1907-1914.
Stegemann JP, Nerem RM. Altered response of vascular smooth muscle cells to exogenous biochemical stimulation in two- and three-dimensional culture. Exp Cell Res 2003;283(2):146-155.
Stegemann JP, Nerem RM. Phenotype modulation in vascular tissue engineering using biochemical and mechanical stimulation. Ann Biomed Eng 2003;31(4):391-402.
Stegemann JP, Dey NB, Lincoln TM, Nerem RM. Genetic modification of smooth muscle cells to control phenotype and function in vascular tissue engineering. Tissue Eng 2004;10(1-2):189-199.
Stegemann JP, Hong H, Nerem RM. Mechanical, biochemical, and extracellular matrix effects on vascular smooth muscle cell phenotype. J Appl Physiol 2005;98(6):2321- 2327.
Stitzel J, Liu L, Lee SJ, Komura M, Berry J, Soker S, Lim G, Van Dyke M, Czerw R, Yoo JJ, Atala A. Controlled fabrication of a biological vascular substitute. Biomaterials 2006;27(7):1088-1094.
Sun Q, Chen G, Streb JW, Long X, Yang Y, Stoeckert CJ, Jr., Miano JM. Defining the mammalian CArGome. Genome Res 2006;16(2):197-207.
Swartz DD, Russell JA, Andreadis ST. Engineering of fibrin-based functional and implantable small-diameter blood vessels. American Journal of Physiology-Heart and Circulatory Physiology 2005;288(3):H1451-H1460.
263 Tanaka H, Sukhova GK, Swanson SJ, Clinton SK, Ganz P, Cybulsky MI, Libby P. Sustained Activation of Vascular Cells and Leukocytes in the Rabbit Aorta after Balloon Injury. Circulation 1993;88(4):1788-1803.
Taylor RS, Loh A, McFarland RJ, Cox M, Chester JF. Improved technique for polytetrafluoroethylene bypass grafting: long-term results using anastomotic vein patches. Br J Surg 1992;79(4):348-354.
Teebken OE, Pichlmaier AM, Haverich A. Cell seeded decellularised allogeneic matrix grafts and biodegradable polydioxanone-prostheses compared with arterial autografts in a porcine model. Eur J Vasc Endovasc Surg 2001;22(2):139-145.
Thomae KR, Nakayama DK, Billiar TR, Simmons RL, Pitt BR, Davies P. The effect of nitric oxide on fetal pulmonary artery smooth muscle growth. J Surg Res 1995;59(3):337-343.
Thyberg J, Hultgardh-Nilsson A. Fibronectin and the basement membrane components laminin and collagen type IV influence the phenotypic properties of subcultured rat aortic smooth muscle cells differently. Cell Tissue Res 1994;276(2):263-271.
Tillman BW, Yazdani SK, Lee SJ, Geary RL, Atala A, Yoo JJ. The in vivo stability of electrospun polycaprolactone-collagen scaffolds in vascular reconstruction. Biomaterials 2009;30(4):583-588.
Tock J, Van Putten V, Stenmark KR, Nemenoff RA. Induction of SM-alpha-actin expression by mechanical strain in adult vascular smooth muscle cells is mediated through activation of JNK and p38 MAP kinase. Biochem Biophys Res Commun 2003;301(4):1116-1121.
Topouzis S, Majesky MW. Smooth muscle lineage diversity in the chick embryo. Two types of aortic smooth muscle cell differ in growth and receptor-mediated transcriptional responses to transforming growth factor-beta. Dev Biol 1996;178(2):430-445.
Torres-Lugo M, Peppas NA. Preparation and Characterization of P(MAA-g-EG) Nanospheres for Protein Delivery Applications. Journal of Nanoparticle Research 2002;4(1):73-81.
Tran T, McNeill KD, Gerthoffer WT, Unruh H, Halayko AJ. Endogenous laminin is required for human airway smooth muscle cell maturation. Respir Res 2006;7:117.
Tranquillo RT, Girton TS, Bromberek BA, Triebes TG, Mooradian DL. Magnetically orientated tissue-equivalent tubes: Application to a circumferentially orientated media-equivalent. Biomaterials 1996;17(3):349-357.
Treisman R. Identification of a protein-binding site that mediates transcriptional response of the c-fos gene to serum factors. Cell 1986;46(4):567-574.
264 Trubel W, Schima H, Czerny M, Perktold K, Schimek MG, Polterauer P. Experimental comparison of four methods of end-to-side anastomosis with expanded polytetrafluoroethylene. Br J Surg 2004;91(2):159-167.
Turla MB, Thompson MM, Corjay MH, Owens GK. Mechanisms of angiotensin II- and arginine vasopressin-induced increases in protein synthesis and content in cultured rat aortic smooth muscle cells. Evidence for selective increases in smooth muscle isoactin expression. Circ Res 1991;68(1):288-299. van der Loop FT, Schaart G, Timmer ED, Ramaekers FC, van Eys GJ. Smoothelin, a novel cytoskeletal protein specific for smooth muscle cells. J Cell Biol 1996;134(2):401-411. van Eys GJ, Niessen PM, Rensen SS. Smoothelin in vascular smooth muscle cells. Trends Cardiovasc Med 2007;17(1):26-30.
Van Tomme SR, van Nostrum CF, Dijkstra M, De Smedt SC, Hennink WE. Effect of particle size and charge on the network properties of microsphere-based hydrogels. Eur J Pharm Biopharm 2008;70(2):522-530. van Tuyn J, Knaan-Shanzer S, van de Watering MJ, de Graaf M, van der Laarse A, Schalij MJ, van der Wall EE, de Vries AA, Atsma DE. Activation of cardiac and smooth muscle-specific genes in primary human cells after forced expression of human myocardin. Cardiovasc Res 2005;67(2):245-255.
Veith FJ, Gupta S, Daly V. Management of early and late thrombosis of expanded polytetrafluoroethylene (PTFE) femoropopliteal bypass grafts: favorable prognosis with appropriate reoperation. Surgery 1980;87(5):581-587.
Walden R, L'Italien GJ, Megerman J, Abbott WM. Matched elastic properties and successful arterial grafting. Arch Surg 1980;115(10):1166-1169.
Wamhoff BR, Bowles DK, McDonald OG, Sinha S, Somlyo AP, Somlyo AV, Owens GK. L-type voltage-gated Ca2+ channels modulate expression of smooth muscle differentiation marker genes via a rho kinase/myocardin/SRF-dependent mechanism. Circ Res 2004;95(4):406-414.
Wang D, Chang PS, Wang Z, Sutherland L, Richardson JA, Small E, Krieg PA, Olson EN. Activation of cardiac gene expression by myocardin, a transcriptional cofactor for serum response factor. Cell 2001;105(7):851-862.
Wang Z, Wang DZ, Pipes GC, Olson EN. Myocardin is a master regulator of smooth muscle gene expression. Proc Natl Acad Sci U S A 2003;100(12):7129-7134.
Wang Z, Wang DZ, Hockemeyer D, McAnally J, Nordheim A, Olson EN. Myocardin and ternary complex factors compete for SRF to control smooth muscle gene expression. Nature 2004;428(6979):185-189.
265 Ward JH, Furman K, Peppas NA. Effect of monomer type and dangling end size on polymer network synthesis. J Appl Polym Sci 2003;89(13):3506-3519.
Watanabe T, Pakala R, Katagiri T, Benedict CR. Serotonin potentiates angiotensin II - induced vascular smooth muscle cell proliferation. Atherosclerosis 2001;159(2):269- 279.
Weber LM, Hayda KN, Haskins K, Anseth KS. The effects of cell-matrix interactions on encapsulated beta-cell function within hydrogels functionalized with matrix-derived adhesive peptides. Biomaterials 2007;28(19):3004-3011.
Weinberg CB, Bell E. A blood vessel model constructed from collagen and cultured vascular cells. Science 1986;231(4736):397-400.
Welser JV, Lange N, Singer CA, Elorza M, Scowen P, Keef KD, Gerthoffer WT, Burkin DJ. Loss of the alpha7 integrin promotes extracellular signal-regulated kinase activation and altered vascular remodeling. Circ Res 2007;101(7):672-681.
West JL, Hubbell JA. Polymeric biomaterials with degradation sites for proteases involved in cell migration. MACROMOLECULES 1999;32(1):241-244.
Weston MW, Rhee K, Tarbell JM. Compliance and diameter mismatch affect the wall shear rate distribution near an end-to-end anastomosis. J Biomech 1996;29(2):187- 198.
Williams B. Mechanical influences on vascular smooth muscle cell function. J Hypertens 1998;16(12 Pt 2):1921-1929.
Williams C, Wick TM. Endothelial cell-smooth muscle cell co-culture in a perfusion bioreactor system. Ann Biomed Eng 2005;33(7):920-928.
Wilson E, Mai Q, Sudhir K, Weiss RH, Ives HE. Mechanical strain induces growth of vascular smooth muscle cells via autocrine action of PDGF. J Cell Biol 1993;123(3):741-747.
Wilson E, Sudhir K, Ives HE. Mechanical strain of rat vascular smooth muscle cells is sensed by specific extracellular matrix/integrin interactions. J Clin Invest 1995;96(5):2364-2372.
Wilson E. Alpha 7 beta 1 integrin: putting the brakes on smooth muscle cell proliferation. Circ Res 2007;101(7):651-653.
Winder SJ, Walsh MP. Smooth muscle calponin. Inhibition of actomyosin MgATPase and regulation by phosphorylation. J Biol Chem 1990;265(17):10148-10155.
Winder SJ, Walsh MP. Calponin. Curr Top Cell Regul 1996;34:33-61.
266 Wright TC, Castellot JJ, Petitou M, Lormeau JC, Choay J, Karnovsky MJ. Structural Determinants of Heparins Growth Inhibitory Activity - Interdependence of Oligosaccharide Size and Charge. J Biol Chem 1989;264(3):1534-1542.
Yamamoto M, Yamamoto K, Noumura T. Type I collagen promotes modulation of cultured rabbit arterial smooth muscle cells from a contractile to a synthetic phenotype. Exp Cell Res 1993;204(1):121-129.
Yayon A, Klagsbrun M, Esko JD, Leder P, Ornitz DM. Cell surface, heparin-like molecules are required for binding of basic fibroblast growth factor to its high affinity receptor. Cell 1991;64(4):841-848.
Yoshida H, Nakamura M, Makita S, Hiramori K. Paracrine effect of human vascular endothelial cells on human vascular smooth muscle cell proliferation: transmembrane co-culture method. Heart Vessels 1996;11(5):229-233.
Yoshida T, Hoofnagle MH, Owens GK. Myocardin and Prx1 contribute to angiotensin II- induced expression of smooth muscle alpha-actin. Circ Res 2004;94(8):1075-1082.
Yoshida T, Kawai-Kowase K, Owens GK. Forced expression of myocardin is not sufficient for induction of smooth muscle differentiation in multipotential embryonic cells. Arterioscler Thromb Vasc Biol 2004;24(9):1596-1601.
Yoshida T, Gan Q, Shang Y, Owens GK. Platelet-Derived Growth Factor-BB Represses Smooth Muscle Cell Marker Genes via Changes in Binding of MKL Factors and Histone Deacetylases to their Promoters. Am J Physiol Cell Physiol 2006.
Yoshida T, Gan Q, Owens GK. Kruppel-like factor 4, Elk-1, and histone deacetylases cooperatively suppress smooth muscle cell differentiation markers in response to oxidized phospholipids. Am J Physiol Cell Physiol 2008;295(5):C1175-1182.
Yoshikawa H, Taniguchi SI, Yamamura H, Mori S, Sugimoto M, Miyado K, Nakamura K, Nakao K, Katsuki M, Shibata N, Takahashi K. Mice lacking smooth muscle calponin display increased bone formation that is associated with enhancement of bone morphogenetic protein responses. Genes Cells 1998;3(10):685-695.
Yue TL, Wang X, Sung CP, Olson B, McKenna PJ, Gu JL, Feuerstein GZ. Interleukin-8. A mitogen and chemoattractant for vascular smooth muscle cells. Circ Res 1994;75(1):1-7.
Zavan B, Vindigni V, Lepidi S, Iacopetti I, Avruscio G, Abatangelo G, Cortivo R. Neoarteries grown in vivo using a tissue-engineered hyaluronan-based scaffold. Faseb Journal 2008;22(8):2853-2861.
Zhang JC, Kim S, Helmke BP, Yu WW, Du KL, Lu MM, Strobeck M, Yu Q, Parmacek MS. Analysis of SM22alpha-deficient mice reveals unanticipated insights into smooth muscle cell differentiation and function. Mol Cell Biol 2001;21(4):1336-1344.
267 Zhang F, Sun AS, Yu LM, Wu Q, Gong QH. Effects of isorhynchophylline on angiotensin II-induced proliferation in rat vascular smooth muscle cells. J Pharm Pharmacol 2008;60(12):1673-1678.
Zhao Y, Xiao W, Templeton DM. Suppression of mitogen-activated protein kinase phosphatase-1 (MKP-1) by heparin in vascular smooth muscle cells. Biochem Pharmacol 2003;66(5):769-776.
Zheng B, Duan C, Clemmons DR. The effect of extracellular matrix proteins on porcine smooth muscle cell insulin-like growth factor (IGF) binding protein-5 synthesis and responsiveness to IGF-I. J Biol Chem 1998;273(15):8994-9000.
Zhou J, Hu G, Herring BP. Smooth muscle-specific genes are differentially sensitive to inhibition by Elk-1. Mol Cell Biol 2005;25(22):9874-9885.
Zhu JM, Beamish JA, Tang C, Kottke-Marchant K, Marchant RE. Extracellular matrix- like cell-adhesive hydrogels from RGD-containing poly(ethylene glycol) diacrylate. MACROMOLECULES 2006;39(4):1305-1307.
Zilla P, Bezuidenhout D, Human P. Prosthetic vascular grafts: wrong models, wrong questions and no healing. Biomaterials 2007;28(34):5009-5027.
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