Ethylene Glycol) Hydrogels to Regulate

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ENGINEERING POLY (ETHYLENE GLYCOL) HYDROGELS TO REGULATE

SMOOTH MUSCLE CELL MIGRATION AND PROLIFERATION

LIN LIN

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Thesis Advisors: Dr. Roger E. Marchant and Dr. Kandice Kottke-Marchant

Department of Biomedical Engineering

CASE WESTERN RESERVE UNIVERSITY

August, 2014

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Lin Lin

candidate for the degree of Doctor of Philosophy*.

Committee Chair

Kandice Kottke-Marchant

Committee Member

Anirban Sen Gupta

Committee Member

Horst von Recum

Committee Member

Stuart Rowan

Date of Defense

May 6, 2014

*We also certify that written approval has been obtained

for any proprietary material contained therein

This work is dedicated to

my dear husband and my beloved family

for their endless love and support in all of my endeavors.

Table of Contents

Table of Contents ...... i List of Figures ...... iv Acknowledgements ...... v Abstract ...... viii CHAPTER 1: Cardiovascular Disease and Treatment Approaches ...... 1 1.1 Significance of Cardiovascular Disease ...... 1 1.2 Pathogenesis of Cardiovascular Disease ...... 1 1.3 Treatment Approaches for Cardiovascular Disease ...... 3 1.4 Failure Mechanisms of Bypass Grafts ...... 8 1.5 Conclusions ...... 12 1.6 References ...... 13 CHAPTER 2 Regulation of Smooth Muscle Cell Migration and Proliferation...... 20 2.1 Introduction ...... 20 2.2 Methods for Measuring SMC Migration and Proliferation ...... 21 2.3 Basic Mechanisms of SMC Migration and Proliferation ...... 28 2.4 Mediators of SMC Migration and Proliferation ...... 30 2.5 Conclusions and Further Directions ...... 38 2.6 References ...... 39 CHAPTER 3 Hydrogels as Extracellular Matrix Mimics for Three Dimensional Cellular Studies ...... 53 3.1 Introduction ...... 53 3.2 Naturally Derived Materials ...... 54 3.3 Synthetic Poly (ethylene glycol) Hydrogels ...... 56 3.4 Studies of Smooth Muscle Cell Migration and Proliferation in 3D Scaffolds ... 60 3.5 Specific Aims ...... 62 3.6 References ...... 66 CHAPTER 4 Engineer a Cell-adhesive Biodegradable Hydrogel for 3D Cellular Studies ...... 73 4.1 Introduction ...... 73 4.2 Materials and Methods...... 75

4.3 Results ...... 82 4.4 Discussion ...... 90 4.5 Conclusion ...... 93 4.6 Acknowledgement ...... 94 4.7 References ...... 95 CHAPTER 5 Smooth Muscle Cell Migration in 3D Bioactive PEG Hydrogels ...... 99 5.1 Introduction ...... 99 5.2 Materials and Methods...... 100 5.3 Results ...... 104 5.4 Discussion ...... 111 5.5 Conclusion ...... 114 5.6 Acknowledgement ...... 115 5.7 References ...... 116 CHAPTER 6 Smooth Muscle Cell Proliferation in 3D Bioactive PEG Hydrogels ...... 119 6.1 Introduction ...... 119 6.2 Materials and Methods...... 121 6.3 Results ...... 125 6.4 Discussion ...... 133 6.5 Conclusion ...... 138 6.6 Acknowledgement ...... 139 6.7 References ...... 140 CHAPTER 7 Conclusions and Future Directions ...... 144 7.1 Summary and Conclusion of Completed Work ...... 144 7.2 Exploring SMC Functions in 3D Scaffolds ...... 146 7.3 Engineering of Improved Scaffold Systems ...... 148 7.4 In vivo Animal Studies ...... 150 7.5 References ...... 152 Bibliography ...... 156

List of Tables

Table 2. 1 ...... 35

Table 3. 1 ...... 64

Table 3. 2 ...... 65

Table 4. 1 ...... 76

Table 4. 2 ...... 86

iii

List of Figures

Figure 4. 1 ...... 78

Figure 4. 2...... 83

Figure 4. 3...... 85

Figure 4. 4...... 88

Figure 4. 5...... 89

Figure 5. 1 ...... 105

Figure 5. 2...... 106

Figure 5. 3...... 107

Figure 5. 4...... 109

Figure 5. 5 ...... 110

Figure 6. 1...... 127

Figure 6. 2...... 128

Figure 6. 3...... 130

Figure 6. 4...... 131

Figure 6. 5...... 132

Figure 6. 6...... 134

Figure 6. 7...... 135

Acknowledgements

It would not have been possible to complete this doctoral work without the help and support of a great many people.

Foremost, I would like to express my deepest gratitude to my advisors, Dr. Roger

Marchant and Dr. Kandice Kottke-Marchant. They have been outstanding advisors and mentors for my PhD study and research. I appreciate very much for the excellent guidance, advice, and support of Dr. Roger Marchant. He was a great advisor and served as an excellent role model to me. He patiently provided not only the vision, knowledge and suggestions necessary for me to develop this PhD project, but also generous encouragement and opportunity to me to pursue independent work. I felt motivated and encouraged every time I had meeting or discussion with him. I am so sad of his passing and the time spent with Dr. Roger Marchant will be dearly missed. I am also extremely grateful to Dr. Kandice Kottke-Marchant for the continuous support of my PhD study and research, even at her most hard time. The great guidance, advice and patience of Dr.

Kandice Kottke-Marchant throughout the research project, as well as her painstaking effort in proof reading the drafts, are deeply appreciated. Without her supervision and constant support, this thesis would not have been completed.

I am also deeply thankful for my experiences as a graduate student in Dr.

Wenguang Liu's lab at Tianjin University in China. My master’s advisor, Dr. Wenguang

Liu, inspired my research interest in the areas of biomaterials. Without his guidance, encouragement and help, I would not be able to pursue my PhD research at Case. Besides my advisors, I would also like to thank the rest of my thesis committees, Dr. Anirban Sen

Gupta, Dr. Stuart Rowan, and Dr. Horst von Recum. They have generously provided their

insightful advice, knowledge, and comments to help me with my project and presentation skills. I would particularly like to thank Dr. Anirban Sen Gupta, who was willing to be my academic advisor at the last year, and Dr. Horst von Recum, who was willing to participate in my final defense committee at the last moment.

My sincere thanks also goes to the past and present members of the Marchant group. I appreciate Dr. Junmin Zhu very much for his numerous helpful advices on my research project and manuscript revision. I am deeply grateful for his kind help when I first came to America and his caring and help after he left the group. I would also like to thank Dr. Chris Hofmann for his guidance on using and maintaining a wide range of equipment, Dr. Lynn Dudash for her assistance with lab maintenance and guidance on cell culture, Dr. Jeffrey Beamish for teaching me hydrogel synthesis and characterization,

Dr. Ping He for teaching me gel electrophoresis experiment. They have been always generously and kindly providing helpful suggestions and advice every time I had inquiries with experimental procedures, lab equipment, and supplies. I am particularly thankful for their continuous caring and help after they left the group. The current

Marchant group members, Dr. Faina Kligman, Jennifer Bastijanic, Derek Jones, Han Xu, have been a daily source of support and encouragement. I would like to thank Dr. Faina

Kligman and Jennifer Bastijanic for their assistance with peptide characterization and lab maintenance, Derek Jones for his help with lab supplies ordering and equipment maintenance. My research work would not have been possible without their support and assistance to keep the lab organized.

I also appreciate the assistance of Maryanne Pendergast in Neurosciences Imaging

Center for teaching me to use confocal microscope, Dale Ray in the NMR lab for training

me to use NMR, Annette Marsolais in the SCSAM center for teaching me SEM.

I would also like to thank the support from the Department of Biomedical

Engineering. I would particularly like to thank Angie Bracanovic for her assistance of

processing my orders, graduate support forms, and other forms through the BME

bureaucracy. I am also thankful to the National Institutes of Health for the financial

support for my studies and research project.

Most importantly of all, I would like to thank my family for their endless love and

support in all of my endeavors. I would like to thank my parents, who have been always supporting me and encouraging me with their best wishes. I am also thankful to my husband, Yaoying Wu, who has always been there cheering me up every time I was feeling down through this process and supporting me whenever I need help. I will be grateful forever for their love.

vii

Engineering Poly (ethylene glycol) Hydrogels to

Regulate Smooth Muscle Cell Migration and Proliferation

Abstract

LIN LIN

The key role of smooth muscle cell (SMC) migration and proliferation in vascular

physiological and pathological remodeling necessitates the exploration of mechanisms

underlying these functions. This work focuses on engineering a poly (ethylene glycol)

(PEG) hydrogel as a model system to evaluate SMC migration and proliferation in three dimensions (3D). We hypothesized that 3D SMC migration and proliferation can be regulated by the properties of a cell-instructive scaffold, including cell-matrix adhesion, degradability, and cross-linking density. To accomplish this, bio-inert PEG-based hydrogels were designed as the scaffold substrate. To mimic the properties of the extracellular matrix (ECM), cell-adhesive peptide (GRGDSP) and enzyme-sensitive peptide (VPMSMRGG or GPQGIAGQ) were incorporated into the PEG macromer chain. Copolymerization of the biomimetic macromers by photopolymerization resulted in the formation of bioactive hydrogels with the dual properties of cell adhesion and proteolytic degradation. Studies of mass swelling ratio as a function of gel compositions indicated that this hydrogel can be engineered quantitatively to allow for uncoupled

viii

investigation of scaffold properties on cell functions. By utilizing these biomimetic

scaffolds, we studied the effect of adhesive ligand concentration, proteolysis, and

network cross-linking density on 3D SMC migration and proliferation. Our results indicated that 3D SMC migration and proliferation were critically dependent on cell- matrix adhesiveness, proteolysis, and cross-linking density. The incorporation of cell- adhesive ligand significantly enhanced SMC spreading, migration and proliferation, with cell-adhesive ligand concentration mediating 3D SMC migration and proliferation in a biphasic manner. The faster degrading hydrogels promoted SMC migration and proliferation. In particular, higher cross-linking density could impede 3D SMC migration and proliferation despite the presence of cell-adhesive ligands and proteolytically degradable sites. Furthermore, the exogenous factor, heparin, exerted significant inhibitory effect on 3D SMC proliferation. These cell-instructive constructs serve as a good model system to study the effect of hydrogel properties on 3D SMC functions and show promise as a tissue engineering platform for vascular in vivo applications.

CHAPTER 1: Cardiovascular Disease and Treatment Approaches

1.1 Significance of Cardiovascular Disease

Cardiovascular disease (CVD), including coronary artery disease (CAD), stroke, heart failure, and peripheral arterial disease (PAD), is the leading cause of mortality in the United States [1] . An estimated 83.6 million American adults (>1 in 3) live with one or more types of CVD, accounting for 31.9% of all deaths on the basis of 2010 mortality data [1]. This represents about 1 of every 3 deaths in the United States [1]. In addition to affecting so many individuals’ health in the United States, CVD also creates a huge financial burden on the health care system. In 2009, CVD resulted in a total inpatient hospital cost of $71.2 billion for one sixth of hospital stays [1]. The combined direct and indirect cost of CVD for 2010 is estimated to be 315.4 billion, and the total direct cost of

CVD is projected to increase to 918 billion [1]. Therefore, there is a critical need to enhance the management and treatment of CVD in the United States.

1.2 Pathogenesis of Cardiovascular Disease

CVD can refer to many different types of diseases that affect the cardiovascular system, many of which are related to a process called atherosclerosis, a buildup of atheromatous plaque in the walls of arteries [2, 3]. In order to better appreciate the underlying pathology leading to atherosclerosis, the important structural and functional characteristics of native vessels will be described first.

1.2.1 Normal Blood Vessel Histology

Normal arteries possess three layers: the tunica intima, the tunica media, and the tunica adventitia [3-5]. The innermost layer, called intima, normally consists of endothelial cells (ECs) and their basement membrane. ECs normally maintain a non-

thrombogenic blood-vessel interface, regulate vessel permeability, modulate homeostasis, and affect the growth of other cell types, particularly vascular smooth muscle cells

(SMCs) [3]. The tunica intima is separated from the tunica media by a thin layer of internal elastic lamina (IEL). Within the layer of tunica media, SMCs align circumferentially around the vessel and are surrounded by the interwoven collagen fibers and elastic lamina [4, 5]. The media contributes the majority of the vessel’s mechanical functionality. As the predominant cellular element of the vascular media, SMCs are responsible for the dilation and constriction of the vessel that occurs in response to the external stimuli [4, 5]. The tunica adventitia consists of fibroblasts, connective tissue, the microvascular supply for the blood vessel itself, and a neural network that modulates the vasotone of the blood vessel [5].

1.2.2 Atherosclerosis

Atherosclerosis is a chronic, inflammatory disease in response to endothelial dysfunction [2, 3]. Possible factors contributing to endothelial dysfunction in early atherosclerosis include elevated and modified low density lipoprotein (LDL), hypertension, toxins from cigarette smoke, homodynamic disturbances, elevated plasma homocysteine concentrations, and combinations of these or other factors [2, 3].

After endothelial injury, the permeability of endothelium is increased and more

LDL cholesterol and other lipids accumulate in the intima, causing further injury and increased adhesiveness of dysfunctional endothelium with respect to the inflammatory cells (e.g. monocytes) and platelets [2, 6, 7]. Monocytes emigrate into the intima, become activated, and transform into macrophages and foam cells [2, 8]. Different cell types, including ECs, inflammatory cells, and activated platelets, release mediators, such as

growth factors and cytokines [7, 8]. These regulatory factors stimulate migration and

proliferation of SMCs from media to intima. The expression of matrix metalloproteinases

(MMPs) is also upregulated after intimal injury. MMPs could catalyze and remove the basement membrane around SMCs, which facilitates SMC migration and proliferation [7,

8]. These processes are accompanied with the deposition of a collagen I-rich matrix secreted from SMCs, which forms a fibrous plaque above the cholesterol filled core [2,

3]. With progression, the advanced plaques can weaken the surrounding vessel wall and inhibit the blood flow, leading to the critical stenosis of the vessel. These plaques can also rupture and erode, which exposes the blood to highly thrombogenic substances and induces acute thrombosis [2]. Such thrombosis can result in vessel occlusion, leading to downstream ischemia and infarction.

1.3 Treatment Approaches for Cardiovascular Disease

1.3.1 Healthy Lifestyle Modifications

Adoption of healthy lifestyle, including cessation of smoking, a balanced diet,

regular exercise, and weight control, plays a key role in reducing risk for CVD [9, 10].

Cigarette smoking is a major risk factor for CVD. As many as 30% of all CAD deaths in the United States each year is attributed to smoking [11]. After cessation of smoking, former smokers had a 30% reduction of non-fatal myocardial infarctions and overall mortality compared with continuing smokers [12]. Maintaining a healthy diet can improve cardiovascular risk factors, such as blood pressure (BP) and LDL cholesterol levels [9, 10, 13]. The American Heart Association (AHA) recommends that individuals consume a variety of fruits, vegetables, grains (especially whole grains), fish, and lean meats, as well as limit the intake of sodium, alcohol, dietary cholesterol and trans-fatty

acid [9, 10]. Regular physical activity is essential for physical fitness and cardiovascular

fitness, which resulted in a 20% reduction in all-cause mortality and a 26% reduction in total cardiac mortality [13]. At least 30 minutes of moderate-intensity physical activity on most days of the week is recommended by the AHA [9]. Obesity is related to increased cardiovascular risk. Reducing caloric intake combined with regular exercise can facilitate the weight loss [10].

1.3.2 Medical Management

For individuals with high cardiovascular risk, such as high blood pressure and

high LDL cholesterol levels, medical intervention may be considered in addition to

lifestyle modifications. A normal BP is a systolic BP < 120 mm Hg and a diastolic BP <

80mm Hg. Antihypertensive medications should be initiated in patients with BP ≥ 140

mm Hg if lifestyle modifications are not effective [9]. Diabetes mellitus is a key risk factor for adverse cardiovascular outcomes. Appropriate hypoglycemic therapy may reduce the risk of cardiovascular disease [9, 10]. Statins are 3-hydroxy-3-methyl glutaryl

coenzyme A (HMG-CoA) reductase inhibitors indicated for reduction of LDL cholesterol

levels. Statin therapy has been shown to significantly reduce CVD mortality in primary and secondary prevention of CVD [9, 13, 14].

1.3.3 Angioplasty and Stent Placement

When medical intervention fails to achieve desired results, more invasive therapy

is required. Percutaneous coronary intervention (PCI), commonly known as angioplasty,

is a non-surgical therapy to open narrowed or blocked blood vessels to restore adequate

blood flow. The procedure of PCI involves inserting a balloon-tipped catheter into the

stenotic artery and inflating the balloon in the blocked region to widen the diameter of

vessel [15]. A stent is often used in company with angioplasty to provide scaffolding to

hold the artery open [15]. Angioplasty is a relatively low-risk and low-cost procedure.

Nearly one million patients have angioplasty procedures each year in the United States

[13, 15].

The major challenge of angioplasty is restenosis, or reclosure of the artery [13,

16]. A 40% chance of restenosis is associated with angioplasty procedure alone.

Angioplasty with stenting reduces the risk of restenosis to 25% [16-18]. Restenosis that

occurred after the use of stent is referred to “in-stent restenosis” [16, 19]. The

development of restenosis can be attributed to the pathogenic changes induced by the

damage to the vessel wall by angioplasty. The disrupted vessel layers stimulate

inflammatory immune response, migration and proliferation of SMCs in the intima, and

tissue accumulation [19]. The introduction of drug-eluting stents has improved the results

of PCI by decreasing the incidence of in-stent restenosis [16, 20].

1.3.4 Bypass Grafting

Coronary artery bypass graft (CABG) surgery is recommended for patients with

multiple areas of coronary artery narrowing or blockage [13, 15]. In bypass grafting, a

non-diseased, autologous vessel is connected, or grafted to the blocked coronary artery,

creating a new path for the blood flow [15]. Compared with PCI, CABG is a more

invasive surgical procedure with greater risks and costs [15]. CABG has been indicated to be associated with higher risk for procedure-related stroke (1.2% versus 0.6% with PCI)

[21]. However, patients who received CABG had a lower frequency of angina and a need for fewer repeat revascularization procedures than did patients who received PCI [21].

The risk rates for repeated revascularization at 5 years were 46.1% for PCI, 40.1% for

PCI employing stents, and 9.8% for CABG [21]. In comparison with PCI using drug

eluting stents, CABG continued to be associated with lower mortality rates, lower rates of

death or myocardial infarction and repeated revascularization for patients with multi-

vessel disease [22, 23].

The used of autologous vessels, including the saphenous vein and the internal

mammary artery, remains a preferred option as bypass grafts for CABG surgery [15]. The

saphenous veins display 85%-90% patency rates at 1 year after surgery, decreasing to

60%-70% at 10 year after surgery [24]. Better long-term patency has been associated

with the internal mammary artery [25]. In 2010, 397,000 coronary artery bypass

procedures were performed in 219,000 patients in the United States [1]. This indicates

that the majority of patients may require two or more grafts. Unfortunately, 20-30% of

patients do not have suitable autologous vessels for graft surgery [26, 27]. The reasons

may include poor quality due to patient’s vessel disease, inappropriate size, and

exhaustion due to previous surgical harvest [28, 29]. Furthermore, the harvest of autologous vessels adds time, cost and the risk of additional morbidity to the surgical

procedure. For all of these reasons, suitable alternative conduits to the autologous vessels

are desirable.

Currently, poly (ethylene tetrephthalate) (PET) (ie, Dacon), expanded

polytetrafluoroethylene (ePTFE) (ie, Goretex) are most commonly used materials to

fabricate synthetic vascular grafts [4, 30]. These materials are readily available, relatively

inexpensive, and have been shown to perform excellent in applications with vessel

diameters > 6mm [5]. ePTFE for aorto-bifemoral bifurcation grafts has about 95%

patency rates at 5 year after surgery, similar to Dacron [31, 32]. However, these synthetic

prostheses are limited to high flow and low resistance applications due to low elasticity, poor compliance and thrombogenicity of synthetic surfaces [33]. A mismatch in mechanical properties (e.g. elasticity and compliance) between grafts and native vessels causes shear rate disturbances and disturbed hemodynamic flow that contribute to graft failure [34, 35]. The compliance of polyurethanes (PUs) is much closer to that of native blood vessels, therefore this synthetic material has been introduced to produce bypass grafts, which has greatly reduced the problems associated with mismatched mechanical properties [36]. However, none of these materials has exhibited satisfactory long-term patency in small diameter applications (<4 mm) and are not available clinically for coronary artery bypass [5].

Tissue engineered blood vessels (TEBVs) as alternative bypass conduits have attracted extensive research interest [5, 30, 33, 37]. Tissue engineering approaches to develop vascular grafts are varied, but generally involve combining scaffolds (e.g. synthetic polymer-based scaffolds, decellularized contructs) with biological components

(e.g. cells, biological factors) to recapitulate the structure and function of native vessels

[5, 38]. An ideal tissue engineered blood vessel should be biocompatible (non-toxic and immunocompatible) and should promote cell attachment, cell invasion and remodeling of the scaffold to possess sufficient mechanical properties to perform its function upon implantation [38]. The tissue engineered blood vessel must also serve as a bridge to regulate vessel wall cell functions to prevent graft failure after implantation [38]. In addition, the vascular graft should be readily available and reasonably economical to

produce and store. Much effort and progress have been made in improvement of TEBV

functions, but a suitable, clinically available TEBV has yet to be developed [39].

1.4 Failure Mechanisms of Bypass Grafts

Failure of bypass grafts may be temporally divided into early and late categories.

The early graft failure is commonly attributable to infection, thrombosis, and/ or technical surgical errors [27]. Infection occurs in 1-6% of arterial vascular grafts [40]. The principal underlying mechanism of early graft failure is thrombosis [25, 36, 41]. Between

3-12% of saphenous vein grafts are occluded by thrombosis within 1 month after graft surgery [41]. For synthetic vascular grafts, the follow up study of 7 mm femoralpopliteal ePTFE bypass grafts has shown that 5% were occluded by thrombosis after 1 month, with rates for smaller diameter ePTFE grafts even higher [42]. Intimal hyperplasia is the most important cause of late failure, accounting for more than 20% of late failures of infrainguinal prosthetic graft revascularizations [43]. The development of intimal hyperplasia can lead to significant stenosis of the lumen as well as the development of atherosclerosis and thrombosis [44].

1.4.1 Thrombosis Challenge

Acute thrombosis is the major contributor to early graft failure, especially in small diameter synthetic vascular grafts, leading to decreased flow or occlusion [4]. The causes of thrombosis may involve endothelial injury and surgical technical errors [25]. ECs normally provide a non-thrombogenic blood-vessel interface [3-5]. Damage to endothelium after surgery causes endothelial dysfunction, leading to upregulation of procoagulants (e.g., von Willebrand factor, thrombin, tissue factor, platelet activating factor) and downregulation of antithrombotics (e.g.: nitric oxide, thrombomodulin) [25,

45]. This leads to platelet adhesion, activation and aggregation. Subsequent activation of the clotting cascade eventually causes thrombosis [25, 45]. The technical errors that

increase the risk of thrombosis include exposure to the high and unaccustomed pressures to detect leaks, mismatched sizes of graft that alters flow dynamics, and excessive or insufficient graft length that results in kinks and linear tension in the graft [25, 35].

Antithrombotic drug therapy along with bypass grafting, including the application of anticoagulants such as heparin, and platelet inhibitors such as aspirin and clopidogrel, has been shown to be beneficial in decreasing graft occlusion and prolonging graft patency after surgery [4, 46-50]. Besides systemic drug therapy, drug coating or binding drugs to grafts to reduce thrombogenicity have emerged as an effective approach for improving graft patency [34]. For example, heparin-bonded ePTFE grafts have shown to provide promising early patency (82% and 97% for the overall primary and secondary 1- year patency rate) after bypass surgery [51]. Several additional graftings have undergone preliminary testing, such as coating ePTFE grafts using Hirudin, a direct thrombin antagonist, in combination with iloprost, an inhibitor of platelet aggregation [52]. The in vivo testing of grafts in a pig model found that Hirudin and iloprost modified ePTFE groups maintained blood flow rates at 6 weeks compared with baseline, while control groups (untreated ePTFE grafts) had markedly reduced flow [34, 52]. Due to the key role of endothelial cells in preventing thrombosis in native tissue, various strategies aspire to induce endothelialization of graft surface either prior to implantation or by accelerating in situ endothelialization [30, 53-56].

1.4.2 Intimal Hyperplasia Challenge

Intimal hyperplasia (IH), particularly pronounced in the area of distal anastomoses, is a major disease process in vascular grafts between 1 month and 3 year after implantation, representing the foundation for later development of atherosclerosis

[25, 45]. In bypass grafting, IH can occur in all types of vascular grafts, including vein, artery and prosthetic grafts. The risk of graft failure due to IH is synthetic vascular graft> vein > artery [25, 41]. The study of vein grafts as bypass conduits has shown that almost all veins implanted into the arterial circulation develop further intimal thickening within 4 to 6 weeks, which may reduce the lumen by up to 25% [41], while only 4.2% of internal mammary artery bypass grafts had more than a 25% decrease in lumen diameter [25, 57].

Besides in response to bypass surgery, intimal hyperplasia occurs in a number of other pathological situations. For example, IH can occur in response to endothelial injury due to angioplasty, leading to the development of restenosis or in-stent restenosis in the vessels or stents [16, 19, 44]. In response to endothelial damage in native vessels due to surgical procedures (e.g, autologous bypass grafting), growth factors, such as platelet-

derived growth factor (PDGF) and basic fibroblast growth factor (bFGF) from platelets and injured endothelial cells, stimulate smooth muscle cells in the media switching from a quiescent, non-proliferative state to a synthetic, proliferative state [25, 45, 58, 59]. The phenotypic switch up-regulates the expression of matrix metalloproteinase, facilitates migration and proliferation of smooth muscle cells. This is required for wound healing and initial vascular repair. However, the over stimulation induces excessive migration and proliferation of SMCs from media to intima as well as excessive ECM deposition in the intima, which leads to the development of intimal hyperplasia [3, 25].

The development of intimal hyperplasia in synthetic vascular grafts is usually found at the distal anastomosis of vascular grafts [60, 61]. After graft surgery, the vessel injury at the artery-graft anastomosis cannot be avoided due to the necessity of suturing.

The vessel injury, along with a mismatch in mechanical properties of native arteries and

grafts, induce the development of distal anastomosis of intimal hyperplasia (DAIH) in vascular grafts, which distributes along the suture line [60-62]. Similar to IH induced by other stimuli, the lesions are developed by extensive SMC migration, proliferation and

ECM synthesis [63].

Statin, 3-hydroxymethyl-3-methylglutaryl coenzyme A (HMG-CoA), has been used extensively for treatment of hyperlipidemias by inhibiting a key enzyme in the pathway of cholesterol synthesis [64]. In addition to lipid lowering, statins also have been reported to have pleiotropic activities of inhibiting SMC migration, proliferation, and

ECM synthesis in both in vitro studies and in vivo animal model studies [65-67]. It has been reported that the pleiotropic properties of statin are related to inhibition of mevalonate synthesis and the isoprenylation reactions important in signal transduction by small G proteins, which are key regulatory proteins participate in SMC migration [68].

Other possible mechanism underlying the inhibitory effect of statin is their inhibition on matrix metalloproteinase secretion, or interruption of growth factor (e.g., PDGF) stimulated cell migration and proliferation [69, 70]. The many benefits of statin mediated vascular effects have established the key role of these drugs in the primary and secondary prevention of CVD [65-67]. Growth factors like bFGF have been shown to promote

SMC migration and proliferation [58, 59]. The introduction of antibodies to bFGF has been reported to reduce SMC proliferation in ePTFE grafts by in vivo testing of animal models [71]. Thus, regulation of SMC migration and proliferation after implantation is important for preventing vascular wall pathogenic remodeling [72, 73].

1.5 Conclusions

Cardiovascular disease continues to be the leading cause of mortality in the

United States. CVD can refer to many different types of diseases, including CAD, PAD,

and stroke, many of which are related to a process called atherosclerosis, which occurs

over time to endothelial injury, involves accumulation of lipids and migration and

proliferation of SMCs from media into intima. Revascularization procedures including

angioplasty and bypass grafting are mainstay treatments for advanced CVD. These

treatments can restore adequate blood flow effectively, but their limited ability of

inhibiting vascular pathogenic remodeling results in the re-narrowing or re-occlusion of the vessels, especially in small diameter blood vessels. As the major cellular components of the blood vessels, endothelial cells and smooth muscle cells play important roles in vascular biology and pathology. To develop effective treatment approaches for CVD, understanding the mechanisms that regulate vessel wall cell functions has been a major focus of research.

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CHAPTER 2 Regulation of Smooth Muscle Cell Migration and Proliferation

2.1 Introduction

Smooth muscle cell (SMC) migration and proliferation occurs under a number of

conditions: during blood vessel formation, in response to vascular injury, and during

development of atherosclerosis [1-8]. During vessel development, endothelial cells (ECs)

form the inner layer of the vessel wall. Growth factors (e.g., platelet-derived growth

factor (PDGF)) released from ECs stimulate the recruitment of pericytes or smooth

muscle progenitor cells that envelop the surface of the endothelial cell tube [1, 2].

Following migration, pericytes or smooth muscle progenitor cells proliferate and

differentiate to a smooth muscle cell layer in the media that is responsible for the

mechanical functionality of blood vessels [1, 9, 10]. It has been reported that mice

lacking PDGF-B or PDGFβ receptors die during late gestation from microvascular

dysfunction [2, 11, 12]. The cause of microvascular dysfunction is severe pericyte/SMC

deficiency on the affected vessels, which is correlated with the failure of pericyte/SMC

migration and proliferation when PDGF-B/PDGFβ signaling is disrupted [2, 13].

Migration and proliferation of SMCs plays a key role in the development of

intimal hyperplasia and other pathologies, such as restenosis and atherosclerosis [3-8]. In

the normal unjured vessels, contractile SMCs that are firmly adherent to the surrounding

matrix, are nonmigratory and not proliferating [7]. After vascular surgery, such as

angioplasty, stent implantation, and bypass grafting, damage to the vessel wall results in

the upregulation of matrix metalloproteinases (MMPs) and other signaling molecules that favor SMC migration and proliferation [3-6, 14]. These events stimulate SMC migration through internal elastic lamina (IEL) where they proliferate and secrete extracellular

matrix (ECM) [3-6]. Excessive SMC migration and proliferation from media to intima

result in the narrowing or occlusion of the vessels, leading to the further development of

atherosclerosis or acute thrombosis [6, 9, 14].

The key role of SMC migration and proliferation in vessel wall remodeling has

stimulated strong interest in the cell and molecular biology of SMC functions. In this

chapter, the common methods to measure SMC migration and proliferation will be

summarized. The cellular events of SMC migration and proliferation will be briefly

reviewed. Finally, the current knowledge and theories on the role of extracellular signals,

including growth factors, ECM proteins, and physical factors, on SMC functions will be

discussed.

2.2 Methods for Measuring SMC Migration and Proliferation

2.2.1 Methods for Measuring SMC Migration

A variety of in vitro approaches have been used to explore the basic mechanisms

of cell migration as well as the effect of extracellular signals on cell migration. Generally, cell migration can be evaluated by either microscopic methods to record changes in cell position and shape or assessment of migrated cell population by fluorescence or staining

[7]. The cell migration assays for 2 dimensional (2D) and 3 dimensional (3D) culture studies are well reviewed elsewhere [7, 15]. Some common assays, including in vitro wound healing assay, transwell migration and invasion assay, microcarrier bead invasion assay, modified 3D invasion assay, and time-lapse/cell tracking assay, will be summarized briefly here.

2.2.1.1 In vitro Wound Healing Assay, Cell Exclusion Zone Assay and Fence Assay

The wound-healing assay is one of the earliest developed methods to study cell migration on 2D surfaces in vitro [16]. Generally, a “wound” (scratch area) is created in a confluent cell monolayer, which is most easily done by a plastic pipette tip [17, 18]. Cell migration can then be monitored by capturing images as cells migrate from the intact areas into the “wound” gap. The different substrates that cells are grown on or the factors added to the medium that alter the motility of cells can lead to the change of time to restore confluence (close of scratch area) [16]. This assay is simple, inexpensive, and the experimental conditions can be easily modified for studies of the effects of cell-matrix and cell-cell interactions on cell migration. The drawbacks of this assay include the damage of cell from scratching and unevenly thickness of scratch that will affect cell migration rate [15].

To avoid these problems, cell exclusion zone assays or fence assays have been introduced to study cell migration [15]. For the cell exclusion zone assay, small silicone stoppers are positioned prior to cell seeding to create an exclusion zone with the tip of the stopper [15]. After cell adhesion, the stoppers are removed and a cell-free area is created by the tip of the stopper. In contrast, the fence assay is to seed cells into the inner area of a Teflon, glass or metal fence (ring) placed on a substrate and to create a cell-free area outside the fence. After cell attachment, the removal of the fence will allow cell migration from the inner area in a radial way outward [15, 19].

In the above assays, the cells are incubated in a uniform concentration of stimuli.

Since there is no chemoattractant concentration gradient during the experiment, the migration of cells is largely a function of increased chemokinesis [20].

2.2.1.2 Transwell Migration and Invasion Assay

The transwell assay, also known as Boyden chamber assay, is based on two chambers filled with culture medium separated by a microporous membrane through which cells transmigrate [7, 15, 21]. In general, cells are seeded on the top surface of the porous membrane in the upper chamber and can migrate through the pores of the membrane into the lower chamber, in which chemoattractants are present. After an appropriate culture time, the number of cells that migrate to the lower chamber is quantified by nuclei staining. The migrated cells can be stained and visualized by microscopy [22, 23]. They can also be detached from the lower chamber, stained with fluorescent markers, with the DNA amount that reflects the number of cells determined by a fluorescence plate reader [7, 15, 21].

In addition to 2D migration, the transwell assay can be modified to assess cell invasion in 3D [15, 24, 25]. To create a 3D invasion assay, the porous membrane is covered by a layer of matrix gel before cell seeding into the upper chamber [15, 24].

Cells seeded on the gel surface can degrade the gel, migrate through the gel and membrane, and adhere to the bottom of the membrane. After a determined culture period, the non-invaded cells on gel surface can be removed (e.g. by a cotton swab) and the invasive cells on the bottom of the membrane can be determined by microscopy or a fluorescence plate reader as described above [15].

The transwell assay is a popular method to investigate cell migration because of its simple set-up. Compared with wound healing assay, the transwell assay can be utilized to evaluate the chemotactic ability of cells [7]. However, the chemotactic gradients cannot be sustained due to diffusion [15, 20]. Moreover, this is an endpoint assay which

requires termination of the experiment to quantify the population of migrated cells [21,

26].

2.2.1.3 Microcarrier Bead Invasion Assay and Modified 3D Cell Invasion Assay

The microcarrier bead assay evaluates cell motility based on quantification of

invasion distance of the cells from microcarrier beads into surrounding matrix [15, 27].

The procedures of this method involve coating gelatin-coated beads (commercially

available) with cells, encapsulating cell-coated beads into matrix gel, and monitoring cell

outgrowth from the beads into surrounding matrix by microscopy [27]. In contrast to the

transwell invasion assay, this method allows to measurement of migration in kinetic

experiments without termination of the assay [15]. However, this method is time- consuming, since there are multiple steps to prepare the beads with a confluent cell

coating.

Similar to the principle of microcarrier beads assay, a modified 3D cell invasion

assay that uses cell aggregates instead of cell-coated microcarrier bead has been utilized

to investigate cell migration in 3D matrix [28-30]. Cells are suspended in an aggregating

solution (media with methocel [29] or naturally derived materials (e.g. fibrin, matrigel)

[28, 30]) and incubated until the aggregate forms. Then the cell aggregate is embedded in

the matrix and cell outgrowth from the central aggregate into surrounding matrix can be

monitored and quantified by microscopy in a kinetic manner [28-30].

For the above assays, there is a disadvantage that cell proliferation in the

surrounding matrix may contribute to the cell number and migration distance [29, 30].

Techniques to separate migration from proliferation, such as mitomyosin C (MMC) to

inhibit cell proliferation in 3D cell invasion experiment have been described [29].

However, there are concerns that MMC may affect secretion of MMs or inhibitors, and

thereafter, affect cell migration [30, 31].

2.2.1.4 Time-lapse/Cell Tracking

Single cell migration can be analyzed by tracking individual cells with computer-

aided videomicroscopy in time-lapse experiments [15, 32]. In this assay, samples are placed on an inverted optical microscope equipped with a software-controlled motorized stage. Cells are maintained at physiological conditions by a stage incubation system. The randomly chosen locations of the samples are repeatedly imaged at determined time

intervals and the recorded migration paths are analyzed [32]. This method can be applied

to investigate cell migration on 2D surface as well as in 3D matrix [33-35]. For 3D cell

tracking, cells are homogeneously encapsulated in the matrix [33, 34]. The analysis of

cell migration on 2D or cell invasion in 3D is based on a random walk model, which is

well described elsewhere [35-38].

The major advantage of this method is that individual movements of cells can be

monitored in real time. The actual length of the individual cell migration as well as

direction and velocity can be exactly determined. However, a specialized microscope for

live imaging and advanced knowledge in data processing are required for this method

[15].

2.2.2 Methods for Measuring SMC Proliferation

Cell proliferation can be evaluated by various assays. The number of cells can be

determined directly by the use of hemocytometer, Coulter counter or flow cytometer [39,

40]. Since the amount of DNA in each cell remains constant (except S phase), the number

of cells can also be measured by quantification of DNA content in the samples [39-41].

Alternatively, the relative metabolic activity can be quantified to reflect the number of

cells because the relative enzymatic activity is consistent among cells [39, 40, 42]. Each

method has its own advantages and limitations. The choice of which assay to use will

depend on a number of factors including the equipment available, the experimental

design, and the questions being addressed.

2.2.2.1 Cell Counting

Cell counting using a hemocytometer is the simplest method to quantify the cell

numbers. This method can be combined with trypan blue dye exclusion to determine the

percentage of viable cells of a cell population [39, 43]. Generally, the cells are harvested

from the surface of substrates using trypsin-EDTA (ethylene diamine tetraacetic acid).

The cell suspension will be subsequently mixed with trypan blue dye solution, loaded to a

hemocytometer, and examined under a microscope [39]. The blue staining cells are

considered non-viable since the intact membrane of live cells will prevent trypan blue

entering cells. The percentage of live cells in the cell suspension can be calculated by

dividing the number of live cell to the total number of cells. A cell counter or flow cytometer, if available, can be utilized to count cells automatically; these techniques are less time consuming and more accurate [44].

2.2.2.2 DNA Quantification

DNA amount can be quantified in different ways depending on the equipment

available and the question being addressed. One method for measuring DNA synthesis is to incorporate [3H]-thymidine into proliferating cells during S phase of the cell cycle [45,

46]. The amount of incorporated radioactivity can be quantified by scintillation counting.

The advantage of this method is that this measurement directly quantifies the amount of

newly synthesized DNA caused by added stimuli [40]. However, some studies have

suggested that this radiochemical induce cell cycle arrest and apoptosis [47].

Another method to assess DNA content is to measure fluorescence by flow cytometry or spectroscopy after DNA staining [41, 48]. Propidium iodide (PI) is a fluorescent dye that binds to double-stranded DNA. Generally, following treatment, cells are fixed, permeabilized, and stained with PI. Through the use of flow cytometer, the amount of DNA per cell can be determined based on the amount of fluorescence per cell.

This analysis can indicate the effect of treatment on the cell cycle progression since the

DNA content per cell reflects the stage of the cell cycle [40, 48]. The total DNA content in the sample can also be quantified by fluorescence plate reader (e.g., PicoGreen assay,

Life Technologies) [39, 41]. After cell lysis, the fluorescent dye selectively binds to the double-stranded DNA and the fluorescence can be subsequently measured by the plate reader. This method does not distinguish between quiescent cell and actively dividing cells [40]. However, by comparing the DNA content after treatment with the content before treatment, the effect of treatment on cell proliferation can be analyzed [49, 50].

2.2.2.3 Metabolic Measurement

Cell proliferation can be detected by the use of metabolic dyes, such as 3-(4 5-

dimethylthiazol-2)-2, 5-diphenyltetrazolium bromide (MTT) [42]. In the presence of

MTT, the NAD(P)H-dependent cellular oxidoreductase enzymes in the live cells can

reduce tetrazolium salt to a formazan product, which can be detected by the resulting

colorimetric change [40]. It is of note that phenol red in cell culture media, fatty acids and

serum albumin have been reported to reduce the sensitivity of this assay [51].

Furthermore, when this assay is utilized to assess cell proliferation in 3D matrix, the

diffusive ability of MTT reagent or resulting formazan product through the matrix might

affect the accuracy of the assay.

2.3 Basic Mechanisms of SMC Migration and Proliferation

2.3.1 Basic Mechanisms of SMC Migration

Cell migration in vivo or in vitro begins with a protrusion of the plasma

membrane-leading lamellae (leading edge) that is in contact with extracellular substrate

[14, 52]. The protrusions are caused by the cytoskeletal actin polymerization and are stabilized through the formation of new focal contacts (focal complexes) just behind the leading edge [53, 54]. These adhesive complexes are formed from the binding of actin cytoskeleton to the underlying ECM by integrin transmembrane receptors [52, 55]. As the cells migrate, focal complexes mature into larger, more organized focal adhesions that secure the adhesion of the cell membrane to the matrix at the front edge [54, 56]. Traction forces are then generated from actomyosin (a complex of myosin and actin filaments) contraction, which promotes cytoskeletal remodeling and detachment of focal contacts at the cell rear to allow cells to move forward in the direction of anchored leading edge [7,

56, 57]. A cascade of intracellular signal transduction events, including G proteins and tyrosine kinases, are involved in the events of cell migration cycle [7, 8, 58].

Mobile cells can be stimulated to migrate in a random, nonvectorial manner

(chemokinesis) or in a directional manner [7, 52]. The migration of SMCs can be directed

by the biochemical cues, including gradients in soluble chemical signals (chemotaxis) or

adhesive ligand density in the ECM (haptotaxis) [7, 52]. Directional cell migration can also be induced by the gradients in substrate mechanical stiffness (durotaxis or mechanotaxis) [59, 60].

2.3.2 Basic Mechanisms of SMC Proliferation

In the normal vessels, quiescent SMCs are maintained in a nonproliferative phase

(G0) [5, 6]. In response to vascular injury, SMCs stimulated by growth factors enter the

cell cycle, which comprises Gap (G) 1, DNA synthesis (S), G2, and mitosis (M) [5, 8].

Cell growth is one of the important events of G1, during which it prepares for DNA

replication in S [61]. In late G1phase, there is a restriction point (R). Before R point, cell

cycle progression is reversible. Growth factors trigger cell cycle entry and lead up to R

point [5]. After passage of this point, cells become irreversibly committed to go through

the rest of the cell cycle and following cell progression does not require further growth

factor stimulation [5]. Once it has duplicated its chromosomes (S), the cells enter another

gap phase (G2), when proteins are synthesized in preparation for mitosis (M). After cell

division, the daughter cells may enter G1 again for another cell cycle, or enter G0 [61].

The cell cycle is controlled by the expression and activities of regulatory proteins.

Cyclin-dependent kinases (CDKs) and their associated cyclins are the core activators of cell cycle progression [5, 62, 63]. Each CDK has a kinase subunit and a cyclin subunit.

As a monomer, the CDK has no enzymatic activity and can be activated by association with a cyclin protein. Different CDK/cyclin complexes are orderly activated at appropriate times in the cell cycle, which function to turn specific proteins on and off by phosphorylation [63, 64].

The CDK inhibitors (CDKIs) are key negative regulators of the cell cycle [62].

The CDKIs are structurally divided into two classes: the INK4 family and the KIP/CIP family [5]. The INK4 family, including p14, p15, p16, p18, and p19, inhibits exclusively

the complexes of CDK4/6-cyclin D [62-64]. The KIP/CIP family, including p21, p27,

and p57, inhibits a broad range of CDK-cyclin complexes [62-64].

2.4 Mediators of SMC Migration and Proliferation

SMC migration and proliferation are stimulated by a variety of extracellular signals, including soluble signaling factors, ECM proteins, proteinases, and physical factors (e.g., cyclic stress) [4, 5, 7, 14]. The extracellular stimulus activates cell surface receptors, which transduce the external signal to the intracellular signaling pathways and then triggers a series of coordinated cellular events, including cell migration and cell

proliferation.

2.4.1 The Role of Soluble Signaling Factors

2.4.1.1 Platelet-Derived Growth Factor

PDGFs are produced by platelets, macrophages, ECs, fibroblasts, and

keratinocytes [13, 65]. PDGFs comprise a family of four ligands, including PDGF-A, -B,

-C, and -D. All PDGFs function as homodimers, but only PDGF-A and -B can form

heterodimers. PDGFs bind to two different transmembrane tyrosine kinase receptor (α

and β), which can homo- and heterodimerize. It is well established and described that platelet-derived growth factors (PDGFs) promote SMC migration and proliferation in both physiological and pathological situations [66-68]. As described above, PDGF-B or

PDGF β receptor knockout mice fail to form normal vessels because of the reduced

capability of pericyte/SMC progenitor cell migration and proliferation [2, 11, 12]. SMCs

have been shown to upregulate the expression of PDGFβ receptor in response to vascular

injury, which contributes to increased SMC migration and proliferation in the intima [69].

The failure to downregulate the expression of PDGFβ receptor after vascular repair may

contribute to intimal hyperplasia and other vascular disease [14]. In vitro studies have

also shown that PDGFs promote SMC migration and proliferation [29, 70, 71]. It is of

note that while PDGF-BB and PDGF-AB are known SMC chemoattractants, PDGF-AA

is indicated to inhibit SMC migration [52]. The proliferative responses to PDGF isoforms

are not consistent between human arterial SMCs and venous SMCs [72]. The

proliferation of human venous SMCs can be stimulated by PDGF-BB but not PDGF-AA,

while the proliferation of arterial SMCs was more sensitive to PDGF-AA stimulation

[72]. In addition to regulating SMC migration and proliferation directly, PDGF can act

indirectly by stimulation of the synthesis of epidermal growth factor and fibroblast

growth factor-2, both of which facilitate SMC migration and proliferation [73, 74].

2.4.1.2 Heparin

Heparin has been reported extensively to inhibit SMC proliferation in vitro and in

vivo [75-77]. Heparin is a highly sulfated glycosaminoglycan and has been widely used

as an anticoagulant drug [78]. It has been well established that the anticoagulant and

antiproliferative properties of heparin are unrelated and reside in different heparin

domains [78-80]. The major structural determinant of heparin antiproliferative activity is

the amounts and distribution of sulfonate groups on the glycosaminoglycan chain, while

the anticoagulant activity is centered in a specific pentasaccharide sequence [78]. Heparin

has been shown to inhibit SMC proliferation induced by many stimulatory signals, such as serum and basic fibroblast growth factor (bFGF) [80-85]. However, there are

inconsistencies on the effect of heparin in inhibiting other stimuli, such as platelet-

derived growth factor (PDGF) [79, 82, 83]. Some studies show that heparin inhibits

PDGF induced proliferation [86-88], whereas other studies indicate it has no effect [82,

83]. Besides affecting SMC proliferation, heparin also has an inhibitory effect on SMC

migration [89-92].

Although the inhibitory effect of heparin on SMC migration and proliferation has

been described for some time, the mechanism of these effects remains unclear. Heparin has been shown to interrupt growth factor signaling by interfering with the binding of a

variety of growth factors to their receptors [93-95]. It might bind to the endogenous

growth factors and cytokines directly or displace growth factors from their binding site,

which then downregulates the regulatory effect of these growth factors on SMC

migration and proliferation [91]. Heparin has also been indicated to affect SMC functions

by modulation of cell cycle progression [96]. The maximum inhibitory effect of heparin on SMC proliferation is produced when heparin is present before cells enter the S phase.

It has been suggested that heparin can be internalized via cell-surface heparin sulfate proteoglycans and can activate the double stranded RNA-activated protein kinase (PKR) by direct binding and results in the block of G1-S transition [79, 84, 97]. The studies of heparin on SMC enzyme secretion have shown that heparin inhibits the expression of matrix-degrading proteinase such as plasminogen activators and matrix metalloproteinases (MMPs) [98, 99]. These proteinases have been demonstrated to

regulate SMC migration and proliferation in both physiological and pathological

conditions [4, 52].

2.4.1.3 Transforming Growth Factor beta

TGF-β has been shown to be a multifunctional cytokine with both stimulatory and inhibitory effects on SMC functions, including SMC migration and proliferation [100].

Active TGF-β is a 25 kDa homodimer of two 112 amino acid polypeptide chains and

mediates its effects by binding to membrane-bound serine/threonine kinase receptors

(TGF-β type I receptor and type II receptor) [79, 101]. Both receptors are necessary for

TGF-β signaling. The TGF-β ligand binds to a type II receptor, which recruits and

phosphorylates a type I receptor, leading to a formation of an interdependent heterodimeric complex. The activated type I receptor phosphorylates downstream

signaling molecules (e.g. the family of Smad signaling molecules) and regulates the

transcription of target genes [100-102]. The effects of TGF-β on SMC functions are

strongly dependent on culture conditions [103]. TGF-β has been shown to stimulate SMC migration [104, 105], while other studies suggest the antimigratory effect of TGF-β on

SMC migration [106, 107]. Furthermore, TGF-β has been shown to inhibit SMC

migration induced by other stimuli, such as PDGF [105]. The effect of TGF-β on SMC

proliferation is also not consistent. TGF-β has traditionally been known to inhibit SMC proliferation induced by serum, PDGF, and epidermal growth factor by inducing cell cycle arrest at G1 phase [45, 108, 109]. However, recent studies have shown that TGF-β promotes SMC proliferation through the Smad 3 and ERK MAPK pathways [110, 111].

It has been reported that TGF-β increases the synthesis of ECM proteins, including fibronectin, collagen type I, III, and V [112, 113], which have been indicated to promote

SMC migration and proliferation [52]. However, it decreases the synthesis of urokinase- type plasminogen activator and of tissue-type plasminogen, which are required for SMC migration and proliferation [103]. The regulatory effect of TGF-β on ECM synthesis and protease production may contribute to the regulatory effect of TGF-β on SMC functions.

2.4.1.4 Other Factors

Besides the factors detailed above, a wide variety of signaling factors have been implicated in regulation of SMC migration and proliferation, including basic fibroblast growth factor (bFGF), vascular endothelial growth factor, insulin-like growth factor-1

(IGF-1), and angiotensin II [4, 14, 52, 71, 103] (Table 2.1). It is notable that these

biochemical signals usually interact with each other, exerting a multifunctional effect on

SMC functions [14]. Species differences or specific culture conditions, such as serum concentration and presence of other stimulators or inhibitors, might induce different

responses.

2.4.2 The Role of Extracellular Matrix

The ECM is an active substrate that regulates SMC adhesion, migration and

proliferation via cell-matrix receptors, particularly integrins (Table 2.1) [3, 114].

Integrins are heterodimer receptors that are composed of α and β subunits. Each subunit

is composed of an extracellular, a transmembrane and a cytoplasmic component, which

transduce signals between the ECM and cell interior [115]. In response to vascular injury,

the expression of a variety of ECM proteins increases, including collagen (e.g. collagen

type I and VIII), osteopontin, fibronectin and vitronectin [116-121]. These proteins are

suggested to promote SMC migration via αvβ3 integrin signaling [115, 122]. Fibronectin

has been shown to promote SMC proliferation and has been used to coat substrates for the cultures of SMCs [123, 124]. Other ECM components, such as hyaluoronic acid, have been shown to stimulate SMC migration by binding of the CD44 receptor and the receptor for hyaluronic acid-mediated motility [3, 52, 121].

In addition to promoting SMC migration and proliferation, the ECM can inhibit

SMC migration and proliferation. In the normal, uninjured vessels, SMCs are firmly

Table 2. 1 Examples of regulatory biochemical factors on SMC migration and proliferation

Effect on smooth Effect on smooth Biochemical factors muscle cell muscle cell References migration proliferation Platelet derived growth factor + + [66-68] (PDGF) Heparin - - [80-85, 89-92] Transforming growth factor beta +/- +/- [100, 104-111] (TGF-β) Basic fibroblast growth factor + + [125, 126] (bFGF) Vascular endothelial growth factor - + [127, 128] (VEGF) Insulin-like growth + + [129, 130] factor-1 (IGF-1) Angiotensin II + +/- [131-134] Collagen + +/- [4, 7, 135] Osteopontin + + [4, 7, 136] Fibronectin + + [7, 123, 124] Vitronectin + + [4, 7] Hyaluoronic acid + + [3, 7, 52] Laminin + - [7, 123] Matrix metalloproteinases + + [7, 137] (MMPs) Tissue inhibitors of - - [7, 137] MMPs (TIMP) +: stimulatory effect; -: inhibitory effect;

adherent to the matrix and have low capability of migration and proliferation partially

due to the stable focal adhesions [14]. The basement membrane has been indicated to

prevent SMC migration and proliferation. In vitro studies, the basement membrane components, such as laminin, fail to promote the ability of SMCs to respond to growth factor stimulation [123, 138, 139]. In vivo, there is a rapid accumulation of fibronectin around proliferative SMCs after vascular injury, whereas the expression of laminin decreases [140]. These studies suggest that the basement membrane present an inhibitory effect on SMC migration and proliferation [138, 141]. Changes in collagen content have

also been suggested to affect SMC proliferation [4]. Mitogen-stimulated SMCs were able to proliferate when grown on monomer coated collagen, but fail to grow on polymerized collagen [135].

2.4.3 The Role of Proteases

The extracellular proteases, particularly matrix metalloproteinases (MMPs), play

a key role in regulation of growth factor availability, cell-matrix and cell-cell interactions

and thereby mediate SMC migration and proliferation [137, 142]. SMCs secrete MMP-1,

-2, -3, -7, -9, and -14 [143]. In response to vascular injury, the expression of MMP-2, -9,

and -14 by SMCs was shown to be upregulated as a result of PI3K activation [144-146].

The enhanced expression of MMPs promotes degradation of ECM, which stimulate SMC

migration and proliferation [79, 147]. Synthetic MMP inhibitors or overexpression of endogenous tissue inhibitors of MMPs (TIMP) proteins reduced neointimal thickening in vivo [148, 149]. The negative effect of synthetic MMP inhibitors on SMC migration also has been observed in vitro [148, 150]. In mice lacking MMP-2 or MMP-9, SMC migration and intima formation decreased [151-154]. Moreover, MMP inhibitors

repressed SMC proliferation in vitro and after angioplasty in vivo [148, 155, 156]. It is

suggested that the positive effect of MMPs is associated with their ability to degrade the

basement membranes, which will disrupt inhibitory effect of basement membrane

components and indirectly facilitate the accumulation of new matrix components (e.g.

fibronectin, osteopotin) [157, 158].

2.4.4 The Role of Physical Factors

The vascular media is subjected to dynamic mechanical stresses in vivo conditions [79, 159]. Cyclic mechanical loading is the dominant mechanical stimulus that

affects SMC functions [159, 160]. It has been reported that the application of cyclic

mechanical strain to cultured SMCs induces SMC proliferation and ECM production

[161, 162]. The in vitro studies of SMC culture on 2D surfaces have suggested that the

effect of cyclic strain on SMC proliferation is associated with ECM proteins [159]. At the

presence of cyclic strain, SMC proliferation was enhanced on fibronectin or vitronectin coated surfaces, compared with collagen or laminin coated surfaces [163]. In addition to dynamic mechanical stimuli, SMC functions can also be affected by changes in static mechanical environment. Studies using 2D cultures have suggested that the mechanical properties of the ECM (matrix stiffness) influence cell spreading, migration, and proliferation [164-166]. Cells preferentially migrate from less stiff to more stiff substrate

[60, 167]. SMC proliferation on 2D surfaces was enhanced on substrate with higher stiffness [168]. In some 3D studies, SMC proliferation has been shown to be not related with matrix stiffness [169].

2.5 Conclusions and Further Directions

The key role of SMC migration and proliferation on vascular physiologic and

pathologic remodeling necessitate the exploration of the mechanisms underlying these

functions. In the last several decades, a better understanding of the cellular and molecular

biology of SMC functions, including the regulatory effect of the extracellular signals on

SMC migration and proliferation, has emerged. These in vitro and in vivo studies conclude with suggestions that inhibiting SMC migration and proliferation might be beneficial for preventing or reducing the risk of intimal hyperplasia or other vascular disease. However, a great many questions remain to be answered regarding SMC migration and proliferation. In vivo, a large number of environmental cues, including growth factors, cytokines, ECM components, and cyclic strain, are working synergistically to regulate SMC functions. Most previous studies have only focused on the regulatory effect of one bioactive factor. The integration of these factors is largely

unstudied. In addition, most in vitro studies are performed on 2D surfaces. There is a growing appreciation that cells may respond differently when cultured in 3D and 2D systems. There is a clear need to develop an appropriate 3D model system to study SMC migration and proliferation. Advances in imaging technology and tissue engineering offer an opportunity to conduct future studies on these interesting processes in 3D cultures.

These approaches may provide some additional insights relevant to the behavior of SMCs in vivo.

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CHAPTER 3 Hydrogels as Extracellular Matrix Mimics for Three Dimensional

Cellular Studies

3.1 Introduction

In natural tissues, most cells are embedded within a three dimensional (3D) extracellular matrix (ECM) that is a highly hydrated, gel-like material composed of collagen fibers, proteoglycans and glycoproteins [1-3]. The ECM is a complex 3D fibrous meshwork that exhibits many biochemical and physical cues. This microenvironment is specific for each cell type and regulates cell functions, including cell adhesion, migration, proliferation, and differentiation [2, 4].

In vitro cell culture systems provide a defined platform that recapitulates many critical aspects of native ECM for investigating cell basic biology outside of the organism. Two dimensional (2D) substrates, such as tissue culture plastic (TCP) substrates or the surface of tissue analogs, have been extensively used as in vitro models

to study cellular events [5, 6]. These 2D experiments have provided much detail on the

mechanisms underlying cell behaviors, including the dynamic interactions between cells

and extracellular cues [3, 6]. However, cells in vivo reside in a 3D microenvironment, and

there is a growing appreciation that cells may respond differently when cultured in 3D

versus 2D systems [3].

One of the major drawbacks of the 2D cell culture systems is the lack of structural

architecture [5, 7]. In a 2D culture, cells are confined to adhere to a flat surface such that

only one face of the cell interacts with the ECM and neighboring cells, while the cell in

3D culture interact with the ECM on all surfaces [7]. The morphology of the cell and the

spatial distribution of adhesions may be fundamentally different. For example, fibroblasts

embedded in 3D matrices adapt an elongated, spindle-like morphology and form focal contacts using a more limited set of integrins than the flat cell on 2D matrices [8, 9]. This leads to differences in distribution of key signaling molecules as well as the differences of the types and rates of biochemical reactions, which will ultimately affect cell behaviors

[7, 8, 10]. Moreover, cells grown on 2D surfaces are exposed to a bulk culture media with a homogeneous concentration of nutrients, growth factors and cytokines. In contrast, the concentrations of soluble factors, such as cell-secreted enzymes or growth factors, often

present a dynamic concentration gradient in vivo [11].

Therefore, advanced 3D cell culture models that are more similar to the in vivo

cell-ECM microenvironment are needed to better understand cell behaviors in vivo.

Hydrogels have emerged as highly attractive materials for developing models for 3D cell

studies. These hydrophilic networks can provide a soft tissue-like 3D environment for cell growth and allow optimal transport of oxygen, nutrients and waste products [6, 12-

16]. Currently, a vast array of natural and synthetic materials has been used to fabricate hydrogels for cell culture [12-14]. This chapter briefly reviews the advantage and limitations of naturally derived materials, and summarizes recent progress of synthetic hydrogels.

3.2 Naturally Derived Materials

Naturally derived materials are widely used to form hydrogels for 3D cell studies.

They are either components of natural ECMs (e.g., collagen, fibrinogen, or hyaluronic acid (HA)) or have macromolecular properties similar to the natural ECM (e.g. alginate, chitosan) [14, 17, 18]. So far, naturally derived gels have been investigated for a variety of tissue engineering applications [13, 17, 18]. They have also been developed as 3D

models for fundamental cell studies [13]. This is because these materials have many

advantageous properties. First, many of the natural polymers, such as collagen and

fibronectin, have cellular binding domains that promote cell adhesion and further

regulation of cell functions [13]. For example, fibronectin has been reported to present a

variety of cellular binding sites to bind with the integrin receptors on the cell surface,

including the peptide sequence: RGD, KQAGDV, REDV, and PHSRN [19]. Second,

these materials are enzymatically or hydrolytically degradable, which allows for cell migration and ingrowth [14]. Collagen, for example, can be degraded by metalloproteinases, particularly collagenase, and serine proteinases [20]. Hyaluronic acid

(HA), which is found in nearly every mammalian tissue and fluid, is naturally degraded by hyaluronidase [20]. Third, many of these materials are intrinsically bioactive and have growth factor binding site to allow for the binding of soluble factors to regulate cell

functions [13]. It has been reported that the composite gel made from HA and fibronectin can modulate angiogenic processes stimulated by angiogenic growth factors and

cytokines [21]. Collagen has been shown to promote osteogenic differentiation of stem

cells activated by differentiation factors [22]. Finally, the gel preparation procedures of

these natural biopolymers are usually nondetrimental, which allows them to encapsulate

cells without affecting cell viability [13]. For instance, Matrigel (BD biosciences, San

Jose, CA), which is a gelatinous protein mixture secreted by Engelbreth–Holm–Swarm

(EHS) mouse sarcoma cells, can encapsulate cells by mixing cells with chilled Matrigel solution, and further incubating at 37 °C to allow for gelation [14]. The unique physiological properties of naturally derived materials have made them gold standard matrices for 3D cell culture models [9].

However, these materials also have several disadvantageous features. First, the resulting gels made from the naturally derived gels usually have weak mechanical properties, which make them difficult to handle and perform various cell studies [23]. For example, the elastic moduli of collagen gels are reported to be less than 5 kPa [23]. The poor mechanical properties of these gels restrict them from the use during long term cell culture, because these gels will readily undergo significant deformation during cell culture due to their limited resistance to cellular contractions [9, 13]. Second, the properties of these materials are difficult to modulate (e.g., biochemical and biophysical) because of the intrinsic bioactive properties of their precursors [9, 13]. For example, a change in the matrix stiffness of these gels by varying the solid content prior to gelation simultaneously alters the ligand densities and presentation [9, 24]. Third, the hydrogel properties are dependent on their derived sources, which results in lot to lot variability affecting experimental reproducibility [19]. As a consequence, these drawbacks may substantially reduce not only their adaptability to a wide range of clinical applications, but also restrict them to become ideal models.

3.3 Synthetic Poly (ethylene glycol) Hydrogels

Recently, synthetic polymeric hydrogels have emerged as an important alternative choice for 3D cell studies [25-28]. Compared with naturally derived materials, synthetic polymers have the distinct advantage of having consistent composition and predictable manipulation of properties, which is important for comparative studies when used as 3D cell culture models [12, 19]. PEG-based polymers are an important type of hydrophilic polymers that is widely used in tissue engineering because of their adjustable mechanical properties, design flexibility and intrinsic resistance to protein adsorption and cell

adhesion [13, 19]. The bio-inert PEG hydrogels can function as a blank slate to

incorporate bioactive factors in a controlled manner, which makes it possible to engineer

the PEG gels with desired bioactivities and examine their effects on cell responses [19,

29]. The current approaches for preparation of bioactive PEG hydrogels with the tunable

properties of biochemical and biophysical cues, such as cell adhesion, biodegradation,

and matrix stiffness, are briefly summarized here.

3.3.1 Approaches for Gel Gelation and Modification

Chemically cross-linked PEG based gels can be fabricated by a variety of

chemical reactions, including free radical polymerization, Michael-type addition

reactions, click reactions, and enzyme reactions [19, 27, 30]. Free radical polymerization,

especially photopolymerization, is the most common method to make PEG hydrogels

[19]. PEG acrylates are the major type of photopolymerizable macromers, including PEG diacrylate (PEGDA), PEG dimethacrylate (PEGDMA), and multiarm PEG acrylate (n-

PEG-Acr) [31, 32]. The liquid solution of these polymers with a biocompatible photoinitiator, such as Irgacure 2959 (4-[2-hydroxyethoxy]-penyl-[2-hydroxy-2-propyl]- ketone; Ciba Specialty Chemicals, Tarry-town, NY), can be converted to solid hydrogels at physiologic temperature and pH by exposure to specific light sources, including UV

light or lasers with a proper wavelength [13, 19, 32]. Therefore, this method allows in

situ encapsulation of cells within the 3D hydrogels [27]. Although photopolymerization

provides an effective method to fabricate gels with cell encapsulations, there is concern

that the use of initiators and UV light may affect cell viability [19, 33]. A mild Michael-

type addition reaction can be utilized to make hydrogels without these concerns.

Appropriately functionalized multiarm PEG (e.g., acrylate, maleimide and vinyl sulfone)

mixed with crosslinkers with thiol groups can form a polymeric network readily under

physiological conditions via a stepwise growth mechanism [34, 35]. Recently, click

reactions have been employed to form PEG hydrogels, which can efficiently link

bioactive components into the PEG polymeric network under physiologic conditions [36-

38]. Moreover, PEG hydrogels can be formed by enzymatic cross-linking reactions [9,

39-41]. The mild gelation conditions (e.g., low temperature, neutral pH, and in buffered

aqueous solutions) and the high selectivity of enzymes to their substrates allow for

fabrication of gels without the concern of side reactions and cellular toxicity [19].

3.3.2 Biochemical Tunability in PEG Hydrogels

Synthetic PEG hydrogels typically exhibit minimal or no intrinsic biological activity due to the resistance to protein adsorption and cell adhesion [19, 29]. This makes

PEG hydrogels able to serve as a blank platform for controlled bioactive modifications

[19]. To promote cellular functions, various strategies have been developed to mimic the

properties of ECM, including cell adhesion and biodegradation.

One possible method to biofunctionalize PEG hydrogels is to link ECM components into the polymeric network of PEG gels [42-44]. Various ECM proteins,

including collagen, fibrinogen, and laminin, have been chemically conjugated with PEG

macromers [13, 45-47]. For instance, fibrinogen was PEGylated with PEG diacrylates to form a bioactive hydrogels through photopolymerization. The biologic domains in the fibrinogen backbone provide cell adhesion sites for endothelial cell (EC) and smooth muscle cell (SMC) attachment as well as proteolytic degradation. The conjugation of fibrinogen into the hydrogel promoted SMC invasion through the hydrogels [46].

Although full proteins incorporated into hydrogels can provide biochemical cues

to encapsulated cells, ECM-derived short peptides have emerged as an attractive

alternative for the biochemical modification of PEG hydrogels [6, 19, 48]. Peptides

contain the same functional domains as the proteins, so they can exert biological

functions similar to those presented by the full proteins [6]. In contrast with the full

proteins, which are subject to denaturation and degradation, bioactive peptides are relatively stable for modifications, tunable for cell binding, and easy to synthesize and purify, which ultimately enables a greater level of biological control than the full proteins

[48].

To date, a variety of bioactive peptides with different biological functions have been incorporated into PEG hydrogels. For example, PEG hydrogels can be rendered cell adhesive by the incorporation of a cell adhesive peptide containing the RGD sequence to the polymeric network [49, 50]. RGD is the cell binding domain derived from ECM proteins, including fibronectin, laminin, and collagen [19, 49]. Extensive studies have suggested that cell functions, including cell adhesion, migration, and proliferation, can be regulated by the quantity of RGD peptides in the hydrogel [9, 51-57]. In addition to RGD sequences, a variety of ECM-derived cell adhesive peptides have been tethered into PEG hydrogels for cell-adhesive modification (Table 3.1).

To generate degradable PEG hydrogels, various enzyme-sensitive peptides have been incorporated to the macromer backbone (Table 3.2). The degradation rate of the hydrogels can be tuned by the sensitivity of peptides to the enzymes, which will subsequently affect cell functions, such as cell migration and proliferation [52, 58, 59].

For example, PEG hydrogels with different degradability have been fabricated by mixing

four-arm-PEG-vinyl sulfone with oligopeptides with different MMP sensitivity through

Michael-type added reactions. Encapsulated fibroblasts showed increased cell spreading,

migration, and proliferation when cultured in 3D hydrogels with faster degrading peptides [58].

3.3.3 Biophysical Tunability in PEG Hydrogels

The biophysical properties of a hydrogel, including stiffness, initial mesh size,

and swelling ratio, are directly related to the cross-linking density of the network. An

important advantage of PEG hydrogels is the ability to tune their network properties over

a wide range simply by varying the molecular weight (MW) and/or concentration of PEG

[13, 24]. Through manipulating the MW and concentration of PEGDA in the hydrogel,

uncoupled investigation of scaffold modulus and mesh size on smooth muscle cell

behavior were permitted [24]. PEG monoacrylates have been commonly used to

conjugate cell adhesive peptides or full proteins for cell adhesive modification of PEG

hydrogels. Investigation of the effect of PEG monoacrylates on the properties of PEGDA

hydrogels has suggested that the addition of PEGMA to gels with fixed PEGDA

composition resulted in a decreased swelling ratio and increased shear moduli only when

the concentration of PEGMA was high relative to the concentrations used to promote cell

attachment [60]. This will allow for investigation of biochemical properties on cell

functions without sacrificing the network properties of hydrogels [52, 53].

3.4 Studies of Smooth Muscle Cell Migration and Proliferation in 3D Scaffolds

As described in Chapter 2, smooth muscle cell (SMC) migration and proliferation

are not only important for guiding the development of vascular tunica media for

functional tissue engineered blood vessels, but also contribute to the development intimal

hyperplasia and other vascular pathologies. Understanding the mechanism of SMC

migration and proliferation would be of great benefit in developing new treatments for

vascular disease, as well as in fabricating functional tissue engineered blood vessels.

Naturally derived materials have been utilized to form gels for 3D SMC

mechanistic studies [61-63], which provides much detail on the mechanisms of 3D SMC

migration and proliferation. However, the poor engineering properties of naturally derived materials (e.g., the interdependence of scaffold variables) have hampered the uncoupled investigation of scaffold properties on SMC functions, inhibiting the rational selection of scaffold variables to achieve desired cell responses [7, 13, 19]..

3D synthetic PEG hydrogels have been rendered cell-adhesive and/or biodegradable to investigate SMC migration and proliferation in 3D [24, 64-66]. For example, modified Boyden Chamber assay has been utilized to investigate SMC migration through 3D biomimetic PEG hydrogels [64]. The studies have shown that both cell-adhesive peptide and degradable peptide are required for cell migration to occur [64].

However, the systematic investigation of both biochemical properties (e.g., cell adhesivity, biodegradability) and physical properties (e.g., cross-linking density) on SMC migration and proliferation is limited [52]. SMC proliferation has been studied by homogeneous seeding of SMCs into 3D PEG gels [24, 65, 67]. Due to the lack of fast degradation mechanisms in gels, cells encapsulated in these gels often present a round morphology instead of a normal spindle-like morphology [24, 67]. The absence of SMC spreading in these 3D constructs might affect SMC viability and proliferation and hamper investigation of the effect of both scaffold properties and exogenous bioactive factors on

SMC proliferation [24, 67, 68].

3.5 Specific Aims

To better understand the mechanisms underlying SMC migration and

proliferation, we developed a synthetic, peptide-bearing hydrogel as a model system for

study of SMC behaviors in 3 dimensions. Our overall hypothesis is that 3D SMC migration and proliferation can be regulated by the properties of a cell-instructive

scaffold, including cell-matrix adhesion, degradability, and cross-linking density, as well

as exogenous bioactive factors. This thesis has 3 specific aims to investigate the

hypothesis:

Aim 1: Engineer a PEG hydrogel with the property of cell adhesion and biodegradation. To accomplish this aim, poly (ethylene glycol) (PEG) based hydrogels were designed as the scaffold substrate. To mimic properties of the extracellular matrix

(ECM), cell adhesive peptide (GRGDSP) and enzyme sensitive peptide (GPQGIAGQ,

VPMSMRGG) were incorporated into the PEG macromer chain. The cell adhesivity and degradability of hydrogels were determined. The swelling ratio of hydrogels was studied to investigate the effect of gel composition on gel network properties.

Aim 2: Examine the effect of scaffold properties on SMC migration in 3D gel.

We hypothesized that SMC migration in 3D hydrogels is governed by cell-matrix

adhesion, proteolysis, and cross-linking density. By manipulating gel compositions, the

effect of biochemical property (e.g.: adhesive peptide concentration) and biophysical

property (e.g., cross-linking denstiy) of 3D gels on SMC migration were evaluated

systematically.

Aim 3: Quantify the ability of biomimetic hydrogel to regulate SMC

proliferation. The hypothesis is that SMC proliferation in 3D gels can be regulated via

the alteration of gel characteristics as well as exogenous bioactive factors. The effect of scaffold properties (e.g., cell-matrix adhesion, MMP sensitivity, cross-linking density) on

SMC spreading and proliferation in 3D was investigated. The optimal gel composition for SMC proliferation was determined, and the effect of heparin as an exogenous factor on SMC proliferation was quantified.

Table 3. 1 Examples of cell-adhesive peptides that have been used for cell-adhesive modification of PEG hydrogels

Cell-adhesive Origin Cell receptor Reference peptides

RGD Fibronectin, laminin, Integrins [9, 51-55, 57]

collagen

KQAGDV Fibronectin Integrins [56, 57, 66, 69, 70]

REDV Fibronectin Integrins [71]

PHSRN Fibronectin Integrins [72-74]

IKVAV Laminin α1 110 kDa protein [43, 75-77]

YIGSR Laminin 67 kDa protein [56, 74-76]

PDGSR Laminin Integrins [75, 76]

LRGDN Laminin Integrins [76]

LRE Laminin Integrins [75]

IKLLI Laminin Heparin [75, 76]

GFOGER Collagen-1 Integrins [70]

VAPG Elastin 67 kDa protein [57, 69, 76, 78]

Table 3. 2 Examples of enzyme-sensitive peptides that have been used for proteolytic modification of PEG hydrogels

Enzyme-sensitive Origin Sensitive enzyme Reference peptide

GPQGIAGQ Collagen-I MMP-2 [52, 53, 79]

GPQGIWGQ Peptide library MMP-8 [9, 53, 54, 80-84]

GPQGILGQ Collagen-I MMP-1 [38]

LGPA Peptide library MMP-1 [66, 85-88]

APGL Peptide library MMP-1 [89]

YKNRD Fibrinogen Plasmin [84, 90]

VRN Fibrinogen Plasmin [89]

AAAAAAAAA Peptide library Elastase [51, 66]

AAPVRGGG Peptide library Elastase [91]

PENFF Aggrecan MMP-13 [92]

VPMSMRGG Peptide library MMP-9 [58, 93, 94]

IPESLRSG Peptide library MMP-2 [58]

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CHAPTER 4 Engineer a Cell-adhesive Biodegradable Hydrogel for 3D Cellular

Studies

Based on: Lin L, Zhu J, Kottke-Marchant K, Marchant RE. Biomimetic engineered poly

(ethylene glycol) hydrogel for smooth muscle cell migration. Tissue Eng Part

A 2014; 20: 864-73.

4.1 Introduction

Two dimensional (2D) cell cultures have been extensively used as in vitro models to study the principles of cell biology, which has provided the basis for understanding how cell functions in response to environmental cues [1-3]. However, the commonly used

2D substrates, such as tissue culture plastic (TCP) substrate or the surface of tissue analogs, introduce an asymmetry that maybe unlike the in vivo conditions since most cells are embedded within a three dimensional (3D) extracellular matrix (ECM) [1-3].

There are increased findings that have elucidated the disparity of cell functions in 2D versus 3D environments [4-6]. Thus, it is of great importance to engineer 3D bioactive scaffolds as in vitro models for 3D cell studies.

An ideal 3D cell culture model should not only present the basic biological properties (e.g., cell adhesivity, biodegradability), but also have the ability to tune their different scaffold properties independently [1, 2, 4, 5]. Hydrogels have demonstrated a distinct efficacy for 3D cell studies due to their numerous similarities with the cells’ native environment. These hydrophilic networks can provide a soft tissue-like 3D environment for cell growth and allow optimal transport of oxygen, nutrients and waste products [7-12].To date, naturally derived materials (e.g. collagen, fibrin) have been frequently used to form hydrogels for 3D cellular studies because of their inherent

biological functions like cell adhesion and biodegradation [13]. However, these inherent

functions also result in the difficulty of modifying the different features of these 3D

microenvironments independently [4]. For example, it is impossible to modulate the

cross-linking density of collagen gel without affecting its density of cell-adhesive binding

site [4, 8]. Besides the limited engineering properties, the relatively poor mechanical

properties and batch-to-batch variability of these naturally derived materials also restrict

their potential to become an ideal model [8, 14].

Synthetic poly (ethylene glycol) (PEG) hydrogels have attracted broad interest as

scaffold materials for tissue engineering applications, because of their tunable mechanical

and chemical properties [8, 14]. The unmodified PEG hydrogels resist protein adsorption

and cell adhesion, which allows for incorporation of biological ligands in a controlled

manner [14, 15]. For example, PEG hydrogels can be rendered cell-adhesive by the

incorporation of a cell-adhesive peptide (e.g. Arg-Gly-Asp [RGD]) to the polymeric

network [16-19]. To tune the degradation rate of PEG hydrogels systematically, enzyme- sensitive peptides or α-hydroxy acids, such as lactic acid, have been conjugated to the macromer backbone [20-28]. Growth factors or other bioactive molecules also have been incorporated in PEG gels to study their effect on cell functions [29-31]. Further, network properties of PEG hydrogels can be tuned by simply varying the molecular weight (MW) and/or concentration of PEG [32-34].

The objective of this work is to engineer a cell-adhesive, proteolytically degradable PEG hydrogel with tunable biochemical and biophysical properties. Photopolymerizable PEG diacrylate (PEGDA) derivatives were used to fabricate hydrogels as scaffold substrates.

These PEG diacrylates were engineered to be proteolytically degradable by incorporation

of enzyme-sensitive peptide, including GPQGIAGQ (GIA) derived from collagen type I

[35], or VPMSMRGG (VPM) derived from a peptide library [26], into the backbone of

PEGDA. It has been reported that VPM peptide is more sensitive to enzyme (e.g. matrix metalloproteinases [MMPs]) degradation than GIA peptide (Table 4.1) [26]. To render

PEG hydrogels cell adhesive, GRGDSP peptides derived from fibronectin [36], were grafted into PEG hydrogels via PEG monoacrylates (PEGMA) during copolymerization with proteolytically degradable PEGDA by photopolymerization. The cell adhesivity, biodegradability, and network properties of these bioactive hydrogels were examined.

4.2 Materials and Methods

4.2.1 Materials

All reagents were obtained from Sigma-Aldrich (St. Louis, MO) and used as received unless otherwise stated.

4.2.2 Preparation of Bioactive Peptides

The cell-adhesive peptide (GRGDSP [RGD, MW: 586 Da]) and diaminopropionic acid (Dap)-capped enzyme-sensitive peptides (VPMSMRGG-Dap [VPM-Dap, MW: 919

Da] and GPQGIAGQ-Dap [GIA-Dap, MW:812 Da]) were synthesized on an amide

(Knorr) resin using standard Fmoc chemistry on a solid phase peptide synthesizer

(Applied Biosystems, Model 433A, Foster City, CA). The peptides were cleaved from the resin using trifluoroacetic acid and purified by reverse-phase high-performance liquid chromatography (Waters, 2690 Alliance system, Milford, MA). Successful peptide synthesis was confirmed by matrix assisted laser desorption/ionization mass spectroscopy

(MALDI-MS, Bruker, Autoflex III, Fremont, CA).

-1 -1 Table 4. 1 -Comparison of kinetic parameters (kcat/Km, M s ) for GIA and VPM peptide from the literature

Peptide Sequence

Enzyme GPQGIAGQ VPMSMRGG

MMP-1 60.6 [37] 1600 [38]

MMP-2 180 [37] 24,000 [38]

MMP-3 16.7 [37] 3900 [38]

MMP-7 110 [37] 7900 [38]

MMP-8 1570 [37] -

MMP-9 93.9 [37] 51,000 [38]

MMP-11 - -

MMP-13 - -

MT1-MMP - 6100 [38] Km: the Michaelis-Menten constant. It is the substrate concentration needed to achieve a half-maximum enzyme velocity.

Kcat: the turnover number. It is the number of times each enzyme site converts substrate to product per unit time.

Kcat/Km: a measure of enzyme efficiency. Either a large value of Kcat (rapid turnover) or a small value of Km (high affinity for substrate) makes Kcat/Km large.

4.2.3 Preparation of Biomimetic Macromers

Biomimetic macromers were synthesized by conjugating bioactive peptides with

Acrylate-PEG-Succinimidyl Valerate (Acr-PEG-SVA, MW: 3400 Da; Laysan Bio, Arab,

AL) (Fig. 4.1). To modify PEG with cell-adhesive peptide, Acr-PEG-SVA was reacted with GRGDSP (15% molar excess) in aqueous sodium bicarbonate solution (50 mM, pH

8.5) at room temperature (RT) for at least 4h. Then, the product of RGD modified PEG

monoacrylate (RGD-PEGMA) was dialyzed against water with membranes of molecular

weight cut off (MWCO) 2000 for 48 h, to remove salts and unreacted peptides. The

purified peptide was lyophilized and stored at -20 °C. The enzyme-sensitive peptide

modified PEG diacrylates (VPM-PEGDA/GIA-PEGDA) were synthesized by the same

method as RGD-PEGMA using Acr-PEG-SVA with VPM-Dap or GIA-Dap peptide in a

molar ratio of 2:1 (PEG: peptide). The final product was dialyzed against water with

membranes of MWCO 5000 for 48 h. The synthesis of biomimetic macromers was confirmed by MALDI-MS.

4.2.4 Hydrogel Preparation

Bioactive hydrogels were prepared at various compositions (RGD-PEGMA: 0-5

mM, VPM/GIA-PEGDA: 4-6% [w/w] in phosphate buffered saline [PBS, pH 7.4]) using

0.1% (w/v) of Irgacure 2959 (4-[2-hydroxyethoxy]-penyl-[2-hydroxy-2-propyl]-ketone;

Ciba Specialty Chemicals, Tarry-town, NY) as the photoinitiator. To convert a liquid polymer to a hydrogel, the hydrogel precursor solution was dispensed into a stainless steel mold (D=6 mm, H=1.2 mm) and then polymerized by 10 min exposure to a UV light (365 nm, 5-10 mW/cm2).

Figure 4. 1 Synthesis scheme of cell-adhesive peptide modified PEG monoacrylates (RGD-PEGMA) and enzyme-sensitive peptide modified diacrylates (VPM-PEGDA and GIA-PEGDA).

4.2.5 Hydrogel Swelling

The hydrogel disks were prepared as described above, and allowed to swell in

excess distilled water at RT for 48 h. The swollen gel was weighed and lyophilized. The

mass of swollen hydrogel (ms) and the mass of the hydrogel after lyophilization (mp) were recorded respectively. The mass swelling ratio, q, was then determined by the following equation:

(1)

4.2.6 Analysis of Hydrogel Network Structure

The hydrogel mesh size (ξ) was determined by Flory−Rehner calculations using

mass swelling ratio (q) [39-41]. First, the molecular weight between cross-links (Mc) was

calculated using the following equation:

(2)

where:

(3)

(4) and M is the number-average molecular weight of the un-cross-linked hydrogel (the

molecular weight of the polymer), V1 is the molar volume of the solvent (water, 18

3 cm /mol), α1 is the polymer−solvent interaction parameter (0.426 for PEG−water and

3 assumed constant for our work), ρp is the density of the dry hydrogel (1.12 g/cm for

3 PEG), and ρs is the density of the solvent (1 g/cm for water).

Mesh size were then determined as described by Canal and Peppas [41] using the

following equations: 79

2 (5)

(6) where l is the average bond length (0.146 nm), Cn is the characteristic ratio of the

polymer (typically 4.0 for PEG), and Mr is the molecular weight of the repeat unit (44 for

PEG).

4.2.7 In vitro Degradation of Hydrogels

Degradation of hydrogels modified with enzyme-sensitive peptide was measured

after treatment with active MMP-2. Briefly, gel disks were incubated in Hank’s buffered

saline solution (HBSS) for 48 h to allow for pre-equilibration. Proenzyme MMP-2 (R&D

systems, Minneapolis, MN) was incubated in p-aminophenylmercuric acetate solution

(APMA, 1 mM in HBSS) at RT for 1 h, which activates the pro-form to activated form.

The swollen gels were weighed and then incubated in the activated MMP-2 solutions for

up to 24 h at 37°C. Proteolytic degradation of the hydrogels was monitored by measuring

the weight loss of the hydrogels at predetermined time intervals. The weight percentage

was determined by dividing the mass of gel sample at each time interval by the mass of

initial swollen hydrogel before MMP-2 incubation. Enzyme-insensitive hydrogels formed

from IGA-PEGDA (synthesized by the same method as VPM/GIA-PEGDA using the

reaction of Acr-PEG-SVA with the scrambled peptide: GPQIGAGQ-Dap) were prepared

as negative controls.

4.2.8 Cell Culture

Human coronary artery smooth muscle cells (HCASMCs, Lonza, Walkersville,

MD) were maintained at 37 ºC, 5% CO2 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. For

all experiments, HCASMCs at passages P8-P11 were used.

4.2.9 Cell Attachment and Spreading on the Hydrogel

To examine cell adhesivity, GIA-PEGDA (5%, w/w) hydrogels with various

concentrations of RGD-PEGMA (0-5 mM) were prepared. Briefly, hydrogel precursor

solution was sterilized by filtration (0.22 µm pore), and then dispensed into a sterilized

stainless steel mold and covered with a cover slip. The gel disks (D=10 mm, H=1.2 mm)

were formed as described previously. Hydrogels were then transferred to a 24-well

culture plate and incubated in PBS to swell overnight. Prior to seeding, gels were rinsed

with PBS three times and secured with a ring of silicone tubing. HCASMCs were seeded

at a density of 3.0×104 cells/cm2 on hydrogels in serum-free medium (SFM: insulin-

trasferrin-selenium supplement [ITS-X, 1×, Invitrogen, Carlsbad, CA], taurine [5 mM],

bovine serum albumin [BSA, 1 mg/ml] in Dulbecco’s Modified Eagle’s Medium

[DMEM, invitrogen]) for 6 h. Then, the medium was changed to SmGM-2 growth

medium. Human fibronectin (FN, 1 µg/cm2) was coated on tissue culture polystyrene

(TCPS) as positive controls. GIA-PEGDA hydrogels with 5 mM of RDG-PEGMA

(synthesized by the same method as RGD-PEGMA using the reaction of Acr-PEG-SVA

with the scrambled peptide: GRDGSP) were prepared as negative controls. Following incubation for 24 h after seeding, cells were imaged on a Nikon Diaphot 200 Inverted

Phase Contrast Microscope (10×). The silicone tubing rings were removed and samples were washed gently with PBS and frozen at -80 ºC. 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

reader (BioTek, Winooski, VT) at the emission wavelength λem of 520 nm (excitation at

λex of 480 nm).

4.2.10 Statistics

Statistical analysis was done using Origin 8.0 and Minitab 1.6. Data are represented as mean ± standard deviation of at least triplicate samples. Single

comparisons were made using an un-paired student’s t-test. Analysis of variance

(ANOVA) followed by Turkey’s post hoc test was used for data sets with multiple

comparisons. A value of p<0.05 was considered statistically significant.

4.3 Results

4.3.1 Synthesis and Characterization of Biomimetic Macromers

The preparation of biomimetic macromers was performed by utilizing the reaction

between the free amino group on the peptide and the N-hydroxysuccinimide (NHS) group

on Acr-PEG-SVA (Fig. 4.1) [42]. Figure 4.2A shows the MALDI spectrum of unmodified Acr-PEG-SVA with a maximum peak at 3820. After conjugation with RGD peptide in a ratio of 1:1, the maximum peak of RGD-PEGMA was shifted to 4255 (Fig.

4.2B), indicating successful conjugation of Acr-PEG-SVA with RGD peptide. To conjugate one molecule of enzyme-sensitive peptide with two molecules of Acr-PEG-

SVA, the C-terminus of VPM or GIA peptide was capped with diaminopropionic acid to provide an additional free amine. The MALDI spectrum of GIA-PEGDA with a maximum peak at 8185 (Fig. 4.2C) and VPM-PEGDA with a maximum peak at 8322

(Fig. 4.2D) confirmed successful conjugation of one molecule of enzyme-sensitive peptide with two molecules of Acr-PEG-SVA.

Figure 4. 2 MALDI characterization of biomimetic macromers. (A) MALDI mass spectrum of Acr-PEG-SVA with a maximum peak at 3820. (B) MALDI mass spectrum of RGD-PEGMA with a maximum peak at 4255. (C) MALDI mass spectrum of GIA- PEGDA with a maximum peak at 8185. (D) MALDI mass spectrum of VPM-PEGDA with a maximum peak at 8322.

4.3.2 Hydrogel Swelling and Network Properties

Hydrogels with various compositions (RGD-PEGMA: 0-5 mM, VPM/GIA-

PEGDA: 4%-6% [w/w]) were prepared and the mass swelling ratio was determined

gravimetrically (Fig. 4.3). For hydrogels with a fixed concentration of VPM-PEGDA,

increasing the concentration of RGD-PEGMA either did not change, or slightly

decreased, the swelling ratio of hydrogels, when the RGD-PEGMA concentration was

relatively low (0-2.5 mM) (Fig. 4.3A). When the concentration of RGD-PEGMA reached

5 mM, there was a significant decrease relative to 0 mM RGD-PEGMA for 4% and 5%

(w/w) VPM-PEGDA hydrogels (Fig. 4.3A).

To study the effect of PEGDA concentration on the hydrogel network, mass

swelling ratio of gels formed from 4%-6% (w/w) of VPM-PEGDA or GIA-PEGDA

(RGD-PEGMA: 0 mM) was determined (Fig. 4.3B). The swelling ratio of hydrogels

decreased substantially with increasing VPM-PEGDA concentration from 4% to 6%

(w/w) (Fig. 4.3B). The substitution of VPM-PEGDA with GIA-PEGDA did not change

the swelling ratio of hydrogels significantly (Fig. 4.3B).

To estimate the network structure of these hydrogels, the mass swelling ratio was further used to calculate the number-average molecular weight between cross-links (Mc) and the average mesh size was determined by Flory-Rehner calculations (Table 4.2). The increase in VPM-PEGDA or GIA-PEGDA concentration results in a decreased Mc and

mesh size, which indicates an increased network cross-linking density. The substitution

of VPM-PEGDA with GIA-PEGDA did not change Mc and mesh size significantly.

Figure 4. 3 The effect of gel compositions on mass swelling ratio of hydrogels. (A) Mass swelling ratio of hydrogels as a function of RGD-PEGMA (0-5 mM) for various VPM- PEGDA compositions (4-6%, w/w). (B) Mass swelling ratio of hydrogels as a function of PEGDA (VPM /GIA-PEGDA: 4-6%, w/w; RGD-PEGMA: 0 mM). *: p<0.05 with regard to VPM-PEGDA hydrogels without RGD-PEGMA; #: p<0.05 with regard to 4% PEGDA hydrogels; ψ: p<0.05 with regard to 5% PEGDA hydrogels.

Table 4. 2-The effect of PEGDA concentration on the network properties of hydrogels

Hydrogel type Polymer concentration Mc (g/mol) Mesh size (Å) (%, w/w) VPM-PEGDA 4 3974±25 210.8±6.2 5 3812±46 177.1±5.4 6 3320±144 135.2±7.9 GIA-PEGDA 4 3915±37 210.2±8.9 5 3721±80 172±8.8 6 3263±48 133.2±2.6 Mc: the number-average molecular weight between cross-links.

4.3.3 In vitro Degradation of Hydrogels

The mass change of enzyme-sensitive peptide modified hydrogels in activated

MMP-2 solution was measured to study the effect of enzyme-sensitive peptide type and

enzyme concentration on hydrogel degradation (Fig. 4.4). At the constant concentration of MMP-2 solution (1 µg/ml), VPM-PEGDA (5%, w/w) hydrogels had faster degradation

time (2 h) than GIA-PEGDA (5%, w/w) hydrogels (7 h) (Fig. 4.4A). As expected,

hydrogels modified with scrambled peptide (GPQIGAGQ) showed no degradation, as

indicated by no significant mass change over the incubation period (Fig. 4.4A). In the

presence of MMP-2, enzyme-sensitive peptide modified hydrogels (VPM-PEGDA and

GIA-PEGDA) initially swelled, likely due to cleavage of enzyme-sensitive sequences,

resulting in a decrease of cross-linking density with increased water diffusion into

hydrogels. Through further incubation in enzyme solution, the hydrogels lost mass with

time reflecting the cross-linking density becoming too low to maintain the hydrogel

network. VPM-PEGDA hydrogels were degraded completely within 2 h in 1 µg/ml

MMP-2 solution. When the MMP-2 concentration decreased, the rate of hydrogel

degradation decreased (Fig. 4.4B). Hydrogels incubated in the buffer solution without

MMP-2 did not degrade over the incubation period of 8 h (Fig. 4.4B), indicating that the

degradation of gels was due to the presence of enzyme.

4.3.4 Cell Attachment and Spreading on the Hydrogel

The effect of adhesive ligand concentration on HCASMC attachment and spreading was assessed by utilizing GIA-PEGDA (5%, w/w) hydrogels with varying

concentrations of RGD-PEGMA (0-5 mM) (Fig. 4.5). Few cells adhered and cell

Figure 4. 4 Degradation of MMP sensitive peptide modified hydrogels in MMP-2 solution at 37 °C. (A) The effect of MMP sensitivity on PEGDA hydrogel (5%, w/w) degradation in MMP-2 solution (1 µg/ml). Hydrogels modified with scramble peptide (GPQIGAGQ-Dap) served as the negative control. (B) The effect of MMP-2 concentration (0-1 µg/ml) on VPM-PEGDA hydrogel (5%, w/w) degradation.

Figure 4. 5 SMC attachment and spreading (after a 24 h seeding period) as a function of RGD concentration (RGD-PEGMA: 0-5mM) on GIA-PEGDA (5%, w/w) hydrogels. (A) Phase contrast micrographs (10×) of cell morphology on RGD modified hydrogels. Hydrogels with 5 mM RDG (RDG-PEGMA) served as the negative control. Human fibronectin (FN, 1 µg/cm2) coated tissue culture polystyrene (TCPS) served as the positive control. (B) Cell attachment on RGD modified hydrogels. Attachment was quantified by measuring DNA content reported relative to the positive control: FN. *: p<0.05 with respect to hydrogels with 5 mM RGD; #: p<0.05 with respect to FN.

spreading was not observed after 24 h on hydrogels without RGD modification (Fig.

4.5A). In contrast, by including RGD-PEGMA into the hydrogels, cell attachment and

spreading was supported (Fig. 4.5A). SMCs on PEG hydrogels with 5 mM of RGD-

PEGMA were well spread, similar to SMCs on the FN coated surfaces (Fig. 4.5A). DNA

quantification showed that cell attachment increased and reached a plateau as the RGD-

PEGMA concentration was increased up to 0.125 mM (Fig. 4.5B). However, SMCs did not adhere to hydrogels with a high concentration of the scrambled peptide GRDGSP (5

mM), which was similar to the result for plain PEGDA hydrogels (Fig. 4.5).

4.4 Discussion

The evidence of discrepancies in cell functions between 2D and 3D studies

necessitates the development of valid 3D models to further elaborate the regulatory effect of extracellular environments on cell functions [1, 2, 4, 5]. The goal of this project is to engineer an ECM-mimetic hydrogel with tunable biochemical and physical properties for further application of 3D cell culture models or tissue engineering scaffolds. Synthetic

PEG hydrogels were selected as the scaffold in this study because PEG resists protein adsorption and cell attachment, providing a biological blank slate for controlled bioactive modifications [14, 15].

The most basic biological functionality of an ideal 3D model is cell adhesivity, which can be achieved by the addition of cell-adhesive peptide [4]. To render PEG hydrogels cell-adhesive, cell-adhesive peptide (GRGDSP, derived from fibronectin [36]) modified PEG monoacrylate (PEGMA) was incoporated into PEG hydrogels by photopolymerization. Our experiments indicate that the incorporation of cell-adhesive peptides into the PEG network facilitates SMC attachment and spreading on hydrogels

(Fig. 4.5),which is attributed to the specific binding of RGD ligands to the integrin

receptors on SMCs [17]. RGD is the most commonly used cell-adhesive peptide to

facilitate the binding of many cell types, such as SMCs, fibroblasts, and endothelial cells

[16-19]. The promotion of SMC attachment has been observed in the scaffolds modified

with other cell-adhesive peptides, such as KQAGDV derived from fibronectin and VAPG

derived from elastin [17, 43]. By varying the concentration of adhesive ligands, cell-

matrix adhesiveness can be modulated [16-19, 43]. Our results show that cell attachment

increased with increasing RGD-PEGMA concentration (up to 0.125 mM), and was

comparable to FN coated surfaces at a RGD concentration of 2 mM (Fig. 4.5B). The

concentration of adhesive ligand to reach optimal cell attachment and spreading varies in

published reports [44, 45]. This is because multiple other factors, such as mechanics [44],

seeding density [42], also affect cell adhesion.

In 3D cellular studies of cell migration, cell proliferation or matrix remodeling,

the embedded cells will need to overcome the steric resistance of their environment [3, 4,

25]. Depending on the type of cell and matrix, either mechanistic strategy of changing

cell morphology and/or matrix degradation are utilized by cells to overcome physical

resistance [3, 4, 25, 46]. Since the mesh size of the synthetic 3D scaffold (Table 4.2) is much smaller than the dimensions of many cells (e.g., smooth muscle cells, fibroblasts, and endothelial cells), the 3D scaffold must contain structural entities that are susceptible

to degradation, either through proteolytic cleavage or hydrolysis [4]. To mimic

proteolytic degradation mechanisms presented in native ECM, various enzyme-sensitive

peptides have been incorporated into PEG network for enzymatic degradation [20-27].

The enzyme-sensitive peptides chosen for this study include GPQGIAGQ (GIA), derived

from collagen type I [35], and VPMSMRGG (VPM), derived from peptide library [26].

To verify the degradability of hydrogels, the mass change of enzyme-sensitive peptide modified hydrogels was monitored after treatment of MMP-2 (Fig. 4.4). MMP-2 has been demonstrated to be expressed by many cell types, including smooth muscle cells, endothelial cells, and fibroblasts [47, 48]. Our results show that both GIA-PEGDA and

VPM-PEGDA hydrogels can be degraded by MMP-2 and the degradation date of enzyme-sensitive peptide modified hydrogels depends on the amount of MMP-2 present

(Fig. 4.4). Compared with GPQGIAGQ peptide, VPMSMRGG peptide has been reported to be more proteolytically sensitive to many MMPs, such as MMP-1, -2, -3, and

-9 (Table 4.1) [26]. As expected, VPM modified hydrogels degrades faster than GIA modified hydrogels in the presence of MMPs [26]. Our in vitro degradation results also

show that VPM-PEGDA hydrogels has a faster degradation rate than GIA-PEGDA

hydrogels at the same MMP-2 concentration (Fig. 4.4).This indicates that simple changes

of the peptide sequence type in the hydrogel network could tune the degradability of hydrogels [26], which allows the examination of proteolytic sensitivity on 3D cellular functions (e.g., cell migration, cell proliferation).

The major disadvantage of naturally derived materials is their difficulty to tune the different features of 3D scaffolds independently [4, 8]. For example, it is impossible to tune the cross-linking density of scaffolds formed from naturally derived materials

(e.g., collagen) without altering the density of biochemical cues (e.g., cell-adhesive binding sites). In order to evaluate the effect of biochemical properties (e.g., RGD concentration, enzyme-sensitive peptide substitution) on network properties of synthetic

PEG hydrogels, studies of mass swelling ratio as a function of gel compositions were

performed (Fig. 4.3). The swelling ratio of hydrogels is an important parameter that is

closely related to the biophysical properties of the hydrogel network (e.g., mesh size,

cross-linking density) [33]. Our results indicate that the hydrogel network is not affected by the inclusion of RGD-PEGMA while adhesive ligand concentration is lower than 2.5 mM for fixed VPM-PEGDA concentrations (Fig. 2A) [33]. This concentration is high relative to the concentration used to promote cell attachment (Fig. 4.5). Therefore, the method of grafting cell-adhesive peptide modified PEGMA into hydrogel network allows

for engineering cell-adhesive hydrogels with minimal altering of hydrogel network properties. The swelling ratio studies and further analysis of network properties (hydrogel mesh size and the molecular weight between cross-links) also indicate that single substitution of enzyme-sensitive peptide does not affect the swelling ratio of hydrogels

(VPM-PEGDA vs. GIA-PEGDA at the same concentration), while variation in VPM-

PEGDA or GIA-PEGDA concentrations results in a significant change in mass swelling ratios of the hydrogels, and thus network property (Table 4.2) [32, 33]. Therefore, by manipulation the composition of hydrogels, the effect of hydrogel properties, including biochemical and biophysical properties, on cell functions can be examined systematically.

4.5 Conclusion

Bio-inert PEG hydrogels can be engineered with the properties of cell adhesion and proteolytic degradation by the incorporation of cell-adhesive peptide (GRGDSP) and enzyme-sensitive peptide (GPQGIAGQ, VPMSMRGG) into the polymeric network. The cell-matrix adhesiveness of hydrogels can be modulated by the concentration of RGD ligand. When the concentration of adhesive ligands stays within a defined range (0-2.5

mM), hydrogels can be rendered cell-adhesive without affecting the network properties.

The degradability of hydrogels can be tuned by the change of enzyme-sensitive peptide type in the backbone of PEG chain, and single substitution of peptide type does not change hydrogel cross-linking density. Furthermore, the concentration of diacrylates (i.e.,

VPM-PEGDA or GIA-PEGDA) plays a major role in controlling the network properties

of hydrogels. This defined, biomimetic hydrogel can be developed as a 3D cell culture

model for the fundamental studies of SMC functions, such as SMC migration and

proliferation (Chapter 5 and Chapter 6).

4.6 Acknowledgement

The project described was supported by Grant Number 5RC1EB010795 and

Grant Number 1R01HL087843 for the National Heart, Lung, and Blood Institute. The

content is solely the responsibility of the authors and does not necessarily represent the

official views of the National Institutes of Health.

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CHAPTER 5 Smooth Muscle Cell Migration in 3D Bioactive PEG Hydrogels

Based on: Lin L, Zhu J, Kottke-Marchant K, Marchant RE. Biomimetic engineered poly

(ethylene glycol) hydrogel for smooth muscle cell migration. Tissue Eng Part

A 2014; 20: 864-73.

5.1 Introduction

Smooth muscle cell (SMC) migration plays a key role in a variety of

physiological and pathological situations, ranging from vascular development to intimal

hyperplasia following vascular injury [1-3]. During vascular development, migration of pericytes and smooth muscle precursor cells occurs following the formation of an endothelial cell tube, assisting in the development of vessel wall construction and biomechanical functionality of the blood vessels [2, 4, 5]. In response to vascular injury,

SMCs up-regulate the secretion of matrix metalloproteinases (MMPs) and increase their

rate of cell migration, which is required for wound healing and vascular repair [3, 6]. The development of materials to facilitate SMC migration has been a critical strategy in vascular tissue engineering because of the essential role of cell migration in vascular remodeling [7-9]. However, excessive SMC migration, followed by SMC proliferation, if uncontrolled, will induce pathogenic vascular remodeling, which is a key step in the development of intimal hyperplasia leading to vascular stenosis [1, 3, 6]. Therefore, understanding the mechanisms involved in SMC migration and development of strategies to regulate this process, have become emerging areas of research.

Published studies of SMC migration on two dimensional (2D) surfaces have suggested that cell migration is largely governed by the balance between attachment and detachment, presenting a biphasic dependence on cell-substratum adhesiveness [10, 11].

However, conditions for cell migration in vivo are more complex. Besides providing a

variety of biochemical cues to guide cell function, the extracellular matrix (ECM) also

imposes biophysical resistance to cell movement [12-15].

The objective of this project is to investigate 3D SMC migration in a defined,

bioactive PEG hydrogel. The bio-inert PEG hydrogels have been functionalized with the

properties of cell adhesion and proteolytic degradation as described in Chapter 4. By

utilizing this biomimetic hydrogel, we studied the effect of adhesive peptide

concentration, proteolysis, and network cross-linking density on 3D SMC migration.

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 Synthesis of Biomimetic Macromers

The cell-adhesive peptide (GRGDSP [RGD, MW: 586 Da]) and diaminopropionic

acid (Dap)-capped enzyme-sensitive peptides (VPMSMRGG-Dap [VPM-Dap, MW: 919

Da] and GPQGIAGQ-Dap [GIA-Dap, MW:812 Da]) were synthesized on an amide

(Knorr) resin using standard Fmoc chemistry on a solid phase peptide synthesizer

(Applied Biosystems, Model 433A, Foster City, CA). The peptides were cleaved from the

resin using trifluoroacetic acid and purified by reverse-phase high-performance liquid

chromatography (HPLC, Waters 2690 Alliance system).

The biomimetic macromers, RGD modified PEG monoacrylate (RGD-PEGMA)

and enzyme-sensitive peptide modified PEG diacrylates (VPM-PEGDA/GIA-PEGDA), were synthesized by conjugating peptides with Acrylate-PEG-Succinimidyl Valerate

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(Acr-PEG-SVA, MW~3400, Laysan Bio, Arab, AL) [16]. For the synthesis of RGD-

PEGMA, Acr-PEG-SVA (added dropwise, 30-40 mg/ml) was reacted with GRGDSP

(15% molar excess) in aqueous sodium bicarbonate solution at room temperature (RT) for at least 4 h. Then, the mixture was dialyzed against water with membranes of

Molecular Weight Cut Off (MWCO) 2,000 Da for 48 h, to remove salts and unreacted peptides. The purified product was lyophilized and stored at -20 ºC. VPM-PEGDA/GIA-

PEGDA was synthesized by the same method as RGD-PEGMA using Acr-PEG-SVA with VPM-Dap or GIA-Dap peptide in a molar ratio of 2:1(PEG: peptide). The final product was dialyzed against water with membranes of MWCO 5,000 for 48 h. The synthesis of biomimetic macromers was confirmed by matrix assisted laser desorption/ionization mass spectroscopy (MALDI-MS).

5.2.3 Cell Culture

Human coronary artery smooth muscle cells (HCASMCs, Lonza, Walkersville,

MD) were maintained at 37 ºC, 5% CO2 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. For all experiments, HCASMCs were used at passages P8-P11.

5.2.4 Gelatin Zymography

To detect enzyme secretion from HCASMCs, cells were seeded at various densities (0-6×104 cells/cm2) in a 48-well culture plate in SmGM-2 growth medium.

After 24 h, the culture medium was collected and analyzed for enzyme activity by gelatin zymography. One part of conditioned medium was mixed with two parts of zymography sample buffer (1×, Bio-Rad, Hercules, CA) and incubated at RT for 10 min. The resulting

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mixed samples were loaded onto a 10% precast polyacrylamide gel with gelatin (Bio-

Rad). Recombinant human MMP-2 (10 ng, R&D, Minneapolis, MN) was used as a

standard. Gel electrophoresis was performed in the Tris-Glycine Sodium dodecyl sulfate

(SDS) Running Buffer at 125V until the dye front reached the bottom of the gel. After

electrophoresis, the gels were incubated in zymography renaturing buffer (Triton-X-100,

2.5% [v/v] in water) with gentle agitation for 60 min to remove SDS. After decanting the

renaturing buffer, gels were incubated in zymography development buffer (Bio-Rad) at

37 ºC overnight. The gels were then stained with 0.1% (w/v) Coomassie Brilliant Blue R-

250 solution and finally destained with destaining solution (methanol: acetic acid: water

= 50: 5: 45). Enzymatic activity was visualized by clear bands against a dark blue

background. Gel samples were imaged and band intensity of each sample was quantified

by Image J analysis.

5.2.5 3D Cell Invasion

A modified 3D cell invasion method was used to evaluate cell migration in 3D

[17, 18]. Generally, a sandwich-like gel was prepared by embedding a cell pellet loaded

gel between two layers of gel (70 µl for each layer) with the same composition. Hydrogel

precursor solutions contained RGD-PEGMA (0-5 mM), VPM/GIA-PEGDA (5-7%,

w/w), and 0.1% w/v Irgacure 2959 (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), dispensed into a sterilized stainless steel mold, and polymerized for 5 min under UV light to form the bottom layer of hydrogel (D=10 mm). To prepare the middle layer of cell pellet loaded gel, 5 µl of cell suspension (1.5×107 cells/ml

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matrigel) was applied onto a separate glass plate mold (a glass plate covered with a

silicone sheet with punched circles). After 5 min of gelation, hydrogel precursor solution

(50 µl) was added into the glass plate mold, covered with a cover slip, and polymerized

for 5 min under UV light. Then, the cell pellet loaded gel was removed from the glass

plate mold and placed on top of the hydrogel layer in the stainless steel mold. Excess

precursor solution was spread evenly on the top of the cell pellet loaded gel, and placed

under a UV light for 10 min to form the top layer of hydrogel. Once completely

polymerized, gel samples (H=1.2 mm) were then transferred to a 12-well culture plate

and cultured in SmGM-2 growth medium for up to 14 d. The medium was changed every

2 d. At pre-determined culture times, cells were imaged by inverted phase contrast

microscopy (10×). Digital images were assembled together in Adobe Photoshop for

quantification [17]. The assembled images were aligned with a grid evenly divided into

36 intervals. The radius of the sample was determined by averaging the distance between

the center and the furthest point on each grid line with an invading chain of cells. The

migration distance was obtained by subtracting the radius of the initial cell pellet from the

radius of the sample.

5.2.6 Statistics

Statistical analysis was done using Origin 8.0 and Minitab 1.6. Data are represented as mean ± standard deviation of at least triplicate samples. 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. A value of p<0.05 was considered statistically significant.

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5.3 Results

5.3.1 Enzyme Detection

To assess the ability of SMCs to degrade the hydrogels, the presence of enzyme in

the conditioned medium from cell culture was determined by gelatin zymography (Fig.

5.1). By a comparison of sample bands with a molecular weight marker and a MMP-2

standard, it was confirmed that the SMCs expressed MMP-2 (Fig. 5.1A) and the amount

of MMP-2 expression increased with increasing cell number (Fig. 5.2B). The in vitro degradation experiments showed that both VPM-PEGDA and GIA-PEGDA hydrogels can be degraded completely by MMP-2 solution (see Chapter 4, Fig. 4.4).

5.3.1 Cell Migration in the Hydrogel

A modified 3D cell invasion method was used to investigate SMC migration

behavior in 3D biomimetic PEG hydrogels [17, 18]. SMCs migrated in the GIA-PEGDA

(5%, w/w) hydrogels with 5 mM of RGD-PEGMA (Fig. 5.2). Initially, the SMC pellet

was embedded in the hydrogel disks (Fig. 5.2A). Then, SMCs migrated from the pellet

into surrounding hydrogels (Fig. 5.2A). The migration distance increased significantly

with culture time (Fig. 5.2B).

5.3.2 The Effect of RGD Concentration on Cell Migration

The effect of cell adhesive ligand concentration on cell migration was

investigated by comparing the migration distance of SMCs after 7 d in GIA-PEGDA

(5%, w/w) hydrogels as a function of RGD-PEGMA concentration (0-2.5 mM in the

precursor solution). Fig. 5.3 shows the RGD ligand concentration mediated SMC migration in a biphasic manner. SMC migration was significantly enhanced in 3D gels with a RGD concentration of 0.625 mM, compared with other tested concentrations. For

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Figure 5. 1 Enzyme detection in conditioned medium (after a 24 h seeding period in SmGM-2 growth medium) by zymography. (A) Zymography image: Lane 1: MMP-2 standard (10 ng); Lane 2-5: conditioned medium with a series of seeding density (Lane 2: 0×104 cells/cm2; Lane 3: 1.5×104 cells/cm2; Lane 4: 3.0×104 cells/cm2; Lane 5: 6.0×104 cells/cm2) (B) Quantification of enzyme secretion by measuring band intensity reported relative to a MMP-2 standard (10 ng). *: p<0.05 with respect to Lane 2: 0×104 cells/cm2.

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Figure 5. 2 SMC migration in 3D hydrogels (GIA-PEGDA: 5%, w/w; RGD-PEGMA: 5 mM). (A) Phase contrast micrographs of cell migration in 3D gels on day 1, 7 and 14. (B) Quantification of migration distance by Image J.

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Figure 5. 3 The effect of cell adhesive ligand (RGD) concentration on SMC migration (on day 7) in 3D hydrogels (GIA-PEGDA: 5%, w/w; RGD-PEGMA: 0-2.5 mM).

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hydrogels without RGD incorporation or with low adhesive ligand concentration (≤0.025 mM), cell migration was absent. Cell migration in GIA-PEGDA (5%, w/w) hydrogels with various concentrations of RDG-PEGMA (synthesized by the same method as RGD-

PEGMA using the reaction of Acr-PEG-SVA with the scrambled peptide: GRDGSP) (0-

2.5 mM) also was determined. Again, no migration was observed in RDG modified

hydrogels (Data not shown).

5.3.3 The Effect of Proteolysis on Cell Migration

To determine the effect of proteolysis on cell migration, VPM-PEGDA, GIA-

PEGDA and IGA-PEGDA (synthesized by the same method as VPM/GIA-PEGDA using

the reaction of Acr-PEG-SVA with the scrambled peptide: GPQIGAGQ-Dap) (5%, w/w)

hydrogels with a constant RGD ligand concentration (0.625 mM in the precursor solution) were utilized (Fig. 5.4). For hydrogels that were enzyme-sensitive, after 7 d, cells were able to migrate from the pellet into the surrounding gel (VPM-PEGDA and

GIA-PEGDA) (Fig. 5.4A and Fig. 5.4B). As expected, SMCs migrated faster in VPM-

PEGDA hydrogels than in GIA-PEGDA hydrogels (Fig. 5.4D). In contrast, little

migration was observed in IGA-PEGDA hydrogels (Fig. 5.4C and Fig. 5.4D), which are

not sensitive to enzyme degradation.

5.3.4 The Effect of Network Cross-linking Density on Cell Migration

Cell migration into 3D gels with various concentrations of GIA-PEGDA (5%-7%,

w/w) was evaluated at a constant RGD ligand concentration of 0.441 mM in the swollen gel (achieved by adjusting the RGD concentration in the precursor solution by correcting for the differences in swelling of the hydrogels) (Fig. 5.5). By increasing the GIA-

PEGDA concentration from 5% to 6%, migration distances decreased significantly from

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Figure 5. 4 The effect of proteolysis on SMC migration (on day 7) in 3D hydrogels. (A) Cell migration in 5% VPM-PEGDA hydrogels with 5 mM RGD-PEGMA. (B) Cell migration in 5% GIA-PEGDA hydrogels with 5 mM RGD-PEGMA. (C) Cell migration in 5% IGA-PEGDA hydrogels with 5 mM RGD-PEGMA. (D) Quantification of migration distance by Image J.

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Figure 5. 5 The effect of network cross-linking density on SMC migration (on day 7) in 3D hydrogels (GIA-PEGDA: 5%-7%, RGD-PEGMA: 0.441 mM in all the swollen gels).

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1688 µm to 378 µm after 7 days. At higher concentrations of GIA-PEGDA (7%, w/w), cell migration was restricted completely.

5.4 Discussion

The role of SMC migration as an essential process in physiological and

pathological vessel wall remodeling makes the study of mechanisms involved in cell

migration a major focus of research [1-3]. In contrast to cell migration on 2D surfaces,

3D cell migration is more complex, because migration is not only mediated by

biochemical factors (e.g. adhesive ligand concentration), but also by biophysical factors

(e.g., network cross-linking density) [12-15].

In this project, a modified 3D cell invasion method was used to investigate SMC migration behavior in 3D biomimetic PEG hydrogels [17, 18]. A pellet of SMCs was embedded in the hydrogel, and SMC outgrowth from the pellet to the surrounding hydrogel was measured. SMC proliferation in the surrounding hydrogel may contribute to the cell number and migration distance. Techniques to separate migration from proliferation, such as mitomyosin C (MMC) to inhibit SMC proliferation in 3D cell invasion experiment have been described [17]. The results showed that MMC treatment attenuated the distance of SMC invasion compared to untreated controls. However, there are concerns that MMC may affect secretion of MMPs or inhibitors [19], and thereafter

affect SMC migration. The observations from our experiments (Fig. 5.2A) clearly show

that SMCs migrated from the pellet into surrounding hydrogel. With culture time

extended, SMCs migrated further (Fig. 5.2). This demonstrates the ability of SMCs to

migrate in this biomimetic hydrogel.

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Cell-matrix adhesion is a governing parameter of cell migration on 2D surfaces

[10], consequently it is reasonable to anticipate that cell-matrix adhesion also will play a key role in 3D cell migration. To explore the effect of a single parameter (e.g., adhesive ligand concentration) on cell migration in a 3D model, the interdependence of variables

(e.g. adhesive ligand concentration vs. hydrogel network property) must be considered.

To investigate the effect of adhesive ligand concentration on the hydrogel network, studies of mass swelling ratio as a function of RGD-PEGMA concentration were performed (See chapter 4, Fig. 4.3). The results indicated that the hydrogel network was not affected by the inclusion of RGD-PEGMA while adhesive ligand concentration was in the range of 0-2.5 mM. Therefore, this concentration range (0-2.5 mM) was chosen to study the effect of adhesive ligand concentration on SMC migration. Similar to previous studies of cell migration on 2D surfaces [10, 20] and within 3D matrices [11, 18, 21, 22], a biphasic relationship between migration distances and cell-matrix adhesiveness was found (Fig. 5.3). As cell migration is a product of the net force between counteracting detachment and adhesion forces, it is hypothesized that at low ligand concentration, weak cell-matrix adhesiveness results in a decrease in traction forces for forward movement, which subsequently slows cell migration. In contrast, in the presence of a high ligand concentration in the 3D network, strong cell-matrix adhesiveness inhibits cell detachment, which also results in decreased migration [11, 18].

In a 3D matrix, cells must not only interact with matrix adhesive ligands for force generation, but also develop additional strategies to overcome the biophysical resistance of the scaffold [12-15]. Depending on the type of cell and matrix, either mechanistic strategy of changing cell morphology and/or matrix degradation are utilized by cells to

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overcome ECM resistance [12-15]. Since the mesh size of the synthetic 3D scaffold (see

Chapter 4, Table 4.2) is much smaller than the dimensions of SMCs, we hypothesize that

proteolysis is required for SMC migration to occur. To investigate this hypothesis, we evaluated the behavior of cell migration in hydrogels that have different sensitivity to

MMP degradation [23]. For hydrogels that were sensitive to MMP degradation (VPM-

PEGDA and GIA-PEGDA hydrogels), cell migration was observed and invading cells assumed a spindle-like shape (Fig. 5.4A and Fig. 5.4B). This is markedly different from the well-spread cell morphology found on the surfaces of GIA-PEGDA hydrogels with the same concentration of RGD-PEGMA [18, 24, 25] (see Chapter 4, Fig. 4.5A).

Furthermore, SMCs in VPM-PEGDA hydrogels migrated faster than cells in GIA-

PEGDA hydrogels, which is suggested to be related with the enzyme sensitivity of incorporated peptide [23, 26]. After the single substitution of an enzyme-sensitive peptide to an enzyme insensitive peptide (GPQIGAGQ) in the PEG backbone, little cell

migration was observed (Fig. 5.4C). A similar inhibition was observed in degradable

matrices when an MMP inhibitor (e.g. GM6001, TIMP-2) was added to the culture

medium, which confirms that cell migration in 3D hydrogels depends critically on the

action of proteolysis by cell-secreted MMPs [18, 21, 27]. Additionally, the incorporated

degradable peptides with different proteinase sensitivity were reported to affect the RGD

concentration for maximal cell migration [11]. The synergistic effect of proteolysis and

adhesive ligand concentration on cell migration remains to be further investigated.

The effect of the hydrogel network on cell migration in 3D gels was evaluated by varying the concentration of GIA-PEGDA in the hydrogels. Mass swelling ratio studies showed that variation in GIA-PEGDA concentrations results in a significant change in

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mass swelling ratios of the hydrogels (see Chapter 4, Fig. 4.3B), which is an important

parameter that is closely related to the biophysical properties of the hydrogel network

(e.g., mesh size, cross-linking density) [28-30]. At a constant concentration of RGD-

PEGMA (0.441 mM, the concentration which shows maximum migration in 5% GIA-

PEGDA gels) in the swollen hydrogel, increasing GIA-PEGDA concentrations

significantly decreases migration distances (Fig. 5.5). This observation is consistent with

previous studies, in which the network properties of PEG hydrogels were tuned by PEG

molecular weight [18]. Therefore, along with cell-matrix adhesiveness and proteolysis,

network cross-linking density plays a critical role in 3D cell migration.

Through characterization of SMC migration in hydrogels with different compositions, we have identified that gels fabricated from high MMP sensitivity, low cross-linking density and an intermediate RGD ligand concentration (0.625 mM RGD-

PEGMA in the precursor solution for 5% PEGDA gel) present an optimal construct to promote SMC migration. This can be utilized to facilitate SMC migration in initial vascular remodeling of tissue engineered blood vessels (TEBVs) [7-9]. In order to regulate 3D SMC migration in lateral stage to prevent failure mechanisms (e.g., intimal

hyperplasia), the spatiotemporal control of biochemical and physical properties of

hydrogels in 3D and in time remains to be further investigated [31].

5.5 Conclusion

Bio-inert PEG gels can be rendered cell adhesive and biodegradable by the

incorporation of cell adhesive peptide (GRGDSP) and collagenase sensitive peptide

(GPQGIAGQ) into the polymeric network. By utilizing this biomimetic scaffold, 3D

SMC migration behavior was investigated in which RGD ligand concentration, MMP

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sensitivity, and network cross-linking density, were varied systematically. Our results indicate that 3D SMC migration was critically dependent on cell-matrix adhesiveness and proteolysis, and cell adhesive ligand mediated cell migration in a biphasic manner.

Further, the network cross-linking density of 3D hydrogels also plays a key role in SMC migration in 3D. Besides cell migration, these biomimetic scaffolds offer a useful tool in the fundamental studies of SMC functions (e.g. cell proliferation) in three dimensions, which will facilitate the development of functional tissue-engineered blood vessels.

5.6 Acknowledgement

The project described was supported by Grant Number 5RC1EB010795 and

Grant Number 1R01HL087843 for the National Heart, Lung, and Blood Institute. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

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5.7 References

1. Willis AI, Pierre-Paul D, Sumpio BE, Gahtan V. Vascular smooth muscle cell migration: current research and clinical implications. Vascular and endovascular surgery 2004;38:11-23.

2. Gerthoffer WT. Mechanisms of Vascular Smooth Muscle Cell Migration. Circulation Research 2007;100:607-21.

3. Louis SF, Zahradka P. Vascular smooth muscle cell motility: From migration to invasion. Experimental and clinical cardiology 2010;15:e75-85.

4. Bergers G, Song S. The role of pericytes in blood-vessel formation and maintenance. Neuro-oncology 2005;7:452-64.

5. Campbell GR, Campbell JH. Development of the Vessel Wall: Overview. In: Schwartz SM, Mecham RP, editors. The Vascular Smooth Muscle Cell. San Diego: Academic Press; 1995. p. 1-15.

6. Schwartz SM. Smooth muscle migration in atherosclerosis and restenosis. The Journal of clinical investigation 1997;100:S87-9.

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;22:3045-51.

8. Almany L, Seliktar D. Biosynthetic hydrogel scaffolds made from fibrinogen and polyethylene glycol for 3D cell cultures. Biomaterials 2005;26:2467-77.

9. Liu Y, Chan-Park MB. A biomimetic hydrogel based on methacrylated dextran-graft- lysine and gelatin for 3D smooth muscle cell culture. Biomaterials 2010;31:1158-70.

10. DiMilla PA, Stone JA, Quinn JA, Albelda SM, Lauffenburger DA. Maximal migration of human smooth muscle cells on fibronectin and type IV collagen occurs at an intermediate attachment strength. The Journal of cell biology 1993;122:729-37.

11. Gobin AS, West JL. Cell migration through defined, synthetic ECM analogs. FASEB journal : official publication of the Federation of American Societies for Experimental Biology 2002;16:751-3.

12. Friedl P, Brocker EB. The biology of cell locomotion within three-dimensional extracellular matrix. Cellular and molecular life sciences : CMLS 2000;57:41-64.

13. Friedl P, Zanker KS, Brocker EB. Cell migration strategies in 3-D extracellular matrix: differences in morphology, cell matrix interactions, and integrin function. Microscopy research and technique 1998;43:369-78.

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14. Even-Ram S, Yamada KM. Cell migration in 3D matrix. Current opinion in cell biology 2005;17:524-32.

15. Baumann K. Cell migration: Switching to 3D. Nat Rev Mol Cell Biol 2012;13:338-9.

16. Zhu J, He P, Lin L, Jones DR, Marchant RE. Biomimetic poly(ethylene glycol)- based hydrogels as scaffolds for inducing endothelial adhesion and capillary-like network formation. Biomacromolecules 2012;13:706-13.

17. Ucuzian AA, Brewster LP, East AT, Pang Y, Gassman AA, Greisler HP. Characterization of the chemotactic and mitogenic response of SMCs to PDGF-BB and FGF-2 in fibrin hydrogels. Journal of biomedical materials research Part A 2010;94:988- 96.

18. 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. Proceedings of the National Academy of Sciences of the United States of America 2003;100:5413-8.

19. Hamner MA, Vernon RB, Gooden MD, Koike T, Reed MJ. Elongation and secretion of tissue inhibitor of metalloproteinases 1 by human microvascular endothelial cells cultured in collagen gels is stimulated by mitomycin c. Endothelium : journal of endothelial cell research 2005;12:97-101.

20. Wu J, Mao Z, Gao C. Controlling the migration behaviors of vascular smooth muscle cells by methoxy poly(ethylene glycol) brushes of different molecular weight and density. Biomaterials 2012;33:810-20.

21. Raeber GP, Lutolf MP, Hubbell JA. Mechanisms of 3-D migration and matrix remodeling of fibroblasts within artificial ECMs. Acta biomaterialia 2007;3:615-29.

22. Raeber GP, Lutolf MP, Hubbell JA. Molecularly engineered PEG hydrogels: a novel model system for proteolytically mediated cell migration. Biophysical journal 2005;89:1374-88.

23. Patterson J, Hubbell JA. Enhanced proteolytic degradation of molecularly engineered PEG hydrogels in response to MMP-1 and MMP-2. Biomaterials 2010;31:7836-45.

24. Beamish JA, Fu AY, Choi AJ, 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;30:4127-35.

25. 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. Journal of biomedical materials research 2002;60:86-93.

26. Patterson J, Hubbell JA. SPARC-derived protease substrates to enhance the plasmin sensitivity of molecularly engineered PEG hydrogels. Biomaterials 2011;32:1301-10.

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27. Cheng L, Mantile G, Pauly R, Nater C, Felici A, Monticone R, Bilato C, Gluzband YA, Crow MT, Stetler-Stevenson W, Capogrossi MC. Adenovirus-mediated gene transfer of the human tissue inhibitor of metalloproteinase-2 blocks vascular smooth muscle cell invasiveness in vitro and modulates neointimal development in vivo. Circulation 1998;98:2195-201.

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. Journal of biomedical materials research Part A 2010;92:441-50.

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;4:713-22.

30. Munoz-Pinto DJ, Bulick AS, Hahn MS. Uncoupled investigation of scaffold modulus and mesh size on smooth muscle cell behavior. Journal of biomedical materials research Part A 2009;90:303-16.

31. DeForest CA, Anseth KS. Advances in bioactive hydrogels to probe and direct cell fate. Annual review of chemical and biomolecular engineering 2012;3:421-44.

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CHAPTER 6 Smooth Muscle Cell Proliferation in 3D Bioactive PEG Hydrogels

6.1 Introduction

Cardiovascular disease (CVD), including coronary heart disease, stroke, heart failure and peripheral arterial disease, is the leading cause of mortality in the United

States [1]. The use of autologous bypass grafts, such as saphenous vein, remains one of the mainstays of treatment for advanced CVD. Due to the limited availability of suitable autologous vessels, tissue engineered blood vessels (TEBVs) have emerged as an attractive alternative for bypass grafts [2-5]. In the development of TEBVs, smooth muscle cell (SMC) proliferation, along with deposition of extracellular matrix (ECM) in vascular media, is necessary for vessel wall construction and biomechanical functionality as blood conduits [2, 6, 7]. Development of strategies to facilitate SMC proliferation in initial vascular remodeling of TEBVs has been critical for vascular tissue engineering [2,

6]. However, excessive SMC proliferation, if uncontrolled, will promote the development of intimal hyperplasia, which is one of the major failure mechanisms for current synthetic vascular grafts [2, 6, 8]. Therefore, exploration of the mechanisms involved in SMC proliferation and the strategies to regulate this function, is critical for development of functional tissue engineered vascular grafts.

SMC proliferation is mediated by various signals, including mechanical stimuli,

ECM proteins, and soluble bioactive factors [6, 8-10]. It has been reported that the application of cyclic mechanical strain to cultured SMCs induces SMC proliferation and

ECM production [11, 12]. The ECM regulates SMC proliferation through cell-surface receptors, which transduce signals between cells and the ECM [10, 13, 14]. Furthermore, a variety of soluble signaling factors have been implicated to affect SMC proliferation

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[15, 16]. Heparin, for example, has been well described to reduce SMC proliferation in

vitro and in vivo [16-18]. The major structural determinant of heparin antiproliferative

activity is the amount and distribution of sulfonate groups on the glycosaminoglycan

chain [19]. Heparin has been shown to be able to inhibit SMC proliferation induced by

many stimulatory signals, such as serum [20-22] and basic fibroblast growth factor

(bFGF) [23, 24]. However, there are inconsistencies on the effect of heparin in inhibiting other stimuli, such as platelet-derived growth factor (PDGF) [6, 23-25]. In general, these

studies have focused on proliferation of SMCs in 2 dimensional (2D) culture. Cells in

vivo reside in a 3 dimensional (3D) microenvironment, and there is a growing

appreciation that cells may respond differently when cultured in 3D versus 2D systems

[26-28]. Thus, it is of great importance to investigate SMC functions in 3D cell culture

models.

In this report, our goal is to examine the effect of scaffold properties and exogenous bioactive factors on SMC proliferation in a 3D ECM-mimetic PEG hydrogel.

SMC functions, such as growth and ECM production, have been studied by homogeneous

seeding of SMCs into 3D PEG gels [29-31]. However, cells encapsulated in these gels

often present a round morphology instead of a normal spindle-like morphology. This might be due to the lack of fast degradation mechanisms in these gels [29, 31]. The

absence of SMC spreading in these 3D constructs might affect SMC viability and

proliferation and hamper investigation of the effect of both scaffold properties and

exogenous bioactive factors on SMC proliferation [29, 31, 32]. VPMSMRGG, a matrix

metalloproteinase (MMP) sensitive peptide derived from a peptide library, has been reported to have higher MMP sensitivity than other naturally derived peptides [33], such

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as GPQGIAGQ derived from collagen type I [34]. In this study, enzyme-sensitive

peptides, including VPMSMRGG and GPQGIAGQ, were incorporated into the PEG

chain, respectively. By photopolymerization, the MMP sensitive peptide modified PEG

diacrylate was copolymerized with cell adhesive peptide (GRGDSP derived from

fibronectin [35]) modified PEG monoacrylate to form a bioactive hydrogel with the

ECM-mimetic properties of cell adhesion and proteolytic degradation. The scaffold properties (e.g. RGD concentration, proteolysis, network crosslinking density) of the hydrogels can be controlled quantitatively and independently [36], which allows systematic examination of these properties on SMC proliferation. Furthermore, the antiproliferative effect of heparin, which has been studied extensively on 2D surfaces

[16-18] but limited in 3D system [32], was examined as an exogenous biochemical factor in this 3D scaffold.

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 Biomimetic Macromers

The cell adhesive peptide (GRGDSP [RGD]) and diaminopropionic acid (Dap)-

capped MMP-sensitive peptides (VPMSMRGG-Dap [VPM-Dap] and GPQGIAGQ-Dap

[GIA-Dap]) were synthesized on an amide (Knorr) resin using standard Fmoc chemistry

on a solid phase peptide synthesizer (Applied Biosystems, Model 433A, Foster City,

CA). The peptides were cleaved from the resin using trifluoroacetic acid and purified by

reverse-phase high-performance liquid chromatography (Waters 2690 Alliance system).

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Successful peptide synthesis was confirmed by matrix assisted laser desorption/ionization

mass spectroscopy (MALDI-MS).

Biomimetic macromers were synthesized by conjugating bioactive peptides with

Acrylate-PEG-Succinimidyl Valerate (Acr-PEG-SVA, MW: 3400 Da; Laysan Bio, Arab,

AL) through the coupling reaction between the primary amine on peptides and the N- hydroxysuccinimide (NHS) group on Acr-PEG-SVA, as described previously [36]. To modify PEG with cell adhesive peptide, Acr-PEG-SVA was reacted with GRGDSP (15% molar excess) in aqueous sodium bicarbonate solution at room temperature (RT) for at least 4 h. Then the product of RGD modified PEG monoacrylate (RGD-PEGMA) was purified by dialysis, lyophilized and stored at -20 °C until use. To conjugate one molecule of MMP-sensitive peptide with two molecules of Acr-PEG-SVA, the C- terminus of VPM and GIA peptide was capped with diaminopropionic acid to provide an additional primary amine [36, 37]. The MMP-sensitive peptide modified PEG diacrylates

(VPM-PEGDA/GIA-PEGDA) were then synthesized by reacting Acr-PEG-SVA with

VPM-Dap or GIA-Dap peptide in a molar ratio of 2:1 (PEG: peptide), respectively. The synthesis of biomimetic macromers was confirmed by MALDI-MS.

6.2.3 Cell Culture

Human coronary artery SMCs (HCASMCs; Lonza, Walkersville, MD) were

maintained at 37 ºC, 5% CO2 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. For all

experiments, HCASMCs were used at passages P5-P8.

6.2.4 Cell Encapsulation within Hydrogels

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To assess the 3D proliferation and spreading of SMCs within the bioactive PEG hydrogels, a sandwich-like gel was prepared by embedding a cell encapsulated gel between two layers of gel (30 µl for each layer) with the same polymer composition. This was to avoid the initial cell loss from the gel surface. Briefly, hydrogel precursor solutions were prepared by dissolving RGD-PEGMA (0-5 mM), VPM/GIA-PEGDA (4-

6%, w/w), and 0.1% w/v Irgacure 2959 (1-[4-[2-Hydroxyethoxy]-phenyl]-2-hydroxy-2- methyl-1-propane-1-one, Ciba Specialty Chemicals, Tarrytown, NY) in phosphate buffered saline (PBS, pH 7.4). The resulting solution was sterilized by filtration (0.22 µm pore), dispensed into a sterilized stainless steel mold, and polymerized for 8 min under

UV light to form the bottom layer of hydrogel (D=6 mm). To prepare the middle layer of cell encapsulated gel, HCASMCs were mixed with the precursor solution (density:

1.25×105 cells/ml, if not otherwise specified) immediately before hydrogel formation.

The cell suspension was applied onto the bottom layer and polymerized for 8 min under

UV light. Then excess precursor solution was spread evenly on the top of the cell

encapsulated gel and placed under a UV light for 8 min to form the top layer of hydrogel.

Once completely polymerized, gel samples (H= 2.4 mm ) were then transferred to a 24-

well culture plate and cultured in SmGM-2 growth medium for up to 3 weeks. The

medium was changed every 2 days.

To assess the effect of heparin on SMC proliferation and spreading in 3D PEG

hydrogels, cell encapsulated gels were prepared as described above and cultured in

SmGM-2 growth medium with increasing concentrations of heparin solution (0-2

mg/ml). To compare the effect of heparin on SMC proliferation on 2D surfaces, SMCs

were seeded on human fibronectin (FN, 1µg/cm2)-coated tissue culture polystyrene

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surfaces at the density of 5000 cells/cm2 and cultured in SmGM-2 growth medium with varied concentrations of heparin solution (0-2 mg/ml).

6.2.5 Cell Viability in 3D Hydrogels

Cell viability after incorporation into 3D bioactive hydrogels by

photopolymerization was determined by live/dead cell viability kit (Life Technologies,

Grand Island, NY). HCASMCs were homogeneously seeded into bioactive PEG

hydrogels at a density of 2.5×105 cells/ml as described previously. Cell encapsulated

hydrogels were then transferred to a 24-well culture plate and incubated in SmGM-2

growth medium. After incubation for 24 h after seeding, the samples were washed gently

with PBS and stained in 500 µl of live/dead stain solution (2 µM calcein AM, 4 µM ethidium homodimer-1 in PBS) at RT for at least 30 min. Subsequently, a Nikon Diphot

100 inverted microscope (Melville, NY) with a 10× objective was used to visualize the green living cells (stained with calcein AM) and the red dead cells (stained with ethidium homodimer-1) in the samples.

6.2.6 Cell Morphology in 3D Hydrogels

For examination of cell morphology, cell encapsulated hydrogels were fabricated

as described above and cultured in SmGM-2 growth medium for up to 3 weeks. At

predetermined time intervals, the samples were washed three times with PBS and treated

sequentially with 4% paraformaldehyde PBS-solution and 0.1% Triton X-100 at RT for

30 min to fix and permeabilize cells. After washing three times with PBS, cytoskeletal F- actin fibers were stained with red fluorescence by incubation with 5 units/mL Alexa

Fluor 568 Phalloidin (Life Technologies) in PBS containing 1% bovine serum albumin

(BSA) at RT for 1 h. Cell nuclei were counterstained with 0.067 mM 4’,6-diamino-2-

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phenylinodole (DAPI; Life Technologies) (blue fluorescence). Subsequently, samples were washed in fresh PBS to remove excess dyes, followed by imaging with a ZEISS

LSM 510 META confocal microscope (Carl Zeiss Microscopy, Thornwood, NY) with 10

× magnification. Z series of approximately 30 equidistant x-y scans at 7 µm intervals were acquired and projected onto a single plane using the software provided by the system.

6.2.7 Cell Proliferation in 3D Hydrogels

Cell proliferation in 3D hydrogels was determined by quantification of DNA content of cell encapsulated gels as previously described [31]. At predetermined time intervals, hydrogels were removed from the culture medium, washed gently with PBS, weighed and then frozen at -80°C. At the time of analysis, hydrogel samples were digested with 0.1 M NaOH (1 ml per 0.2 g hydrogel wet weight) overnight at 37°C.

Digested hydrogels were then neutralized with HCl. DNA content of the digested, neutralized hydrogels was determined using PicoGreen assay (Life Technologies) [36].

Calf thymus DNA served as a standard and was subjected to the same association with hydrogel samples, including incubation with 0.1 M NaOH overnight and then neutralization with HCl. The proliferation data were expressed as the fold change of

DNA, representing the ratio between the DNA content per gel measured at different time points and the DNA content at day 1.

6.3 Results

6.3.1 Cell Viability

To demonstrate that cells remained viable after photopolymerization, HCASMCs were homogeneously seeded in the bioactive hydrogels (VPM-PEGDA: 5%, w/w, RGD-

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PEGMA: 1.25 mM) and were examined at 6 h after gel formation. Live cells, indicated

by green fluorescent staining with calcein AM, were evident after photopolymerization

(Fig. 6.1). HCASMCs viability was greater than 95% in the hydrogels.

6.3.2 The Effect of RGD Concentration on Cell Morphology and Proliferation

The effect of cell adhesive ligand concentration on HCASMCs morphology and

proliferation (at day 21) was assessed by utilizing VPM-PEGDA (4%, w/w) hydrogels

with varying concentrations of RGD-PEGMA (0-2.5 mM) (Fig. 6.2). In the absence of

RGD incorporation, encapsulated SMCs presented round morphologies (Fig. 6.2A). By

including RGD-PEGMA into the hydrogels, cells started to elongate and adopted a spindle-shaped morphology (Fig. 6.2A). DNA quantification showed that RGD ligand concentration mediated SMC proliferation in a biphasic manner (Fig. 6.2B). The incorporation of 0.625 and 1.25 mM RGD-PEGMA significantly enhanced cell proliferation in 3D gels, compared with other tested concentrations (Fig. 6.2B). For hydrogels without RGD incorporation or with low adhesive ligand concentration (≤ 0.125

mM), minimal or no cell proliferation occurred (Fig. 6.2B).

6.3.3 The Effect of MMP Sensitivity on Cell Morphology and Proliferation

The effect of MMP sensitivity on 3D SMC morphology and proliferation was

investigated using VPM-PEGDA, GIA-PEGDA and IGA-PEGDA (4%, w/w) hydrogels

with a constant RGD ligand concentration (1.25 mM in the precursor solution). On Day 1

after cell encapsulation into hydrogels, all the hydrogels showed a rather uniform

distribution of cells. For hydrogels modified with MMP-insensitive peptide

(GPQIGAGQ), encapsulated SMCs predominantly remained round during a culture

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Figure 6. 1 Cell viability in 3D bioactive PEG-based hydrogels (VPM-PEGDA: 4%, w/w; RGD-PEGMA: 1.25 mM) at 6 h after gel formation. Live cells were stained green while dead cells were stained red. The arrow indicates a representative dead cell.

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Figure 6. 2 The effect of cell adhesive ligand (RGD-PEGMA) concentration on SMC morphology and proliferation (at day 21) in 3D functionalized PEG-based hydrogels (VPM-PEGDA: 4%, w/w; RGD-PEGMA: 0-2.5 mM). (A) Confocal laser scanning micrographs of cell morphology in RGD modified hydrogels. Cells were double-stained with Alexa Fluor 568 phalloidin (F-actin filaments, red) and DAPI (nuclei, blue). (B) SMC proliferation in 3D RGD modified hydrogels. Proliferation was quantified by measuring DNA content using PicoGreen assay and reported relative to the DNA content at Day 1. *: p<0.05 with regard to hydrogels with 0.625 mM RGD; #: p<0.05 with regard to hydrogels with 1.25 mM RGD.

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period of 21 days. Limited cell spreading (indicated by arrows in Fig. 6.3) was observed

in GIA-PEGDA hydrogels. For VPM-PEGDA hydrogels, cells started to elongate at early

time points (Day 7), with increased cell spreading as well as cell proliferation observed by Day 21 (Fig. 6.3). Similar to the behavior of cell spreading, the fold change in DNA

content, representing cell proliferation, was significantly increased in VPM-PEGDA

hydrogel after 2 and 3 wk (Fig. 6.4). In contrast, little cell proliferation occurred within

GIA-PEGDA and IGA-PEGDA hydrogels (Fig. 6.4).

6.3.5 The Effect of Cross-linking Density on Cell Morphology and Proliferation

SMCs morphology and proliferation in 3D gels with various concentrations of

VPM-PEGDA (4-6%, w/w) were evaluated while maintaining a constant RGD ligand

concentration of 0.502 mM in the swollen gel (achieved by adjusting the RGD

concentration in the precursor solution by correcting for the differences in swelling of the

hydrogels). Increasing the VPM-PEGDA concentration from 4% to 6%, resulted in less

cell spreading (Fig. 6.3). On day 21, both 4% and 5% VPM-PEGDA hydrogels showed a

significant increase of DNA content compared to day 1. In contrast, the DNA content was

observed to decrease in 6% VPM-PEGDA hydrogels. Proliferation of SMCs was

decreased significantly with increasing VPM-PEGDA concentration (Fig. 6.5).

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Figure 6. 3 Morphology of SMCs in differently functionalized PEG-based hydrogels. The final RGD-PEGMA concentration in all the swollen hydrogels was 0.502 mM. The arrows indicated the limited spreading of SMCs in GIA peptide functionalized hydrogels.

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Figure 6. 4 The effect of MMP sensitivity on SMC proliferation in 3D PEG-based hydrogels. (VPM/GIA/IGA-PEGDA: 4%, w/w; RGD-PEGMA: 1.25 mM). Proliferation was quantified by measuring DNA content using PicoGreen assay and reported relative to the DNA content at Day 1. *: p<0.05 with regard to DNA content at day 1 of hydrogels with the same composition; #: p<0.05 with regard to DNA content of IGA-PEGDA hydrogels at the same day; ψ: p<0.05 with regard to DNA content of GIA-PEGDA hydrogels at the same day.

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Figure 6. 5 The effect of network cross-linking density on SMC proliferation in 3D hydrogels (VPM-PEGDA: 4%-6%, w/w; RGD-PEGMA: 0.502 mM in all the swollen gels). Proliferation was quantified by measuring DNA content using PicoGreen assay and reported relative to the DNA content at Day 1. *: p<0.05 with regard to DNA content at day 1 of hydrogels with the same composition; #: p<0.05 with regard to DNA content of 4% VPM-PEGDA hydrogels at the same day; ψ: p<0.05 with regard to DNA content of 5% VPM-PEGDA hydrogels at the same day.

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6.3.6 The Effect of Heparin on Cell Morphology and Proliferation in 3D and 2D

Cell proliferation as a function of heparin concentration was also examined in both 3D bioactive hydrogels (VPM-PEGDA: 4%, RGD-PEGMA: 1.25 mM) and on 2D surfaces. In 3D gels, the inclusion of heparin into SmGM-2 growth medium resulted in limited cell spreading (at Day 21) even when the heparin concentration was relatively

low (0.5 mg/ml) (Fig. 6.6A). Cell proliferation was dramatically inhibited in the presence

of heparin (Fig. 6.6). The inhibitory effect of heparin on SMC proliferation was also

observed on 2D surfaces, however, is not as evident as in 3D gels (Fig. 6.7).

6.4 Discussion

An understanding of the effect of scaffold properties on SMC proliferation would

be of great benefit in developing strategies to regulate SMC proliferation and new

treatments for vascular disease, as well as in fabricating functional tissue engineered

blood vessels. Compared to a 2D culture system, a 3D engineered scaffold is more similar to the in vivo cell-ECM microenvironment and more appropriate for exploration of fundamental cell biology [26-28]. Hydrogels have been utilized for 3D studies of cell functions, because they can provide a soft tissue-like 3D environment for cell growth and allow diffusion of exogenous bioactive factors through the highly swollen 3D network

[27, 28]. 3D biomimetic PEG hydrogels with the tunable properties of cell adhesion and

proteolytic degradation were selected as the scaffold to study SMC proliferation in 3D.

Our results have shown that the incorporation of cell adhesive peptides into the

PEG network facilitates SMC spreading in hydrogels and cell adhesive ligand concentration (0-2.5 mM) mediates 3D SMC proliferation in a biphasic manner (Fig.

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Figure 6. 6 Morphology and proliferation of SMCs in 3D bioactive hydrogels (VPM- PEGDA: 4%, w/w; RGD-PEGMA: 1.25 mM) as a function of heparin concentration (0-2 mg/mL). (A) Confocal laser scanning micrographs of cell morphology in 3D hydrogels with treatment with varied concentrations of heparin. (B) The effect of heparin concentration on cell proliferation in 3D hydrogels. Proliferation was quantified by measuring DNA content using PicoGreen assay and reported relative to the DNA content at Day 1. *: p<0.05 with regard to DNA content at day 1 of hydrogels treated with the same concentration of heparin; #: p<0.05 with regard to DNA content of hydrogels without heparin treatment at the same time point.

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Figure 6. 7 Proliferation of SMCs on 2D human fibronectin (FN, 1µg/cm2)-coated tissue culture polystyrene surfaces as a function of heparin concentration (0-2 mg/mL). Proliferation was quantified by measuring DNA content using PicoGreen assay and reported relative to the DNA content at Day 1. *: p<0.05 with regard to DNA content at day 1 of surfaces with the same concentration of heparin; #: p<0.05 with regard to DNA content of surfaces without heparin treatment at the same time point.

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6.2). Mann et al. found that increasing ligand density decreased SMC proliferation on 2D

RGD modified surfaces and in 3D PEG based hydrogels [38]. However, they did not see the initial increase in proliferation with increasing peptide concentration on 2D surfaces and in 3D hydrogels, while Neff et al. found a maximum proliferation of fibroblasts at an intermediate peptide concentration on 2D RGD modified surfaces [39], similar to our observation for SMC proliferation in 3D gel (Fig. 6.2B). These inconsistencies may be related to differences of the applied adhesive peptide concentration, the additional dimensionality or other factors that may affect cell proliferation, such as mechanics [29].

In contrast to our finding that the RGD incorporation facilitated SMC proliferation in 3D scaffold, less proliferation was observed in scaffolds grafted with cell adhesive ligands than in nonadhesive control scaffolds in Mann et al. studies [38]. The contrasting observations might be attributed to the differences of other scaffold properties, such as degradability and network crosslinking density.

To investigate the effect of proteolytic degradation on SMC behavior in 3D gels,

SMC spreading and proliferation in the hydrogels modified with MMP-sensitive peptides with different enzyme sensitivity were examined at the constant RGD concentration and network structure (Fig. 6.3-6.4). VPM-PEGDA hydrogels were shown to enhance SMC spreading and proliferation more significantly than GIA-PEGDA gels and non- degradable IGA-PEGDA hydrogels (Fig. 6.3-6.4). SMCs secrete a variety of MMPs, including MMP-1, -2, -3, -7, -9, and membrane type 1-MMP (MT1-MMP) [6].

Compared with GPQGIAGQ peptide, VPMSMRGG peptide has been reported to be more proteolytically sensitive to many MMPs, such as MMP-1, -2, -3, and -9 [33]. As expected, VPM modified hydrogels degrades faster than GIA modified hydrogels in the

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presence of MMPs [33]. This indicates that faster proteolytic degradation is required in order to create sufficient space for cell proliferation within these biomimetic hydrogels. A similar dependency on cell-mediated proteolytic degradation of the hydrogel network has previously been observed for cell migration in 3D [36], as well as cell proliferation in 3D for other cell types [33]. Additionally, our previous studies have shown that SMCs were able to spread and migrate in the hydrogels modified with GIA peptide by encapsulation of a cell pellet in the gels [36]. By utilizing the cell pellet method, more enzymes would likely be secreted from the cell pellet than a single cell in the homogeneously seeded hydrogels, which could facilitate SMC invasion.

The effect of the hydrogel network on cell proliferation in 3D gels was evaluated by varying the concentration of VPM-PEGDA in the hydrogels. At a constant concentration of RGD-PEGMA (0.502 mM, the concentration which shows maximum proliferation in 4% VPM-PEGDA gels) in the swollen hydrogel, increasing VPM-

PEGDA concentrations significantly decreases SMC spreading and proliferation rate

(Fig. 6.3 and Fig. 6.5). This observation is in contrast with previous studies on 2D surface, in which SMC proliferation was enhanced with increasing crosslinking density

[29]. This opposite trend may be attributed to the physical barrier and confinement posed by the 3D network, which may alter the mechanical signaling from the scaffolds [26, 27].

The distribution of key signaling factors may be quite different in flat cells on a 2D surface versus more elongated spindle-shaped SMCs in 3D gels [26, 40]. Furthermore, the levels of MMPs produced by the cells responded to the network microarchitecture might be altered and need to be further investigated [41].

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Through characterization of SMC proliferation in hydrogels with different

composition, we have identified that gels fabricated from 4% VPM-PEGDA and 1.25

mM RGD-PEGMA (in precursor solution) present optimal SMC spreading and

proliferation. By utilizing this optimal composition, we examined the effect of heparin

concentration on SMC spreading and proliferation at day 21. The presence of heparin,

even at the relatively low concentration (0.5 mg/ml), has dramatically inhibited the

spreading and proliferation of SMCs in 3D hydrogel, while on 2D surfaces SMCs could

still proliferate at this concentration (Fig. 6.6-6.7). This indicated that heparin has more

evident effect on inhibition of SMC proliferation in 3D scaffolds than on 2D surfaces.

Studies of heparin on cell enzyme secretion have shown that heparin inhibits the

expression of matrix-degrading proteinase such as plasminogen activators and MMPs, including MMP-2, -3 and -9 [22, 42, 43]. Since proteolysis plays a predominant role in

3D SMC proliferation, the decrease of MMP secretion from SMCs by heparin treatment might result in a significantly greater influence of heparin on inhibition of SMC proliferation in 3D gels than on 2D surfaces.

6.5 Conclusion

PEG based hydrogels can be rendered cell adhesive and biodegradable by the incorporation of bioactive peptides (e.g. cell adhesive peptide and enzyme sensitive peptide) into the polymeric network through photopolymerization. Through utilizing this defined, biomimetic PEG scaffold, 3D SMC proliferation has shown to be critically dependent on cell-matrix adhesiveness and proteolysis. The incorporation of cell adhesive ligand significantly enhanced SMC spreading and proliferation and the concentration of cell adhesive ligand mediated 3D SMC cell proliferation in a biphasic

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manner. The faster degrading hydrogels can facilitate SMC proliferation and spreading.

In particular, higher crosslinking density could impede SMC proliferation and spreading

despite the presence of cell adhesive ligand and proteolytically degradable sites. Further,

heparin could act as exogenous inhibitory factors to regulate SMC proliferation in 3D.

These synthetic ECM-mimetic scaffolds may be useful for 3D mechanistic studies of

many aspects of SMC functions, including the effect of scaffold properties on SMC phenotypic switch and ECM production, and the effect of other exogenous bioactive factors (e.g., PDGF, transforming growth factor-β) on 3D SMC functions.

Furthermore, the optimal gel composition to enhance SMC spreading and proliferation has been determined, which can be utilized in the promotion of vascular regeneration. The evident antiproliferative effect of heparin in 3D gels suggests that development of heparin releasing hydrogels by encapsulation of heparin into gels could show promise to regulate SMC proliferation without exogenous stimulation. Further exploration of SMC mechanisms by this 3D model and engineering of improved scaffold system will facilitate the development of functional tissue-engineered blood vessels.

6.6 Acknowledgement

The project described was supported by Grant Number 5RC1EB010795 and

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35. Pierschbacher MD, Ruoslahti E. Cell attachment activity of fibronectin can be duplicated by small synthetic fragments of the molecule. Nature 1984;309:30-3.

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38. 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. Journal of biomedical materials research 2002;60:86-93.

39. Neff JA, Tresco PA, Caldwell KD. Surface modification for controlled studies of cell-ligand interactions. Biomaterials 1999;20:2377-93.

40. Gerthoffer WT. Mechanisms of vascular smooth muscle cell migration. Circ Res 2007;100:607-21.

41. 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. Proceedings of the National Academy of Sciences of the United States of America 2003;100:5413-8.

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CHAPTER 7 Conclusions and Future Directions

7.1 Summary and Conclusion of Completed Work

This work investigated the hypothesis that 3D SMC migration and proliferation can be regulated by the properties of a cell-instructive scaffold, including cell-matrix adhesion, degradability, and cross-linking density. Photopolymerizable poly (ethylene glycol) diacrylate (PEGDA) derivatives were used to fabricate hydrogels as scaffold substrates. This system allows for in situ cell encapsulations, which can be utilized for 3D cell culture. Further, the unmodified PEGDA hydrogel resists protein adsorption and cell attachment. This bio-inert PEG hydrogels can function as a biological blank slate to incorporate bioactive factors in a controlled manner, which makes it possible to engineer the PEG gels with desired bioactivities and examine their effects on cell responses.

In Chapter 4, bioactive PEG hydrogels was fabricated by copolymerization of cell-adhesive peptide (GRGDSP) modified PEG monoacrylates and enzyme-sensitive peptide (GPQGIAGQ, VPMSMRGG) modified PEG diacrylates. The scaffold properties, including cell adhesivity, proteolytical degradability, and network properties, were characterized. These experiments showed that the cell adhesivity of PEG hydrogels can be modulated by the quantitative control of incorporated cell-adhesive peptide concentration, and the degradability of gels can be tuned by the simple change of enzyme-sensitive peptide type present in the network. Analysis of these networks showed that the biochemical properties of hydrogels (e.g., cell adhesivity, biodegradability) can be controlled independently without affecting the network properties of hydrogels under certain conditions. This defined, biomimetic hydrogel showed potentials as a3D cell culture model for exploration of cell biology.

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In Chapter 5, SMC migration behavior in 3D bioactive hydrogels developed in

Chapter 4 was investigated. The effect of scaffold properties, including cell-matrix

adhesiveness, proteolysis, and network cross-linking density, on 3D SMC migration was

evaluated. The results showed that 3D SMC migration had a biphasic dependence on

adhesive ligand density, and both adhesive and enzyme-sensitive peptides were required for migration to occur. Furthermore, network cross-linking density was shown to dramatically influence the behavior of 3D cell migration. These results supported the hypothesis that 3D SMC migration can be regulated by the properties of biomimetic PEG scaffolds, including cell-matrix adhesion, degradability, and cross-linking density.

In Chapter 6, 3D bioactive hydrogels developed in Chapter 4 continued to be utilized as 3D culture models to investigate 3D SMC proliferation. The effect of both scaffold properties and exogenous bioactive factors on 3D SMC proliferation was quantified. The studies indicated that 3D SMC proliferation was critically dependent on cell-matrix adhesiveness, degradability and cross-linking density. SMC spreading and proliferation was significantly enhanced by the incorporation of cell-adhesive ligand and the concentration of cell-adhesive ligand mediated 3D SMC proliferation in a biphasic manner. The faster degrading hydrogels promoted SMC proliferation and spreading, while higher cross-linking density could impede SMC proliferation and spreading.

Further, heparin could act as exogenous inhibitory factors to regulate SMC proliferation in 3D. These results supported the hypothesis that 3D SMC proliferation can be regulated by the properties of a cell-instructive scaffold, including cell-matrix adhesion, degradability, and cross-linking density, and suggested that the use of soluble bioactive factors, such as heparin, is an effective approach to regulate SMC proliferation in 3D.

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Compared with 2D culture studies, the exploration of SMC migration and

proliferation in 3D biomimetic scaffolds provides information that is more relevant to

behavior of cells in vivo. Besides SMC migration and proliferation, the defined, synthetic

scaffolds can be useful as tools for 3D mechanistic studies of other aspects of SMC

functions. The potential applications of 3D scaffolds in studies of SMC biology will be

described below. Further, this scaffold system can be developed with improved scaffold

properties for 3D cell culture models and vascular tissue engineering applications.

Components of this system that require additional engineering will be discussed below.

Additionally, the investigation of the in vivo performance of this scaffold by using animal

studies will be proposed below.

7.2 Exploring SMC Functions in 3D Scaffolds

7.2.1 Investigation of SMC Phenotype and ECM Remodeling

In normal vascular vessel, SMCs are nonmigratory and not proliferating. They are

responsible for the dilation and constriction of the vessels, which enables blood vessels to

maintain an appropriate blood pressure [1, 2]. However, SMCs also perform other

functions, which play a key role in a variety of physiological and pathological conditions,

such as vascular remodeling after vascular injury [3]. In response to injury, SMCs

upregulate the secretion of matrix metalloproteinases (MMPs), increase migration and

proliferation, and synthesize large amounts of extracellular matrix (ECM) components

[4-7]. These processes define a transition from a contractile SMC phenotype to a synthetic phenotype, which is essential for tissue regeneration and vascular repair [8-10].

Later in the development of vascular remodeling, synthetic SMCs must switch back to a contractile phenotype, which can minimize the development of intimal hyperplasia and

146

assist in the biomechanical functionality of blood vessels [8, 9]. Therefore, understanding

the mechanisms underlying SMCs phenotypic modulation is the subject of intense

research.

Previous studies of SMC phenotypic modulation on 2D surfaces have indicated

that SMC phenotypic switch is reversible and is regulated by a variety of factors, including soluble bioactive factors, ECM components, and physical factors [8-10]. To gain further insight of SMC phenotypic modulation in vivo, 3D bioactive PEG scaffold can be utilized to investigate a variety of biochemical and biophysical factors on SMC phenotypic transitions. . The SMC phenotype can be studied by migration, proliferation, expression of collagen and vimentin (synthetic markers), gene expression and immunofluorescent localization of smooth muscle α-actin, calponin and SM22α

(contractile markers) [9, 11, 12] .

7.2.2 Investigation of Additional Mediators

As described in Chapter 2, there are a variety of mediators that affect SMC functions, including soluble bioactive factors, extracellular matrix components, and physical factors [5, 6, 9, 13]. This work has indicated that heparin could act as an exogenous inhibitory factor on SMC proliferation. The other regulatory bioactive factors, such as platelet-derived growth factor (PDGF) [14-16], transforming growth factor β

(TGF β) [17, 18], or angiotensin II [16], can be used as exogenous factors to study their individual effect on SMC functions in 3D scaffolds. Besides their individual role, the

synergistic effect of these bioactive factors on SMC functions can be explored by using

this 3D culture model.

147

In vivo, the vascular SMCs are subjected to dynamic mechanical stimuli [8, 9]. It is clear that the dynamic mechanical stimuli play an important role in mediating SMC functions, such as phenotypic modulation, SMC proliferation, and ECM production [19-

23]. However, the mechanisms that regulate these responses are not well understood [9].

By using the 3D bioactive scaffold, the effect of dynamic mechanical stimuli (e.g., cyclic strain) on 3D SMC functions can be investigated. Moreover, the tunable biochemical and biophysical properties of this scaffold make it possible to evaluate the synergistic effect of dynamic mechanical stimuli and other environmental cues (e.g., matrix stiffness, RGD concentration, and degradability) on 3D SMC functions.

7.3 Engineering of Improved Scaffold Systems

7.3.1 SMC-selective Adhesive Ligand

This work employed the cell-adhesive peptide sequence, RGD, to bind with the

integrin receptors on SMCs. RGD is the most extensively studied cell-adhesive peptide,

which has been demonstrated to adhere to many cell types, such as endothelial cells

(ECs) and fibroblasts, as well as platelets [24-27]. For in vitro 3D cell culture models,

particularly in single cell culture, it is acceptable to use the cell nonspecific adhesive

peptide to study the effect of cell-matrix adhesiveness on cell functions. However, in

vascular tissue engineering applications, biospecific and cell-selective adhesion within

each of the layers (i.e., intima, media, and adventitia) are required to ensure development

of the appropriate tissue structure [28]. For example, EC selective peptide,

CRRETAWAC, has been used to modify the surface of ePTFE grafts, which has been

shown to facilitate the endothelization on ePTFE grafts and prevent platelet adhesion

[29]. In normal vessels, the SMCs in vascular media are surrounded primarily by elastin

148

and laminin. VAPG is a peptide sequence derived from elastin, which has been shown to

be specific for adhesion of SMCs [30]. Fibroblasts, ECs, and platelets could not adhere to

VAPG [30]. Although SMC attachment has been reported on PEGDA hydrogels

modified with VAPG, other studies found that SMCs did not adhere to VAPG [11]. The

inconsistent findings might be due to SMC species differences, lot-to-lot differences in cell response, or changes in cell receptor library [11]. Therefore, further attempt to find a robust SMC selective cell-adhesive peptide are needed to facilitate the development of a functional vascular media.

7.3.2 Modification of Bifunctional Peptides with Cell Adhesivity and Biodegradability

In this project, the RGD peptide was incorporated into the polymeric network by

copolymerization of monoacrylated RGD peptides and PEG diacrylates. It has been

shown that the inclusion of the monacrylated peptides did not affect the swelling ratio

and mechanical properties of the hydrogels when the concentration of incorporated

peptides was relatively low [31, 32]. However, increasing the concentration of peptides over the range will subsequently affect the network properties of hydrogels [31].

Therefore, to independently control the scaffold properties, the extent of incorporated peptide is limited by using this method. To better control the biochemical and biophysical properties of hydrogels, a bifunctional peptide constituted by cell-adhesive peptide and enzyme-sensitive peptide can be incorporated into the scaffolds. By utilizing this method, the cell-adhesive peptide will be allocated on the backbone of the polymeric network, which will achieve improved control of peptide incorporation and scaffold properties

[33].

7.3.3 Improved Heparin Delivery

149

Heparin was utilized as an endogenous factor to regulate SMC functions in this work. To engineer a functional vascular media in tissue engineering, a cell-instructive scaffold must be able to not only stimulate vascular regeneration, but also regulate SMC functions to prevent failure mechanisms [8]. For example, promoting the shift of synthetic, proliferative SMCs toward a contractile, non-migratory, non-proliferative SMC phenotype at an appropriate cell development stage is one strategy to minimize the development of intimal hyperplasia (IH) [9]. To accomplish this, a microparticle delivery system encapsulated with regulatory factors, such as heparin, can be developed to control the release of the factors to regulate cell functions. The delivery system can be subsequently incorporated into 3D biomimetic PEG scaffold, which will facilitate the regulation of SMC functions without exogenous stimulation. There are several strategies to generate the delivery systems. For example, poly (lactide-co-glycolide) microspheres have been utilized for controlled release of a variety of bioactive factors, such as heparin and TGF β [34, 35]. Highly cross-linked hydrogel microparticles have also been employed for drug delivery applications [36-38]. It is of note that enzyme-sensitive peptides can be incorporated into the delivery system to establish the cell-mediated release mechanism.

7.4 In vivo Animal Studies

Although 3D cell culture models recapitulate many critical aspects of in vivo extracellular environments, in vivo testing of hydrogel scaffolds is still needed. The complex, dynamic in vivo environment might affect the behaviors of hydrogel scaffolds.

For example, hydrogel degradation in vivo is mediated not only by SMCs, but also by other cell types secreting MMPs, such as macrophages, fibroblasts, and ECs. In

150

preparation for in vivo testing, these materials must be incorporated into existing vascular

grafts since the scaffold alone is too weak to serve as a vascular prosthesis. To reinforce the gel, the biomimetic PEG scaffolds can be incorporated into a more mechanically robust material, such as ePTFE materials. The modified ePTFE grafts can subsequently

implanted into an animal model, such as a porcine model, to investigate the effect of

scaffolds on tissue regeneration and also the development of vascular disease. To

accomplish this, a robust protocol must be developed to incorporate bioactive PEG

hydrogels into ePTFE grafts.

151

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