ELECTROSPINNING OF RESORBABLE AMINO-ACID BASED

POLY(ESTER UREA)S FOR REGENERATIVE MEDICINE

A Dissertation

Presented to

The Graduate Faculty of The University of Akron

In Partial Fulfillment

of the Requirements for the Degree

Doctor of Philosophy

Yaohua Gao

May, 2016 ELECTROSPINNING OF RESORBABLE AMINO-ACID BASED

POLY(ESTER UREA)S FOR REGENERATIVE MEDICINE

Yaohua Gao

Dissertation

Approved: Accepted:

______Advisor Department Chair Dr. Matthew L. Becker Dr. Coleen Pugh

______Committee Member Dean of the College Dr. Darrell H. Reneker Dr. Eric J. Amis

______Committee Member Dean of the Graduate School Dr. Abraham Joy Dr. Chand Midha

______Committee Member Date Dr. Li Jia

______Committee Member Dr. Ge (Christie) Zhang

ii ABSTRACT

Electrospinning generates very fine fibers with diameters that bridge nanometer to micrometer length scales and loosely connected 3D fibrous structure which can mimic native extracellular matrix (ECM). Some other advantages of electrospun nanofibers include high surface area per unit mass, variety of surface functionalization strategies, simplicity of process, and tunable morphology. These advantages make electrospun nanofibers very attractive candidate for regenerative medicine applications, such as tissue engineering and drug delivery. Degradable polymers, including PCL, PLA, PGA and their copolymers or blends are the most commonly known and used materials to fabricate electrospun scaffolds for regenerative medicine. However, these materials are limited in that they create acidic degradation byproducts that cause inflammation and cell phenotypes changes in surrounding tissues. In the first part of this work, a new class of resorbable L-leucine based poly(ester urea)s (PEUs) was developed by interfacial polymerization. The chemical, thermal and mechanical properties of the resulting polymers were characterized. This new class of PEUs showed wider tunable range of thermal, mechanical and degradation properties than most commonly studied degradable polymers. The in vitro hydrolytic degradation study indicated the PEU polymers are degradable and it did not cause significant pH drop during the degradation process. The protein adsorption study showed this kind of materials had PRP blocking effect after pre-adsorbing fibrinogen. Vascular cells, such as smooth muscle cells and endothelia cells,

iii were found to be able to spread and attach on the PEU polymer film. Taken together, the data suggest the new PEUs are promising candidates for vascular tissue engineering.

As a continuous work to explore the application of PEUs in vascular tissue engineering, small diameter vascular grafts with 1 mm inner diameter were fabricated using electrospinning. Long term performance of two types of PEU grafts with different wall thickness (type A: 250 μm; type B: 350 μm) were evaluated in an abdominal infra-renal aortic mouse model over 1 year. Significantly, the small diameter vascular grafts did not rupture or lead to acute thrombogenic events in the intervals tested. The pilot study in vivo showed long term patency and extensive tissue remodeling with type A grafts, while the type B grafts experienced occlusion over the 1 year interval due to intimal hyperplasia. This study affords significant findings that will guide the design of future generations of small diameter vascular grafts.

In the third part of my doctoral research, the application of electrospun PEU scaffold in protein drug delivery was explored. Recombinant human growth hormone

(rhGH) for protein therapeutics is in great demand. However, as a consequence of short half-life, it is still quite a challenge to develop effective rhGH sustained delivery systems that can exceed a month in duration. In this part, a new sustained release strategy of rhGH was developed by encapsulating sugar glass nanoparticle protected rhGH in the electrospun PEU nanofibers. The protein was found to be randomly dispersed throughout the electrospun fibers in an aggregate form. Sustained rhGH release with modest burst release was observed for up to 6 weeks in vitro as confirmed by protein assay. These results clearly suggest the feasibility of using electrospun PEU nanofibers as new, long-term sustained release strategy for rhGH.

iv ACKNOWLEDGEMENTS

This dissertation summarizes five years of my challenging but rewarding doctoral research in the department of Polymer Science at the University of Akron. I appreciate the guidance, mentoring, and help of many people. This dissertation would not have been possible without you. Foremost, I acknowledge my advisor, Dr. Matthew L. Becker, for providing me the opportunity to work on these challenging and impactful projects. Dr.

Becker has been extremely helpful for my progress in my Ph.D work. Thank you for teaching me dream big, know how to have fun with work, and make bold decisions in face of uncertainty. Besides my advisor, I want to acknowledge my collaborators, Dr.

Darrell H. Reneker, Dr. Christopher Breuer, Dr. Tai Yi, Dr. Todd Ritzman, and Dr. Adam

Land. My collaborators have been incredibly helpful in developing new strategies to solve challenges in the projects. I also want to acknowledge my dissertation committee members - Dr. Abraham Joy, Dr. Li Jia, Dr. Ge Zhang, Dr. Darrell H. Reneker, and Dr.

Matthew L. Becker for their guidance and time. Thank you very much for challenging me to answer tough questions to help me understand the projects. Last but not least, I thank my family and friends. Thank my dear parents, brother, sister and husband for their unconditional love and support through my whole Ph.D study. Thanks for being always proud of me. Thank my friends for always being there for me, I won’t point out your names here, but you know who you are!

v TABLE OF CONTENTS

Page

LIST OF TABLES ...... x

LIST OF FIGURES ...... xi

LIST OF SCHEMES ...... xiv

CHAPTER

I. INTRODUCTION ...... 1

1.1 Electrospinning...... 1

1.1.1 Theory of electrospinning ...... 1

1.1.2 Parameters affecting electrospinning process and fiber properties ...... 3

1.1.3 Applications in regenerative medicine...... 8

1.2 Small diameter vascular grafts (SDVGs) ...... 15

1.2.1 Clinical needs and challenges for SDVGs ...... 15

1.2.2 Resorbable Scaffold guided SDVGs tissue engineering ...... 16

1.2.3 Electrospun scaffolds for SDVGs tissue engineering ...... 29

1.3 Controlled protein delivery ...... 32

1.3.1 Pharmaceutical proteins and delivery issues...... 33

1.3.2 Electrospun nanofibers for protein delivery ...... 33

1.4 Poly(ester urea)s (PEUs) ...... 39

II. MATERIALS AND INSTRUMENTS ...... 41

2.1 Materials ...... 41 vi 2.2 Instruments ...... 42

III. L-LEUCINE BASED POLY(ESTER UREA)S FOR VASCULAR TISSUE ENGINEERING ...... 46

3.1 Outline ...... 46

3.2 Introduction ...... 47

3.3 Experimental Section ...... 49

3.3.1 Materials and cells ...... 49

3.3.2 Instrumentation ...... 50

3.3.3 Synthesis of di-p-toluenesulfonic acid salts of bis-L-leucine diester monomer ...... 51

3.3.4 Synthesis of L-leucine-based poly(ester urea)s using interfacial polymerization ...... 55

3.3.5 Mechanical property measurements...... 57

3.3.6 In vitro degradation tests...... 57

3.3.7 Water uptake study ...... 58

3.3.8 Quartz crystal microbalance (QCM)...... 59

3.3.9 Cell attachment and spreading studies ...... 59

3.3.10 Statistics ...... 60

3.4 Results and discussion ...... 61

3.4.1 Polymer synthesis and characterization ...... 61

3.4.2 Thermal properties ...... 64

3.4.3 Mechanical properties ...... 67

3.4.4 In vitro degradation studies ...... 69

3.4.5 Protein adsorption ...... 73

3.4.6 Cell culture studies ...... 75

3.5 Conclusion ...... 77 vii 3.6 Acknowledgements ...... 78

IV. PILOT RAT STUDY OF 1 MM INNER DIAMETER (ID) VASCULAR GRAFT USING ELECTROSPUN POLY(ESTER UREA) NANOFIBERS ...... 79

4.1 Outline ...... 79

4.2 Introduction ...... 80

4.3 Experimental section ...... 83

4.3.1 Materials ...... 83

4.3.2 Polymer synthesis and characterization ...... 83

4.3.3 Graft fabrication ...... 84

4.3.4 Graft characterization ...... 85

4.3.5 Biomechanical evaluations ...... 85

4.3.6 Biological activity evaluations ...... 87

4.3.7 Animal study ...... 89

4.3.8 Statistical analysis ...... 92

4.4 Results and discussion ...... 92

4.4.1 Polymer synthesis and characterization ...... 92

4.4.2 Scaffold characterization ...... 93

4.4.3 Biomechanical properties of scaffolds ...... 95

4.4.4 In vitro cell study results ...... 98

4.4.5 Survival of the animals ...... 102

4.4.6 Ultrasound and micro-CT ...... 102

4.4.7 Histological assessment ...... 104

4.5 Conclusion ...... 106

4.6 Acknowledgments ...... 106

viii V. NEW SUSTAINED RELEASE STRATEGY OF RECOMBINANT HUMAN GROWTH HORMONE (rhGH) FROM BIORESORBABLE POLY(ESTER UREA) NANOFIBERS ...... 107

5.1 Outline ...... 107

5.2 Introduction ...... 108

5.3 Experimental section ...... 110

5.3.1 Materials ...... 110

5.3.2 Polymer synthesis and characterization ...... 111

5.3.3 SGnPs preparation and characterization ...... 111

5.3.4 Electrospun fiber mat fabrication and characterization ...... 112

5.3.5 In vitro protein release study...... 114

5.4 Results and discussion ...... 114

5.4.1 Sugar glass protein nanoparticle ...... 114

5.4.2 Electrospun fiber mat fabrication and characterization ...... 116

5.4.3 In vitro release study ...... 118

5.5 Conclusion...... 120

5.6 Acknowledgements ...... 121

REFERENCES ...... 122

APPENDIX...... 142

ix LIST OF TABLES

Table Page

1.1. Biodegradable polymers and their degradation properties…………………….……26

1.2. Tissue engineered SDVGs fabricated with biodegradable synthetic polymers…..…28

1.3. A summary of electrospun scaffolds for SDVGs (ID < 6 mm)………………...……31

1.4. A summary of protein delivery systems based on electrospun nanofibers………….38

3.1. Characterization data summary of the L-Leucine-based PEUs…………………..…62

3.2. Mechanical properties of L-leucine-based PEUs with diols of different length…….69

3.3. Water uptake of L-leucine-based poly(ester urea)s with diols of different length after being soaked in PBS buffer at 37 °C for one week……………………………….…71

3.4. Summary of protein and PRP adsorption on PEU surface……………………….....74

4.1. Characterization data summary for the amino acid-based poly(ester urea)………....93

4.2. Physical properties summery of poly(1-LEU-10) electrospun grafts…………….....97

x LIST OF FIGURES

Figure Page

1.1. (a) Schematic illustration of electrospinning. (b) Bending instability of electrically charged liquid jets in electrospinning…………………………………………...……2

1.2. Scaffold architecture affects cell adhesion and spreading. (a) and (b) Cells adhere to scaffolds with microscale architecture, flatten and spread as if cultured on flat surfaces. (c) Scaffolds with nanoscale architecture have larger surface area to adsorb proteins, presenting many more binding sites to cell membrane receptors...... 9

1.3. Schematic illustration of in vitro, in vivo, and in situ vascular tissue engineering….19

1.4. Decellularized-matrix vascular tissue engineering……………………………….…24

1.5. Schematics of various experimental setups for the electrospinning process for the fabrication of tubular scaffolds. (a) A multilayer scaffold combining electrospinning and hydrogel fabrication. (b) Co-electrospinning of two polymer solutions concurrently. (c) Electrospinning with simultaneous electrospraying………………30

1.6. Fabrication techniques of bioactive electrospun nanofibers. (a) Physical adsorption. (b) Blend electrospinning. (c) Coaxial electrospinning. (d) Covalent immobilization………………………………………………………………………37

1 3.1. H NMR (DMSO-d6) of di-p-toluenesulfonic acid salts of bis-L-leucine- diester monomers………………………………………………………………………...….53

13 3.2. C NMR (DMSO-d6) of di-p-toluenesulfonic acid salts of bis-L-leucine- diester monomers…………………………………………………………………………....54

3.3. FT-IR of di-p-toluenesulfonic acid salts of bis-L-leucine-diester monomers…….…54

1 3.4. H NMR (DMSO-d6) spectra of L-leucine-based poly(ester urea)s……………..….62

13 3.5. C NMR (DMSO-d6) spectra of L-leucine-based poly(ester urea)s…………..……63

3.6. FT-IR spectra of L-leucine-based poly(ester urea)s………………………………....63

3.7. TGA analysis of L-leucine-based poly(ester urea)s…...... …………...... ….....65

xi 3.8. DSC trace of L-leucine-based poly(ester urea)s…………………………………….66

3.9. DMA results of L-leucine-based poly(ester urea)s………………………………….66

3.10. Stress-strain plots of L-leucine based PEUs measured at room temperature (20 ± 1 °C) (a, b)) and physiological temperature (37 ± 1 °C) (c, d) using an Instron 3365 universal materials testing machine…………………………………………………68

3.11. In vitro degradation tests were performed according to ASTM standard F1635-11: “In vitro degradation testing of hydrolytically degradable polymer resins and fabricated forms for surgical implants”……………………………………………..72

3.12. Protein adsorption properties of PEU monitored by QCM-d. The experiment contains four processes: (i) baseline in PBS buffer; (ii) adsorption of proteins; (iii) adsorption of PRP; (iv) PBS buffer washing. The shift in QCM frequency corresponds to adsorption on the polymer surface. A decrease in frequency indicates a mass increase due to adsorption, and the extent of decrease is directly proportional to the amount of increased mass…………………………………………………….74

3.13. A-10 smooth muscle cells (A-10 SMCs) and human umbilical vein endothelial cells (HUVECs) attachment and spreading on PEU thin films (glass coverslips were studied as positive control)………………………………………………………….76

4.1. (a) The gross appearance and SEM images of small diameter electrospun PEU grafts, (b) entire (×40), (c) surface (×2500), and (d) cross-sectional (×2500) morphologies………………………………………………………………………...95

4.2. Stress-strain curve of PEU graft in wet condition from uniaxial tensile testing….....97

4.3. Suture retention strength of PEU whole grafts test with commercial 5-0 Prolene sutures…………………………………………………………………………….....98

4.4. A-10 smooth muscle cells (A-10 SMCs) and human umbilical vein cells (HUVECs) attachment and spreading on PEU electrospun nanofibers (cell seed density: 78 mm-2; red: F-actin stained by rhodamine phalloidin; blue: nucleus stained by DAPI)…...100

4.5. Cell proliferation of A-10 smooth muscle cells (A-10 SMCs) and human umbilical vein cells (HUVECs) cultured in direct contact with electrospun PEU nanofibers after 1, 3 and 7 days of cell seeding, as determined by PrestoBlue assay. Blank glass coverslips were studied as positive controls………………………………………101

4.6. (a) Intraoperative photograph demonstrating poly(1-LEU-10) vascular grafts during surgical implantation. (b) Serial doppler ultrasound examinations were performed on all implanted grafts. All grafts remained patent to the experimental end point according to the ultrasound tests. (c) Graft inner diameter change was calculated by ImageJ software…………………………………………………….……………...103

xii 4.7. In vivo micro computed tomography (CT) angiography was performed at 12 months (a, b, c, d). All type A grafts (250 μm) showed long term patency at the time point 12 months while all type B grafts (350 μm) demonstrated occlusion……………...…104

4.8. H&E image of harvested poly(1-LEU-10) grafts at 12 months after implantation (a, d, h, k); Endothelial layer of graft lumen stained by CD31 markers (b, e, i, l); Smooth muscle cells stained by immunohistochemical smooth muscle actin (aSMA) (c, f, j, m)…………………………………………………………………………………..105

5.1. (a) Schematic presentation of a biomolecule encapsulated in a SGnP. (b) rhGH-SGnPs suspended in isooctane. (c) Representative TEM image of rhGH-SGnP. (d) Particle size distribution of rhGH-SGnP from DLS ...... …………..116

5.2. SEM image of polymer nanofibers (a) plain PEU, (b) rhGH-SGnP loaded PEU, (c) RB-SGnP loaded PEU, (d) plain PCL, (e) rhGH-SGnP loaded PCL, (f) RB-SGnP loaded PCL…………………………………………………………………………118

5.3. Sustained release profile of rhGH-SGnPs from PEU and PCL electrospun nanofibers within 6 weeks in vitro (insert: schematic presentation of rhGH-SGnPs distribution across the fiber diameter)…………………………………………………………..120

xiii LIST OF SCHEMES

Scheme Page

1.1. The two-step general synthetic scheme of amino acid-based poly(ester urea)s (PEUs)……………………………………………………………………………….40

3.1. A two-step general synthetic route of the L-leucine-based poly(ester urea)s with diols of different chain length……………………………………………………………..51

4.1. The two-step synthetic route of L-leucine-based poly(ester urea)s (P(1-LEU-10))...93

xiv CHAPTER I

INTRODUCTION

1.1 Electrospinning

Electrospinning is a polymer processing technique that produces very fine fibers with diameters that bridge nanometer to micrometer length scales.1-3 Electrospun nanofibers are valued for their ultra-high specific surface area per unit mass, variety of surface functionalization, simplicity of processing, ease of large scale up production, and tunable morphologies, among other benefits.1-3 These materials are explored for a broad range of regenerative medicine applications. In this chapter, the theory of electrospinning, the parameters affecting the process and the application and impact of electrospinning in the biomedical field will be discussed.

1.1.1 Theory of electrospinning

Electrospinning is a polymer processing technique that produces fine fibers by the application of high voltage to a polymer solution or melt.4 The typical set up of electrospinning has three important components (Figure 1.1(a))5 : a high voltage power supply, a spinneret (e.g., a needle with blunt tip, pipette tip), and a grounded collector

(e.g., metal plate, aluminum foil, conductive film, and rotating drum or mandrel). As the

1 high voltage is applied, the body of the polymer solution or melt drop at the spinneret tip will be charged, producing a repulsive electrical force acting opposite to surface tension.

At a certain point (known as a Taylor Cone), a stream of the polymer liquid erupts from the surface of the droplet. The jet of solution is unstable after leaving the Taylor Cone and accelerates towards the collector going through a rapid whipping and bending process

(Figure 1.1(b)).5,6

Figure 1.1.5 (a) Schematic illustration of electrospinning. (b) Bending instability of electrically charged liquid jets in electrospinning. Figures reproduced with permission from Ref. (5) (Copyright © 2014 Royal Society of Chemistry).

The solvent evaporates and the jets solidify into fibers meanwhile, depositing on a collector. The base for fiber collection is either grounded or supplied with a negative charge to further attract the fiber jets. The base can be a flat collector, such as a metal

2 plate, aluminum foil, or screen. It can also be a rotating drum/mandrel to produce circularly aligned fiber or two grounded parallel electrodes (e.g., aluminum strips with a certain gap and metal frame) to produce aligned fibers.7,8 Electrospinning is a clean polymer processing technique, and can be conducted at atmospheric conditions and temperatures.

1.1.2 Parameters affecting electrospinning process and fiber properties

The electrospinning process can be affected by many factors, such as molecular weight of the polymer, viscosity, surface tension, electrical conductivity, applied voltage, solution flow rate, needle size (inner diameter), and gap distance (distance between the needle tip and collector).2,9,10 Fiber size and morphology are also greatly affected by these factors. Therefore, properly selecting parameters during the electrospinning process allows fabrication of polymer nanofibers with tailored size and morphology.

Molecular weight and viscosity. Polymer chain entanglement plays an important role in the electrospinning process. Molecular weight has an effect on polymer chain entanglement in solution. It is reported that solutions with higher polymer molecular weight tend to form more stable fibers, while low molecular weight solution gives fibers with bead defects.11-13 Furthermore, molecular weight is directly proportional to the viscosity of a polymer solution. After leaving the needle tip, the polymer solution is elongated into fibers under the stretching of electrical repulsion force. If the polymer solution is not sufficiently viscous, there is not enough polymer chain entanglement for fiber formation, causing disruption of the polymer jets and bead formation along the fibers. Conversely, if the polymer molecular weight is too low, electrospray formation can

3 occur. Therefore, it is necessary to prepare polymer solutions with sufficiently high molecular mass and viscosity to maintain the consistency of fiber spinning. Finally, solution concentration can also influence the formation of homogenous fibers by changing the solution viscosity. It was reported by Zong et al.14 that fibers with beads were produced at low solution concentration and the beads can be still wet when they reach the collector. Low solution concentration leads to fiber jets with low viscosity that break up into droplets easily as a consequence of surface tension. As increase the polymer concentration to a certain level, uniform fibers without beads are produced. However, if the polymer solution is too viscous it is not easy to maintain the consistency of the fiber spinning process as the droplet dries out before it can form a fiber jet. Taken together, there are optimal polymer molecular mass, viscosities, and solution concentration values for specific polymers in the electrospinning process that have significant effects on the resulting fiber size and morphology.

Surface tension. Surface tension of polymer solutions also plays a critical role in the electrospinning process.15-17 High surface tension values promote molecular aggregation into spherical shapes, thereby inhibiting the fiber spinning process. High surface tension also favors the production of beads in the electrospun fibers. However, this does not imply that a polymer solution with lower surface tension will create fibers without beads or droplets. Surface tension is mostly contributed by the solvent used to dissolve the polymers. Therefore, choosing an appropriate solvent is an important step in consistently producing bead-free fibers.

Electrical conductivity. Polymer solution conductivity is another factor that is critical for the fiber spinning process.14,15,17-19 It was reported that polymer solutions with

4 high electrical conductivity tend to produce fibers with smaller diameters, while polymer solutions with lower electrical conductivity tend to produce fibers with beads. Fibers are created by fiber jet stretching during electrospinning as a consequence of the electrical repulsion of the charges on the jet surface. The number of charges carried on the jet surface is directly proportional to the solution conductivity. Significant decreases in fiber diameter are observed for high conductivity solutions compared to low conductivity solutions. However, Hayati et al.20 reported that highly conductive polymer solutions are very unstable under a high voltage power supply, producing dramatic jet bending instability and broader fiber size distributions. Most synthetic and natural polymers are electrically conductive for electrospinning, with some exceptions such as dielectric materials. Salts are suggested to increase electrical conductivity of the less conductive polymers, stabilize fiber jets and to reduce bead formation. The effect of salts on the morphology and diameter of electrospun fibers were studied by Zong et al.,14 who found that bead-free fibers with relatively small diameters (200-1000 nm ) were created by adding ionic salts like KH2PO4, NaH2PO4, and NaCl to the poly(D,L-lactic acid) (PDLA) solution. Additional examples of the effects of added salts have been studied on other polymers such as poly(vinyl alcohol) (PVA),16 poly(acrylic acid) (PAA),19 and polyamide-6 (Nylon 6).21

Voltage applied. Applied voltages (kV) provide charges on the spinning fiber jet surface. Small changes in the applied voltage can lead to big changes in the produced electrospun fiber diameter and morphology. However, the exact relationship between applied voltage and fiber size is unclear. Some researchers have claimed the fiber diameter increases with increasing applied voltage,22-25 suggesting when higher voltages

5 are applied there is more polymer ejection at the needle tip that facilitates the formation of fibers with larger size. Conversely, others reported there is no significant effect of the applied voltage on electrospun fibers.4,26 Finally, a few research groups even observed that the fiber diameter decreases as the applied voltage increases,6,27-29 reducing the fiber diameter in half with a doubling of applied voltage. It is likely that the applied voltage may affect the fiber diameter in two different processes. Higher applied voltage pushes more polymer solution at the needle tip and leads to a larger diameter initial fiber jet, which facilitates large diameter fiber production. However, higher applied voltage may also cause many charges on the initial fiber jets and this further leads to the spilt of the larger initial jet into many smaller jets and production of fibers with smaller size. Besides creating larger fibers, bead defects were formed at higher applied voltage as the fiber jet instability increased.

Flow rate. The flow rate of polymer solution during the electrospinning process is another important process parameter that is influential on the resulting fiber diameter and morphology. During the electrospinning process, polymer solution passing through the needle tip is carried away under the electrical field and solidifies into nanofibers as the solvent evaporates. The fiber diameter increases as the flow rate increases.30 However, if the flow rate is too high, the polymer solution ejected at the needle tip will be more than it can be carried away, causing bead defects along the fibers to occur.17,19,30,31 The residual solvent in the beads may cause the fibers to collapse together and form webs upon contact. Generally speaking, a lower flow rate is more desirable since it allows the solvent enough time to evaporate.32,33

6 Needle size (inner diameter). The diameter of the needle used in the electrospinning process is another important parameter. The primary functions of the needle are to support the applied voltage used to charge the polymer solution and concurrently act as a narrow vent for the polymer solution to be drawn through. At the beginning of the electrospinning process, a Taylor Cone is formed at the needle tip under the competition between the electrical repulsive force and surface tension. The initial fiber jet is erupted from the Taylor Cone as the electrical field strength is increased. A

Taylor Cone has the inner diameter of the needle as the base and an apex angle of 50o, which is observed in nearly all liquids.34 Therefore, needles with larger diameter will result in a larger Taylor cone and thus larger initial fiber jets. Many studies were performed to understand the effect of the needle size on the electrospinning process.35 It was found that the fiber diameter increases with the needle size as other parameter were kept same. However, clogging and beading can happen if the needle size is too small or too large.

Gap distance. The distance between the needle tip and the grounded collector is called gap distance. It is examined as another approach to control the fiber size and morphology. The effect of the gap distance can be understood by considering the fiber jet path during the electrospinning process.5,6,36,37 As a droplet leaves the Taylor Cone and travels towards the grounded collector, the fiber jets are going through a rapid bending and whipping instability as the solvent evaporates, thinning the fiber. If the gap distance is too short, there is not enough time for the solvent to evaporate when the fibers reach the collector and beads can form. Therefore, it is important to maintain adequate distance and time for the solvent to evaporate during the electrospinning process.

7 1.1.3 Applications in regenerative medicine

Electrospinning is a polymer processing technique rediscovered in recent years.

One main advantage of the electrospinning technique is the production of very fine fibers with diameters that bridge nanometer to micrometer length scales, which are similar to the fibrous structure of natural extracellular matrix (ECM).38-41 Additional advantages of nanofibers include high surface area per mass unit, variety of surface functionalization strategies, simplicity of processing, ease of scale up, and tunable morphology.1-3 All of these advantages make the electrospinning technique very attractive for applications such as filtration,42-48 electrical and optical applications,49-52 protective clothing,53-56 and biomedical applications.1,57-61 The applications of electrospinning in tissue engineering and drug delivery are highlighted in the following sections.

1.1.3.1 Tissue engineering applications

Tissue engineering is an interdisciplinary field typically involving scaffolds and cells or growth factors to regenerate new tissues/organs for implantation into the donor patients due to disease, injury or congenital defects.62 The three dimensional scaffold plays a significant role in tissue engineering by providing structural support and binding sites for cells.63-65 The scaffold architecture affects cell binding and spreading (Figure

1.2).66 Cells adhering to scaffolds with microscale architectures tend to flatten and spread as if cultured on flat surfaces (Figure 1.2(a) and 1.2(b)), while those adhering to scaffolds with nanoscale architectures have larger surface areas to adsorb proteins (Figure 1.2(c)).

The adsorbed proteins may also change conformation, exposing additional cryptic binding sites. A number of approaches were reported for the fabrication of 3D scaffolds

8 for use in tissue engineering, including solvent casting,67,68 particulate-leaching,69-72 gas foaming,73-76 phase separation, 3D printing77-80 and electrospinning.1,57-61 Among them, the electrospinning technique is an excellent candidate because it allows the fabrication of micro-scale interconnected pores, a 3D structure that can resemble the fibrous structure of the natural ECM.38-41 The interconnected pores are also essential for cell nutrition, proliferation, and migration for new tissue regeneration.78,81

Figure 1.2.66 Scaffold architecture affects cell binding and spreading. (a) and (b) Cells binding to scaffolds with microscale architectures flatten and spread as if cultured on flat surfaces. (c) Scaffolds with nanoscale architectures have larger surface areas to adsorb proteins, presenting many more binding sites to cell membrane receptors. Figures reproduced with permission from Ref. (66) (Copyright © 2005 Science).

9 There is a wide range of material choice in making electrospun scaffolds for tissue engineering applications, including natural and synthetic polymers. Among natural polymers, collagen, gelatin, chitosan, silk, and elastin are extensively studied.82-90 For synthetic polymers, poly (ε-caprolactone) (PCL), poly(lactic acid) (PLA), poly(glycolic acid) (PGA) and their copolymers poly(lactic-co-glycolic acid) (PLGA) are mostly used because of their non-toxicity as well as biodegradability.35,91-95 Listed below are a few examples of using electrospun scaffolds for various tissue engineering applications.

Nanofibers for vascular tissue engineering. The nanoscale fibrous structure of electrospun fibers mimics the fibrous network structure in native ECM and provides enhanced cell interaction with the scaffolds.38-41 Electrospinning is employed to fabricate grafts for vascular tissue engineering application. The materials involved are synthetic polymers, natural polymers or a combination of the two materials. For example, He et al.96 fabricated tubular nanofiber scaffolds based on synthetic poly(L-lactic acid)-co-poly(ε-caprolactone) P(LLA-CL 70:30) polymers using electrospinning followed by collagen coating. Human coronary artery endothelial cells (HCAECs) were rotationally seeded onto the lumen of the scaffolds through a custom seeding device. The scaffold was further cultured under static condition. Results showed evenly distributed and well-spread HCAECs throughout the lumen of the scaffold starting from day 1 up to

10 days after seeding. Further, HCAECs maintained phenotypic expression. To prove the basic concept of using the scaffolds as vascular grafts, acellular tubular P(LLA-CL) nanofiber scaffolds (inner diameter 1 mm) were implanted into a rabbit inferior superficial epigastric vein model. Results showed the scaffolds sustained the surgical process, kept the structure integrity, and showed the patency for 7 weeks. Furthermore,

10 Zhang et al.97 developed a natural silk fibroin polymer based electrospun graft that mimicked the structure of blood vessels with suitable mechanical properties. Human coronary artery smooth muscle cells and human aortic endothelial cells were sequentially seeded onto the luminal surface of the tubular electrospun silk fibroin scaffold and cultured under physiological pulsatile flow conditions. The flow condition provided improved mass transport and aerobic cell metabolism. Under the dynamic culture conditions, enhancement of tissue formation, extracellular matrix production, cell alignment and the retention of differentiated cell phenotype were induced. Moreover, a multilayered hybrid vascular graft with tailored mechanical properties was also reported by Wise et al. by electrospinning human elastin and polycaprolactone blends.98 This mutilayered graft presented a synthetic elastin internal lamina to circulate blood with enhanced endothelial cell interactions and low thrombogenicity, and meanwhile excellent suturability and mechanical durability in a small scale rabbit carotid interposition model.

Nanofibers for neural tissue engineering. The aim of neural tissue engineering is to restore nerve system for patients with injured or diseased nerves. Recently, various electrospun synthetic nerve grafts have been studied as alternatives for autologous nerve grafts.95,97,99-101 Electrospinning is used to fabricate scaffolds for this specific application as a consequence of the production of aligned nanofibers, which greatly influences cell growth and differentiation during never regeneration. For example, Yang et al.95 studied nano/micro scale poly(L-lactic acid) (PLLA) aligned fibers and their potential in neural tissue engineering. Their study demonstrates the possibilities of fabricating aligned PLLA nano/micro fibrous scaffolds by electrospinning technique tailored by the processing parameters. The suitability of PLLA scaffolds for the neural stem cells (NSCs) culture

11 was studied on fibers with different fiber alignment and dimension. It was found that the

NSCs elongated and their neurite grew along the fiber direction for the aligned scaffolds, whereas the fiber diameter did not show any significant effect on the cell orientation.

Furthermore, the NSCs differentiation rate was higher on the nanofibers than that on the micro fibers, but independent with respect to the fiber alignment. All in all, the aligned nanofibers highly supported the NSCs growth, differentiation and improved the neurite outgrowth. This study demonstrated the aligned nanofibrous scaffold as a promising candidate in neural tissue engineering.

Nanofibers for bone tissue engineering. The ideal scaffolds for bone tissue engineering should have comparable mechanical property and 3D porous architecture to native bones. Composite nanofibers comprised of biodegradable polymers and hydroxyapatite (HA) were studied to compositionally and structurally resemble native mineralized collagen fibers and further promote bone regeneration. For example, Cui et al.102 developed a novel strategy to make fibrous composites of hydroxyapatite (HA) and poly(DL-lactide) (PDLLA). Electrospinning of the blends led to the formation of non-stoichiometric nanostructured HA and a good dispersion of HA in the produced

PDLLA fibers. To control the nucleation and growth of HA crystals, gelatin was grafted on the electrospun fibers. The amount of HA formed on the fibrous composites and the crystal size of formed HA can be tuned through the content of gelatin grafted on the fibers. The fibrous nanocomposites showed potential as scaffolds for bone tissue engineering. In another study, Zhang et al.103 reported novel nanocomposite nanofibers comprised of hydroxyapatite/chitosan (HA/CTS) by combining a two-step in situ co-precipitation synthesis method using electrospinning. Poly(ethylene oxide) (PEO)

12 with ultrahigh molecular mass was used as the fiber-forming additive. Model nanocomposite HA/CTS fibers with about 30% HA loading mass was fabricated with proper structural preservation of HA crystallites. The biological evaluation results indicate that the HA/CST nanofibrous scaffolds significantly stimulated the bone forming ability as supported by the cell proliferation, mineral deposition, and morphology observation. As compared to CTS scaffolds alone, it seemed the excellent osteoconductivity of HA helped promote bone formation. This study showed the great potential of using the HA/CTS nanocomposite nanofibers for bone tissue engineering applications.

1.1.3.2 Drug delivery

Similarly to tissue engineering, electrospun nanofibers show great promise in drug delivery as well from their special characteristics.104-106 For example, polymer nanofibers have high surface area to volume ratio per mass unit which provides an efficient method for the delivery of poorly water soluble drugs. Additionally, the drug release rate can be easily tuned by the polymer composition, dimension, and morphology of the fibers. Moreover, the dosage of drug loaded in the nanofiber carrier is facile to manipulate to achieve maximum drug delivery effects. Finally, compared to other drug delivery carriers, electrospun nanofibers have higher drug loading efficiency and capacity.

The first electrospun nanofiber based drug delivery was reported by Kenawy et al. in

2002.107 Polymer nanofibers made either from poly(lactic acid) (PLA), poly(ethylene-co-vinyl acetate) (PEVA), or from a 50:50 blend of the two were studied for the controlled delivery of hydrophobic tetracycline hydrochloride as a model drug.107

13 Release of the drug from these polymer nanofibers was followed by UV-Vis spectroscopy.

Their results demonstrated that PEVA nanofiber based delivery system showed a higher release rate (65% release of tetracycline hydrochloride within 120 hours) when compared with the PLA or the 50/50 PLA/PEVA blend nanofiber meshes (about 50% release for the same time duration). These results indicated that the release rate of loaded drugs in electrospun nanofiber based delivery carriers can be controlled by modulating the ratio of polymer compositions. Another study by Kim et al. demonstrated the successful incorporation and sustained release of a hydrophilic drug from poly(lactide-co-glycolide)

(PLGA) electrospun nanofiber based delivery system without the loss of structure and bioactivity.91 These studies demonstrate that electrospun nanofibers can be used as efficient drug delivery carriers for both hydrophilic and hydrophobic drugs and the drug release rate can be finely tuned by modulation of the polymer composition, size and morphology of the electrospun fibers. Apart from hydrophobic/hydrophilic drugs, there is also a great need for bioactive molecule delivery (e.g., DNA, protein, enzyme, growth factors) for various therapeutic applications. Several challenges for delivering bioactive molecules include low gene transfection efficiency, protein instability and difficulties in release kinetic control.59 Polymer nanofibers, especially those produced from coaxial electrospinning, have been explored for delivering bioactive molecules. Coaxial electrospun nanofibers have a core/sheath structure: the sheath is comprised of polymer and the bioactive molecules are encapsulated in the core. Such a structure has great potential in stabilizing bioactive molecules during the electrospinning process. In addition, the core/sheath structure also allows better release kinetics control of the encapsulated bioactive molecules by modifying the core/sheath material compositions,

14 and morphologies. To date, the delivery of many different types of bioactive molecules including lysozyme,108 platelet-derived growth factor-bb (PDGF-bb),109 nerve growth factor (NGF)110 and fibroblast growth factor (FGF)111 have been studied within coaxially electrospun scaffolds. These studies demonstrated controlled release of the entrapped biomolecules from the electrospun scaffolds with efficient bioactivity to interact with surrounding cells. Besides, blend electrospinning of polymers with biomolecules protected by nanoparticles is also widely studied to achieve effective bioactive molecules delivery.112-114

1.2 Small diameter vascular grafts (SDVGs)

A vascular graft is a synthetic tube that can be used to repair, replace and bypass damaged blood vessels.57 Vascular grafts are used to treat patients with cardiovascular disease. There are three sizes of vascular graft products - large diameter (inner diameter >

10 mm), medium diameter (10 mm > inner diameter > 6 mm) and small diameter (inner diameter < 6 mm).57,115,116 The most challenging construct is small vascular graft applications due to the hydrodynamics within these small size blood vessels (e.g., lower blood flow rate and higher resistance). 63

1.2.1 Clinical needs and challenges for SDVGs

Cardiovascular disease is the number one cause of death worldwide, claiming about 17 million lives each year and accounting for 1/3 of all deaths globally.117 The coronary artery is a small blood vessel (with inner diameter about 3 mm) located on the surface of heart. The coronary artery carries oxygen rich blood and nutrients to the heart.

15 When blocked, severe damage in the form of heart attack results. When this happens, surgery is required to allow for blood flow around the blockage. The most common treatment is to use an autologous venous graft, which is use of the patients own blood vessels from the legs (saphenous veins) or chest (mammary arteries) to bypass the blockage. Autografts are considered a gold standard material for this application because of a low failure rate and lack of immune response.57,115,116 In the United States, more than

750,000 coronary artery bypass surgeries are performed each year.118 However, this surgery has been limited by the availability of the autografts as there are other pre-existing conditions preventing the use of the patients own blood vessels. Synthetic vascular grafts made of materials such as polyethylene terephthalate (PET, Dacron) or expanded polytetrafluoroethylene (ePTFE, Gore-Tex), which are very easy to produce at a large scale with unlimited supply, work well with large and medium size grafts.57,115,116

However, when applying these materials to small diameter applications, there are high failure rates due to the challenging blood flow hydrodynamics within small size blood vessels. Currently, the development of clinically acceptable synthetic small diameter vascular grafts would be transformative.

1.2.2 Resorbable scaffold guided SDVGs tissue engineering

Vascular tissue engineering, which typically involves scaffolds, cells, and/or growth factors to regenerate new blood vessels, provides promising approaches to solve the problem of small diameter vascular grafts.63 There are three different tissue engineering approaches: in vitro, in vivo, and in situ (Figure 1.3.).63 In vitro vascular tissue engineering is the traditional approach to construct functional living vascular grafts

16 outside the body. In this technique, a scaffold is designed and cells are seeded to the material and grown in culture using a bioreactor. The cells will grow on and into the scaffold and begin producing matrix that will create a tissue-like substitute for the damaged blood vessel. This technique requires several weeks to months in order to grow enough cells on the scaffolds before transplantation. In vivo vascular tissue engineering replaces the bioreactor from in vitro engineering with subcutaneous incubation, where the same cell-seeded scaffold is implanted under the skin of the patient to grow and vascularize the tissue substitute. Again, the in vivo vascular tissue engineering technique requires significant time to grow the tissue and incubate it in the patient before being transplanted into the patient. In situ vascular tissue engineering strategy is an alternative approach where a synthetic graft usually made of a functional bioresorbable polymer is implanted directly where it is needed. The functionality of the material will signal the surrounding cells and tissue to populate the scaffold, and as the polymer degrades, a regenerated blood vessel will be formed in its place. The in situ vascular tissue engineering does not involve time consuming cell culture, and can be made for use as a readily available device off-the-shelf.

Among the numerous factors that affect new blood vessel regeneration in vascular engineering, the scaffold material plays a vital role. The three dimensional scaffolds provide structural support and guidance during the new tissue formation, and affect a variety of cellular behaviors, such as adhesion, differentiation, migration, and proliferation.119,120 The bioresorbable materials used for scaffold fabrication during vascular engineering are mainly divided into two categories: natural materials-based and synthetic polymers-based. Natural polymers, such as collagen, gelatin, fibrin, hyaluronic

17 acid, alginate and decellularised matrices, are produced from biological sources.

Therefore, natural materials usually have excellent biological performance and non-toxic degradation.121 However, natural materials show inferior performance in durability and mechanical strength to synthetic polymers. Synthetic polymers, such as poly(lactic acid)

(PLA), poly(glycolic acid) (PGA), poly(lactic-co-glycolic acid)

(PLGA), polyhydroxyalkanoates (PHAs), and their copolymers or blends, have performed better in durability and mechanical properties compared to natural materials.

However, the bulk degradation of the synthetic polymers leads to a decrease in the pH, which causes local acidification and adverse inflammatory reactions.122,123 In addition, synthetic polymers suffer from a lack of bioactivity.124 Recently, hybrid scaffolds, made of synthetic and natural polymers that combine the superior mechanical properties of synthetic polymers with the excellent biological performance of natural materials have been extensively studied.98,122,125,126 In the following sections, a bioresorbable scaffold based on natural materials and synthetic polymers and their use in scaffold-guided vascular tissue engineering is discussed.

18 Figure 1.3.63 Schematic illustration of in vitro, in vivo, and in situ tissue engineering of vascular grafts. Figures reproduced with permission from Ref. (63) (Copyright © 2014

John Wiley and Sons).

1.2.2.1 Natural materials based scaffold

The natural materials used for tissue engineering scaffolds includes polysaccharides (e.g., chitosan, and hyaluronic acid derivate), proteins (e.g., collagen, fibrin, elastin and silk fibroin), or decellularised ECM.127 For vascular tissue engineering, collagen, fibrin, and decellularised ECM are mostly used.57

Collagen. Collagen is the major component of ECM in many body tissues. It supplies mechanical support and maintains biological integrity of the body tissues.128

There are mainly four types of collagen (collagen I, II, III, and V). Collagen shows non-toxicity, biodegradation, low antigenicity and low inflammatory response.128-132

Therefore, scaffolds based on collagen were widely explored in the field of tissue

19 engineering. Among the four different types of collagen, type I collagen is extensively studied for vascular tissue engineering as the vascular wall is mainly composed of collagen type I.128 The first tissue-engineered vascular graft using cell-seeded collagen was demonstrated by Weinberg and Bell in 1986.133 A multilayered structure based on a collagen scaffold resembling an artery was reported. Collagen and bovine smooth muscle cells were casting in a tubular mold. Endothelial cells and fibroblasts were seeded separately into the lumen and outer layer to create a pseudo-intima and a pseudo-adventia.

The resulting graft was weak in mechanical strength and ruptured at very low burst pressures (less than 10 mmHg). A synthetic polyethylene terephthalate (PET) mesh was added to enhance the burst pressure. Experimental results showed promising tissue remodeling with endothelial cell lining in the lumen and the smooth muscle cells in the wall were well differentiated. Since then, vascular scaffolds based on collagen have been extensively reported.129,134-137 One limitation with using collagen fibers and gel for vascular scaffolds application is their low mechanical strength and rigidity. Various methods, including physical and chemical, were investigated to improve the mechanical properties of collagen for its use in vascular scaffolds. The methods include blending collagen fibers with elastin to improve the viscoelasticity (compliance) of the grafts122,138,139 and cross-linking to enhance mechanical strength.140-142

Fibrin. Fibrin is an insoluble body protein largely involved in blood clotting and wound healing.128,143 Fibrin clot is a mesh of fibrillar gel formed by polymerization of fibrinogen which can be purified from patient’s own blood. Therefore, fibrins are widely studied as natural bioresorbable scaffolds since they show a lack of adverse inflammatory reaction at the implantation site.144,145 Fibrin scaffolds are used in the form of fibrin

20 hydrogels, fibrin glue, and fibrin microbeads. When used as gels, fibrin can stimulate smooth muscle cells into synthesizing more elastin, which is one important component of native blood vessels.146 Fibrin based scaffolds used for vascular tissue engineering was reported by Ye et al.138, 139 Three-dimensional constructs were fabricated from fibrin gel.

The degradation rate of the gel was controlled by a proteinase inhibitor called aprotinin, which can slow down or inhibit the process of fibrin degradation. However, the pure fibrin scaffolds suffered from inadequate mechanical strength. To strengthen fibrin scaffolds for vascular engineering, collagen was introduced. Neidert et al. reported by adding TGF-β and insulin either individually or in combination with plasmin, more collagen was secreted and the mechanical property of the fibrin scaffold increased by about 10 times.147 Grassl et al. showed that 148 by optimizing the ratio of TGF-β, insulin and a proteinase inhibitor, fibrin scaffolds showed comparable mechanical properties to a native rat abdominal aorta. Another promising strategy to improve the mechanical properties of fibrin based vascular scaffolds is making composite constructs by mixing fibrin with collagen.139 Collagen stiffens the scaffolds as fibrin is more elastic. Recently, fibrin gel-based scaffolds seeded with vascular cells were studied as tissue engineered small diameter vascular grafts by Swartz et al.149 The implanted grafts integrated well with the native blood vessels and showed similar blood flow rates as native blood vessels.

At 15 weeks post-implantation, the grafts showed excellent tissue remodeling with production of collagen and elastin fibers with orientation of smooth muscle cells perpendicular to the direction of the blood flow.

Decellularised ECM. Extracellular matrixes have three dimensional fibrous structures that provide mechanical support for surrounding tissues and attachment sites

21 for surrounding cells.38-41 When used as scaffolds, ECM plays significant role in tissue engineering by guiding cell behavior, such as cell proliferation, morphogenesis and differentiation.78,81 The source of ECM can be from either animals (xenografts) or humans (homografts). ECMs from humans are more favorable for use as tissue engineering scaffolds, since there is less immune reactions. However, the homografts are limited by availability. Xenografts from animals do not have the limitation of supply but are associated with the risk of immune rejections or disease transmission.150 To reduce or eliminate the immune reaction of ECM obtained from xenografts, ECM from xenogenic tissues (e.g., bovine, sheep, monkeys, pigs, and rabbits) are decellularised.151-153 The decellularization process generally consists of mechanical shaking, chemical detergent treatment, and enzymatic digestion.154 The purpose of decellularization is to remove all cellular and nuclear matter that might cause adverse immune reactions but keep the structure integrity of the remaining ECM.155,156 A general overview of decellularised

ECM vascular tissue engineering process is shown in Figure 1.4.115 Cells are isolated from patients and expanded in cell medium in vitro. Meanwhile, arteries are harvested from a xenogenic source (pigs for example), and decellularized to obtain ECM matrix.

The matrix is seeded with patient’s own cells and cultured in a bioreactor to engineer a fully functional graft before it can be transplanted into patients. Decellularised ECMs from xenografts, such as bovine, pigs and rats, were widely studied in vascular tissue engineering and showed promising results. In one study, Huynh et al. reported a small diameter (4 mm) graft constructed from decellularised porcine small intestinal submucosa and type I bovine collagen.157 The potential of the graft to integrate into the host tissue was evaluated using a rabbit arterial bypass model. Within 3 months after implantation,

22 the implanted grafts demonstrated excellent patency without hyperplasia and active tissue remodeling. In another study, tissue-engineered small-diameter vascular grafts (4 mm) based on a decellularized porcine iliac vessel were studied by Mayer’s group.158

Endothelial progenitor cells (EPCs) that isolated from peripheral blood of sheep were seeded on the decellularized ECM scaffold to make a cellular scaffold prior transplantation. Experiment results in a sheep carotid interposition model demonstrated the EPC-seeded grafts remained patent for 130 days, showing similar properties to native carotid arteries. More interestingly, Dahl et al. recently reported a tissue engineered vascular graft by using human allogeneic or canine smooth muscle cells grown on a tubular polyglycolic acid scaffold.159 The cellular material was later removed from the engineered grafts using a decellularized process to minimize immune reaction. Grafts tested in a dog peripheral and coronary artery bypass model demonstrated excellent patency with no dilatation, calcification, or intimal hyperplasia. These tissue-engineered approaches provide a readily available option for patients who do not have available autologous vessels for the bypass surgery.

23 Figure 1.4.115 Decellularized-matrix vascular tissue engineering. Cells are harvested from a patient, isolated according to type, and expanded in culture. At the same time, arteries are harvested from an allogenic, heterogenic, or xenogenic source, and decellularized to leave a matrix. After recellularizing the matrix with the patient’s cells, the construct matures in a bioreactor to engineer a fully functional graft. Figures reproduced with permission from Ref. (115) (Copyright © 2013 Nature).

1.2.2.2 Synthetic polymers based scaffold

Synthetic biodegradable polymers are widely studied as tissue engineering scaffolds. As compared to natural resorbable materials (e.g., collagen, fibrin, decelluarized ECM), synthetic polymers show better control over mechanical properties, degradation rates (Table 1.1), porosity, etc.57,128,160,161 Furthermore, synthetic polymers can be produced at a large scale with less expensive cost and also have longer storage duality. Among the large number of existing synthetic polymers, aliphatic polyesters

24 composed of glycolide/lactide and their copolymers, including poly(glycolic acid) (PGA), poly(lactic acid) (PLA), poly(ε-caprolactone) (PCL), and poly(lactic-co-glycolic acid)

(PLGA) are the mostly investigated for vascular tissue engineering. Besides, other degradable polymers, such as polyhydroxyalkanoates (PHAs), polyglycerolsebacate

(PGS) are also extensively used as vascular tissue engineering scaffolds.

PGA is a semicrystalline polyester synthesized by the ring-opening polymerization of glycolide.128,132,162 Degradation of PGA occurs through hydrolysis of its ester bonds. When implanted in vivo, the mechanical strength of PGA scaffolds is lost within four weeks and is completely absorbed in about six months.128 The degradation byproducts of PGA are glycolic acids, which can be metabolized and eliminated from the body as water and carbon dioxide.

Similarly, PLA is thermoplastic aliphatic polyester synthesized by ring-opening polymerization of lactic acid.123, 127 Compared to PGA, PLA is more hydrophobic and thus has a slower degradation rate. To finely tailor degradation of the polymers, glycolide is introduced to the polymer structure to make PLGA copolymer. The degradation properties of the resulting copolymers can be finely tuned by varying glycolide and lactide ratios in the repeating units. PLA has two enantiomeric semicrystalline forms, poly(D-lactic acid) (PDLA) and poly(L-lactic acid) (PLLA).163 The properties of PLA are also highly affected by the L/D ratio of the lactate units. Generally, if the L/D ratio is decreased, crystallinity of the polymer increases, and the degradation rate is slower. For example, degradation of PDLA is slower than PLA due to the higher crystallinity of

PDLA.

25 PCL is another semicrystalline, aliphatic polyester synthesized by ring-opening polymerization of -caprolactone. 125,128,132,162,164 PCL degrades slowly in vivo through hydrolysis of its ester bonds followed by release of monomeric hydroxycaproic acid by-products. The hydroxycaproic acid byproducts are ultimately eliminated by macrophages and giant cells.

PHAs are a class of natural polyesters produced by PHA-polymerase catalyzed polymerization in vitro, or by bacterial fermentation of sugar or lipids in vivo.165,166 PHAs can be modified to display a wide range of mechanical and degradation properties.

Among the PHAs, poly(3-hydroxybutyrate) (PHB), copolymers of 3-hydroxybutyrate and

3-hydroxyvalerate (PHBV), poly(4-hydroxybutyrate) (P4HB), copolymers of

3-hydroxybuturate and 3-hydroxyhexanoate (PHBHHx), and poly(3-hydroxyoctanoate)

(PHO) are the extensively studied for tissue engineering applications.167

PGS is a degradable elastomer synthesized by polycondensation of glycerol and sebacic acid.168 PGS degrades by hydrolysis in vivo. Due to the good elastomeric and degradation properties, PGS has been investigated as scaffolds for soft tissue engineering, such as vascular grafts.169-171

Table 1.1. Biodegradable polymers and their degradation properties.172

Polymer Degradation time Degradation products Reference PGA 6-12 months Glycolic acid 173 PLA >24 months L-lactic acid 173 PDLA 12-16 months D,L-lactic acid 173 D,L-lactic acid PDLGA 50:50 2 months 173 Glycolic acid D,L-lactic acid PDLGA 15:85 5 months 173 Glycolic acid

26 PCL >24 months Caproic acid 173 P4HB 2-12 months 4-hydroxbutyrate 174 Collagen ~2 weeks Amino acids 175 Fibrin degradation Fibrin few days 176 products (FDP)

Food and Drug Administration (FDA) approved devices from PGA, PLA, PCL,

PLGA for several human clinical use.128 The FDA also approved devices from polymers containing sebacic acid for several medical applications.170 The first tissue engineered vascular graft based on degradable polymers was reported by Niklason and colleagues,177 in which vascular cells were seeded on a PGA scaffold. After performing culture and biomimetic perfusion in vitro, these grafts were implanted into the right saphenous artery of a pig model. The grafts showed 100% patency 4 weeks post-implantation with a burst pressure strength over 2,000 mmHg and suture retention strength up to 90 grams. Since then, many other studies based on PLA, PCL, PHA, PGS, and their copolymers or blends, have been explored for vascular tissue engineering (Table 1.2).57 Synthetic degradable polymers are promising as vascular engineering scaffolds, but they suffer from a common lack of bioactivity.124 To address this limitation, cell adhesion peptides or growth factors can be incorporated onto the polymer surface or in the polymer bulk to allow sustained release.178-185 Another concern is their acidic degradation by-products. The hydrolysis by-products of polyesters are acidic due to the formation of a carboxylic acid, and this could cause decreased local pH and further inflammation at the implantation site.28, 29, 35

The pH change is also found to be responsible for the abnormal change of surrounding vascular cell phenotype.186-189 Therefore, a new class of resorbable polymers with adequate mechanical properties and non-acidic degradation by-products is highly desired. 27 Table 1.2. Tissue engineered SDVGs fabricated with biodegradable synthetic polymers.57

Materials ID Animal study Reference Model: rat abdominal aorta replacement (12 weeks); Perfect patency with no thrombosis; for grafts with low porosity adventitia, cell PCL 2 mm 190 invasion and neovascularization were reduced; no significant difference between grafts in endothelialization rate Model: rat carotid arterial replacement (18 months); Patency = 72.5%; no 0.7 PCL aneurysm formation; no calcification; 191 mm endothelium; not complete degradation of graft materials Model: rat abdominal aorta replacement (18 months); Excellent structural integrity with no aneurysmal dilation; perfect patency with no PCL 2 mm 164 thrombosis and limited intimal hyperplasia; rapid endothelialization within 6 months; cellular regression is observed after

Model: rat inferior vena cava or aortic interposition (6 weeks); Excellent PGA or PLLA patency without thromboembolic nonwoven felts + 0.7/0.9 complications or aneurysm formation; 192 PCL/PLA sealant mm a foreign body immune response was solution observed in 3 weeks; extensive vascular tissue remodeling observed in 6 weeks Model: rat abdominal aorta replacement (3 months); Rapid host Heparin-coated remodeling in 3 months with no PGS porous tube aneurysm and stenosis; confluent wrapped with a 0.7 endothelium, contractile smooth 170 PCL thin mm muscle layers and expression of electrospun layer elastin/collagen/glycosaminoglycan

were observed; neovessels had tough and compliant mechanical properties

28 1.2.3 Electrospun scaffolds for SDVGs tissue engineering

Ideal scaffolds for small diameter vascular graft tissue engineering should be biocompatible and bioactive. Furthermore, the mechanical properties, degradation properties and microstructure of the scaffolds should be able to mimic the native ECM as closely as possible.193 The electrospinning technique produces nanofibers with very fine size ranging from nanometer to micrometer length scales, which allows mimicking the fibrous structure of ECM. Electrospinning also offers precise control over the mechanical properties and microstructure of the scaffolds by tuning composition, dimension, and alignment of the fibers.1 Natural polymers or bioactive growth factors can be incorporated into the fibers during the electrospinning process or immobilized onto the fiber surface to enhance biological interaction of the scaffolds with the surrounding tissues/cells. Electrospinning has been widely used to fabricate tissue engineering scaffolds. Specifically, tubular scaffolds from various electrospinning techniques (e.g., co-spinning, co-spraying, coaxial setups, and use of hydrogels) are extensively studied for small diameter vascular tissue engineering (Figure 1.5).58 Besides using various electrospinning techniques, a variety of synthetic polymers, natural polymers, or a combination of the two have been electrospun for small diameter vascular tissue engineering and have demonstrated promising results. A summary of electrospun scaffolds for small diameter vascular engineering is shown in Table 1.3.58

29

Figure 1.5.58 Schematics of various experimental setups for the electrospinning process for the fabrication of tubular scaffolds. (a) A multilayer scaffold combining electrospinning and hydrogel fabrication. (b) Co-electrospinning of two polymer solutions concurrently. (c) Electrospinning with simultaneous electrospraying. Figures reproduced with permission from Ref. (58) (Copyright © 2014 Elsevier).

30 Table 1.3. A summary of electrospun scaffolds for SDVGs (ID < 6 mm).58

Electrospun Properties Biology study ID Reference materials in vitro in vivo Model: rabbit carotid Mechanical interposition (1 PCL and properties closely month); elastin matched the human High patency (recombinant 2.8 mm artery, enhanced 98 with no human endothelial cell dilatation, tropoelastin) interaction, improved anastomotic blood compatibility dehiscenc, or seroma Comparable biomechanical PEG- collagen properties to mimetic autologous grafts, protein - 4 mm reduced platelet - 194 hydrogel with adhesion, or electrospun activation, higher PEU endothelia cell migration speed Model: rabbit epigastric-free flap Close elastic modulus (7 weeks); PLLACL to native artery, 1 mm No blood leaking, 96 (70:30) phenotype expression kept structure of endothelia cells integrity during surgery and good patency

Model: carotid artery of adult sheep ( 6 and 12 Superior handling months); PEtU–PDMS 5 mm and compliance High patency 195 characteristics rates, extensive tissue remodeling with no sign of calcification

31 Model: rat Good infrarenal aortic anti-thrombogenic replacement PCL/chitosan property, enhanced (4 weeks); hybrid growth of endothelial 1.5 mm Reduced 196 immobilized cells, while thrombus with with heparin moderately improved suppressed growth of endothelialization smooth muscle cells and patency

Model: rabbit Accelerated carotid artery PLLACL/ endothelia cells replacement chitosan proliferation in the (4 weeks); hybrid loaded first 6 days, and 2.2 mm Extensive 185 with growth modulated slow vascular cells factors (VEGF smooth muscle cells remodeling, no and PDGF) proliferation in the thrombus or burst initial 3 days rupture appeared

1.3 Controlled protein delivery

The first recombinant protein therapeutic, human insulin, was introduced about three decades ago.197 Since then, pharmaceutical proteins, including peptides, recombinant therapeutic proteins, enzymes, monoclonal antibodies, and antibody-drug conjugates have been rapidly explored. So far, more than 200 proteins and peptides, targeting at cancer, inflammatory diseases, vaccines, diagnostics and tissue engineering, have received approval from US Food and Drug Administration (FDA) and over 900 pharmaceutical protein products are currently in development for treating a variety of human diseases.198,199 Pharmaceutical protein drugs account for nearly 20% of the total number of drugs (Medicines in development-biologics, 2013, Pharmaceutical Research and Manufacturers of America Report). The current protein delivery issues and electrospun nanofiber based delivery system are highlighted in the following sections.

32 1.3.1 Pharmaceutical proteins and delivery issues

The delivery of protein drugs is very challenging as protein drugs have:199 1) poor oral bioavailability, 2) inadequate stability and shelf life, 3) immunogenicity, 4) short plasma half-life, and 5) poor penetration across biological membranes. To date, the most frequently used protein drug administration is by parenteral injection. However, due to the short half-life of protein drugs and the fast renal clearance by the body, multiple injections of protein drugs daily or weekly are needed. The frequent injections lead to poor patient compliance and increased protein drug dosage (increased cost of burden as well). Therefore, effective and non-invasive protein delivery approaches are highly desired, and ideally should be able to:200 1) maintain plasma protein-drug concentration over an extended period of time, 2) protect the active therapeutic from premature degradation, 3) enhance drug efficacy, while reducing non-target side effects, and 4) avoid frequent administration and lower drug dosage.

1.3.2 Electrospun nanofibers for protein delivery

The electrospinning technique allows for the production of fine nanofibers that have high surface to volume ratio per mass unit, controlled porosity, and a structure that mimics the fibrous structure of extracellular matrix (ECM).38-41 Recently, electrospinning is used to develop protein delivery systems. Compared to other widely studied delivery systems (e.g., micro/nanoparticles, microspheres, liposomes, hydrogels), electrospun nanofiber based delivery systems showed a number of advantages, including:201-203 1) high drug loading efficiency, 2) facilitated drug diffusion, and 3) improved solubility of various bioactive molecules. Furthermore, the release profile of loaded drugs can be

33 modulated by the biodegradability/hydrophilicity/hydrophobicity of the polymers used, and fiber dimensions or porosity,204 since drug release from the electrospun fibers occurs primarily by diffusion and degradation of the materials used. In addition, to decrease the amount of protein drugs administrated and the non-target delivery toxicity, local delivery of proteins can be achieved by using protein loaded electrospun fibers as an implantable device. Also, to protect the active protein from premature degradation, various electrospinning techniques such as emulsion electrospinning and coaxial electrospinning are available. So far, a number of protein drugs have been loaded to electrospun nanofibers for application in many areas, such as anti-cancer therapy, tissue engineering scaffolds, anti-bacterial dressings.59,205 A summary of electsospun nanofibers based protein delivery system is listed in Table 1.4. In general, biomolecules can be delivered either directly from the electrospun nanofibers or from additional separate carriers (i.e., micro/nanospheres) loaded into the nanofibers, in which the electrospun nanofibers perform only as a supporting structure. Currently, the most frequently explored fabrication techniques for loading proteins to electrospun nanofibers are physical adsorption, blend electrospinning, coaxial electrospinning and covalent immobilization

(Figure 1.6).59

Physical adsorption. In this approach, biomolecules are added to the electrospun nanofibers by dipping into a pure solution or an emulsion of the biomolecules (Figure

1.6(a)). This method is one of the easiest ways to load biomolecules to electrospun nanofibers, and there is minimal bioactivity loss during the absorption process. However, the physical adsorption approach is not favorable in protein drug delivery because of the uncontrolled release profile of the loaded drugs, since all physically absorbed protein

34 drugs are located on the fiber surface and the protein release is in a pure diffusion-controlled release manner.

Blend electrospinning. In blend electrospinning, biomolecules, biomolecules protected in microspheres, or emulsified solutions containing biomolecules are mixed with a polymer solution (Figure 1.6(b)). The mixed solution is electrospun together to produce hybrid nanofibers. In hybrid nanofibers, the biomolecules are located within the fibers. The polymer fiber degradation allows the loaded biomolecules to diffuse into the surrounding environment, and a sustained release profile is expected. However, one significant concern associated with blend electrospinning is the bioactivity loss of biomolecules when exposed to harsh organic solvents. To address this limitation, several strategies have been developed, including using salt complexation, or hydrophilic additives to stabilize the biomolecules.206-208 Recently, other strategies using hydroxy-apatite (HAp) or sugar glass nanoparticles (SGnP) were also reported to avoid denaturation of biomolecules during the hash electrospinning process.112-114 A typical release profile for blend electrospun fibrous scaffolds begins with an initial burst release, followed by a sustained release close to a linear mode. The initial burst release usually occurs within first 24 hours, mostly attributed to biomolecules on the fiber surface via diffusion. After the burst release, the biomolecule release is driven by diffusion or a combination of polymer fiber degradation and diffusion, which means the biomolecule release profile of blend electrospun nanofibers can be modulated by polymer fiber degradation rates, polymer hydrophilicity/hydrophobicity, and fiber dimensions.

Coaxial electrospinning. Coaxial electrospinning, also known as co-electrospinning, uses different capillary channels at the spinneret and coaxially

35 electro-spins the polymer solution and biomolecule solution simultaneously (Figure

1.6(c)). During coaxial electrospinning, a solution of biomolecules is loaded in the inner capillary while polymer solution is loaded in the outer capillary. This experimental setup allows for fabrication of polymer nanofibers with a core-shell structure (core: biomolecules, shell: polymers) and such structure provides a sustained release of loaded biomolecules from the electrospun scaffolds, which is quite similar to that from a blend electrospun scaffold. Furthermore, the drug loading can be modulated by controlling the inner and outer capillary feeding rates. Multi-drug delivery is possible by using the co-electrospinning technique. The one limitation with coaxial electrospinning is the complicated experimental setup, since special designed spinnerets with multiple capillary channels are needed.

Covalent immobilization. Covalent immobilization strategy immobilizes biomoeclules onto the fiber surface via chemical bonds (Figure 1.6(d)). The primary goal of covalent immobilization is to enhance surface properties of electrospun scaffolds.

However, by choosing stimuli-responsive bonds which can be cleaved with certain triggers, e.g., enzyme, this approach can be used for protein delivery with controlled release profiles. Covalent immobilization is not a routine way to delivery biomolecules due to its complexity. However, this strategy provides another option for protein delivery.

36

Figure 1.6.59 Fabrication techniques of bioactive electrospun nanofibers. (a) Physical adsorption. (b) Blend electrospinning. (c) Coaxial electrospinning. (d) Covalent immobilization. Figures reproduced with permission from Ref. (59) (Copyright © 2011

Springer Link).

37 Table 1.4. A summary of protein delivery systems based on electrospun nanofibers.59

Fabrication Loaded Scaffold Results/application Reference technique protein material Physical Serum Enhanced cell attachment PCL 209 adsorption protein and spreading Sustained enzyme release BSA PVA 210 in bioactive form Sustained enzyme release Lysozyme PDLLA 211 in bioactive form Sustained growth factor Blend release, enhanced bone electrospinning bFGF PLGA marrow stem cell 111 attachment and proliferation Aided the wound healing EGF Silk Fibroin by increasing the time of 212 wound closure Modulated protein release BSA PCL 213 from co-electrospun fibers Promoted vascular smooth PDGF-bb PLCL muscle cell attachment 214 and proliferation

Coaxial Promoted human primary keratinocyte and fibroblast electrospinning bFGF, PCL-PEG cells proliferation, 215 EGF enhanced wound closure rates of diabetic ulcers Sustained dual growth VEGF, Chitosan/ factors delivery for blood 185 PDGF PELCL vessel regeneration PEG-PDLL Enhanced human dermal RGD 216 A fibroblast adhesion Covalent Enhanced fiber surface immobilization RGD, properties for Schwann PCL 101 YIGSR cell attachment and alignment

38 1.4 Poly(ester urea)s (PEUs)

Current widely studied degradable polymers for vascular tissue engineering and protein delivery are PGA, PCL, PLA and their copolymers or blends. These materials are limited because they create acidic degradation byproducts that cause inflammation and cell phenotype changes in surrounding tissues.217-222 For vascular tissue engineering, these materials do not have the same mechanical properties as the native blood vessels, and this causes unusual cell activity resulting in intimal hyperplasia and occlusion.3, 10

These challenges typically cause graft failure. In order to solve this issue for vascular engineering and to avoid adverse inflammatory reactions during protein delivery, novel materials that have similar mechanical properties to native tissues and can degrade into non-acidic/non-toxic byproducts are highly desired.

Amino acid-based poly(ester urea)s (PEUs) are a new class of materials that have been developed in recent years. Synthetic PEUs show promise in biomedical applications because of their excellent blood/tissue compatibility, tunable mechanical properties, and nontoxic hydrolysis byproducts.72,223-228 PEUs are comprised of amino acids and diols that are reacted through esterification to form the diester monomers. p-toluene sulfonic acid is added to prevent amidation and exchange reactions during the monomer synthesis.

The monomers can then be coupled with triphosgene via an interfacial polymerization to produce PEU polymer with degradable ester bonds. The final degradation products are amino acids and diols that can readily be metabolized by the body. The urea bonds at every repeat unit neutralize the acidic degradation byproducts. By using different amino acids and diols with different chain lengths, several different types of poly(ester urea)s

39 with different mechanical properties can be created and potentially used for a variety of applications. A general two-step synthesis route for the PEUs is shown in Scheme 1.1.223

Scheme 1.1.223 The two-step general synthetic scheme of amino acid-based poly(ester urea)s (PEUs). In the first step, diol is condensed with 2.1 equivalents of amino acids protonated with p-toluene sulphonic acid to produce monomers. Following the diamine monomer synthesis, interfacial polymerization with triphosgene yields the PEU polymer.

Figures reproduced with permission from Ref. (223) (Copyright © 2015 American

Chemical Society).

40 CHAPTER II

MATERIALS AND INSTRUMENTS

2.1 Materials

All chemical reagents and solvents were purchased from Sigma-Aldrich, Fisher

Scientific, or Alfa Aesar and used as received without additional purification unless specifically noted. Cell culture medium and staining regents were purchased from Lonza

(Basel, Switzerland) or Invitrogen (Carlsbad, CA). BCA (bicinchoninic acid) assays were obtained from Pierce Biotechnology (Rockford, IL). Additional thanks to Prof. Fayez F.

Safadi and Douglas C. Crowder from the Northeast Ohio Medical University (NEOMED) for the donation of platelet rich plasma (PRP), Prof. Hossein Tavana in Biomedical

Engineering at the University of Akron for the source of human umbilical vein endothelial cells (HUVECs) and Dr. Todd Ritzman from the Akron Children’s Hospital for the supply of recombinant human growth hormone (rhGH).

L-leucine: Sigma-Aldrich, 98%

1,6-Hexanediol: Sigma-Aldrich, 99%

1,8-Octanediol: Sigma-Aldrich, 98%

1,10-Decanediol: Sigma-Aldrich, 98%

1,12-Dodecanediol: Sigma-Aldrich, 99%.

p-Toluenesulfonic acid: Fisher Scientific, 99% 41 Triphosgene: Alfa Aesar, 98%

Sodium carbonate (Na2CO3): Fisher Scientific, 99%

Dioctyl sulfosuccinate (AOT): Alfa Aesar, 96%

D-(+)-trehalose dehydrate: Sigma-Aldrich, 99%

Rhodamine B: Sigma-Aldrich, ≥95%

2.2 Instruments

Nuclear Magnetic Resonance (NMR): NMR spectra were obtained using either a

Varian 300 MHz or 500 MHz NMR spectrometer. The solvents were CDCl3 or DMSO-d6.

Data analysis was performed by ACDLABS 12.0 1D NMR Processor software. Chemical shifts were reported in parts per million (δ) and referenced to the chemical shifts of the

1 residual protonated solvent peak ( H NMR: CDCl3 7.27 ppm, DMSO-d6 2.50 ppm) or the

13 central carbon peak of the solvent ( C NMR: CDCl3 77.00 ppm, DMSO-d6 39.50 ppm).

Fourier Transform Infrared (FT-IR): FT-IR spectra were recorded on a Shimadzu

MIRacle 10 ATR-FTIR spectrometer (600 to 4000 cm-1). The resolution was 4 cm-1 and scan numbers were 64. Data analysis was performed by Win-IP Pro and plotted using

Origin 8.1 software. Samples were prepared either by solvent casting or compression molding with KBr.

Size Exclusion Chromatography (SEC): Size exclusion chromatography (SEC) analyses were performed using a TOSOH HLC-8320 gel permeation chromatograph equipped with a refractive index (RI) detector. The eluent is 10 mM LiBr in DMF at a flow rate of 0.8 mL/min. The column and detector temperatures were maintained at 50 °C.

42 The molecular mass and mass distribution were calculated from a universal calibration based on polystyrene standards.

UV-Visible Spectroscopy (UV-Vis): UV-Vis absorption tests were performed on a

Synergy MX spectrophotometer (Biotek Inc.) using a quartz 96 well plate. 300 μL solution was used for each sample. Absorbance at 562 nm was measured. The data experiments were controlled and data was analyzed by BioTeck’s Gen5TM Data Analysis and plotted using Origin 8.1 software.

Quartz Crystal Microbalance (QCM): A Q-sense E4 operator from Biolin

Scientific AB was used to study protein adsorption on polymer thin films. Polymers (0.5 wt% in dioxane) were spin-coated on SiO2-coated crystal sensors X301 (5 MHz resonant frequency) at 2000 rpm for 1 min, and the acceleration time was 10 seconds. The thickness of the thin films was measured using spectroscopic ellipsometry. All experiments were performed at 20 ± 0.1 °C using a flow rate of 0.15 mL/min. The concentration of protein solutions was typically 1 mg/mL. In a typical experiment, the polymer-coated QCM sensors were pre-soaked in PBS buffer overnight to reach a hydrated equilibrium. Afterwards, the baseline was established over a 20 min interval.

The protein solution was applied to the QCM sensors in a continuous flow until a plateau was reached. Subsequently, continuous flow of protein solution was applied for 30 min.

After that, the sensors were rinsed thoroughly with PBS buffer for 20 min. The normalized frequency of the third overtone was used for analysis. Based on Sauerbrey model,229 the frequency shift was converted into the mass change per area according to the following equation (where Δm is the adsorbed mass on the surface, C is the mass

43 sensitivity constant which is 17.7 ng·cm−2·Hz-1, Δf is the resonance frequency change at the nth harmonic, and n is the overtone number):

m - f n

Thermal Analysis: All samples were vacuum dried thoroughly at 60 °C prior to the thermal property testing. The thermal stability of polymers was measured using thermogravimetric analysis (TGA, TA Q500) across a temperature range of 0 - 600 °C at a scanning rate of 20 °C/min under nitrogen. Decomposition temperature data at 5% weight loss was collected. The thermal transitions of the polymers were studied by using differential scanning calorimetry (DSC, TA Q2000) from -20 to 120 °C at a scanning rate of 20 °C/min. Equilibrium time at highest/lowest temperature was 2 min. Approximately

10 mg sample was used for the testing. The glass transition temperature (Tg) was determined as the midpoint in the second heating cycle of DSC.

Destructive Tensile Testing: To determine the elastic properties of the polymers, tensile tests were performed using an Instron machine (Instron 5567 universal testing machine with temperature control chamber). Thin films of each polymer were fabricated using vacuum melt-compression (TMP Technical Machine Products Corp.). Each large film was cut using an ASTM stamp into small rectangular samples with dimensions of 40 mm in length, 5 mm in width, and 0.5 mm in thickness for testing. The gauge length and crosshead speed were set as 20 mm and 5 mm/min respectively. The tensile tests were studied both at room temperature (20 ± 1 °C) and physiological temperature (37 ± 1 °C).

Stress-strain data was reported using the Instron Bluehill software. The data was plotted using Origin 8.1 and elastic modulus values were calculated as the slope of the tangent

44 line of the data curve prior to the yield point. Results presented are average values for three individual measurements (n=3).

Scanning Electron Microscopy (SEM): SEM testing was performed using field emission SEM (JSM-7401F, JEOL, Peabody, MA). The acceleration voltage for SEM imaging was 5 kV. Samples were vacuum dried overnight, stabilized on an aluminum cylinder SEM mount using conductive tapes and sputter coated with silver for testing.

Transmission Electron Microscopy (TEM): Bright field TEM images were recorded on a JEOL-1230 TEM microscope with an accelerating voltage of 120 kV using a digital CCD camera. For samples dispersed in organic solvents, 20 µL of sample diluted solution was dropped onto a 300 mesh copper TEM grid (PELCO, Redding, CA). Excess sample solution was removed by filter paper and the sample was dried under vacuum for overnight.

Light Scattering (LS): The hydrodynamic radius of nanoparticles was determined by LS using a Brookhaven light scattering spectrometer (BI-200SM) equipped with a temperature-controlled solid state laser (λ 532 nm). A CONTIN method was used to obtain the average hydrodynamic radius of the particles in solution. The sample solutions were filtered through a 0.45 µm syringe membrane filter to remove any dust or aggregated particles and equilibrated at room temperature for 30 minutes before the measurements.

Fluorescence Microscopy Testing: Florescence Images were recorded on a IX 81 microscope (Olympus, Center Valley, PA) and are unaltered. Fluorescent mages were analyzed with the ImageJ software to determine average cell area.

45 CHAPTER III

L-LEUCINE BASED POLY(ESTER UREA)S FOR VASCULAR TISSUE

ENGINEERING

Portions of this work have been published previously as

Yaohua Gao, Erin Childers, and Matthew L. Becker

ACS Biomater. Sci. Eng., 2015, 1, 795-804

3.1 Outline

Resorbable poly(ester urea)s (PEUs) derived from α-amino acids are promising for vascular engineering applications. The objective of this work was to synthesize and characterize L-leucine-based PEUs and evaluate their suitability for vascular tissue engineering. Four different PEUs were prepared from di-p-toluenesulfonic acid salts of bis-L-leucine esters and triphosgene using interfacial condensation polymerizations.

Mechanical testing indicated that the elastic moduli of the respective polymers were strongly dependent on the chain length of diols in the monomers. Three of the resulting

PEUs showed elastic moduli that fall within the range of native blood vessels (0.16 MPa to 12 MPa). The in vitro degradation assays over 6 months indicated that the polymers are surface eroding and no significant pH drop was observed during the degradation process.

Human umbilical vein endothelial cells (HUVECs) and A-10 smooth muscle cells (A-10

SMCs) were cultured on PEU thin films. Protein adsorption studies showed the PEUs did 46 not induce the rupture of platelets in platelet rich plasma (PRP) after pretreatment with fibrinogen. Taken together, our data suggest that the L-leucine-based PEUs are viable candidate materials for use in vascular tissue engineering applications.

3.2 Introduction

Over the past few decades, biodegradable polymers have been extensively studied as resorbable scaffold materials for vascular tissue engineering applications.63,230-233

Among these, aliphatic polyesters including poly(ε-caprolactone) (PCL),125,164,190,234-236 poly(lactic acid) (PLA),161,237 poly(glycolide) (PGA),177,238,239 poly(glycerol sebacate)

(PGS)169-171 and their blends or copolymers192,240-243 are the most widely investigated family of degradable polymers. However, polyesters have shown several limitations in vascular engineering studies (e.g. slow degradation, localized acidification, mechanical property mismatch, etc.).217,218,244,245 For example, the degradation of PCL with an initial molecular mass of 55,000 g/mol showed no significant mass loss until after 80 weeks in vivo.244 Capsules of linear PCL with an initial molecular mass of 66,000 g/mol remained intact for 24 months in rats, and retained their mechanical strength for 30 months.245

Vascular grafts engineered from slow degrading polymers typically show limited host cell infiltration and remodeling even 6-12 months after implantation.170 Attempts to tune the degradation rate via copolymerization or blending PCL with other polymers have yielded mixed results.246-250 PGA generally degrades much faster than PCL. However, the main concern for with regard to PGA is that the intermediate degradation products lead to a localized pH drop, which in turn induces an inflammatory reaction at the implant site.217-219 Furthermore, the acidic degradation byproducts of these polymers have been

47 reported to be cytotoxic and negatively affect the vascular smooth muscle cell (VSMC) phenotype.220-222 Intimal hyperplasia and graft rupture owing to mechanical property mismatch between implanted scaffold materials and native blood vessels are also considered as leading causes of graft failures.251 Significant efforts have been made to diversify the pool of degradable polymers to meet design criteria for vascular tissue engineering applications.

Amino acid-based PEUs are synthetic biodegradable polymers that have shown promise in biomaterials and regenerative medicine applications.224,227,228,252,253 They are amino acid-based and degradable with tunable mechanical properties and bioactivity.72,225,226 Phenylalanine-based PEU (poly(1-PHE-6)) was shown to be resorbable and nontoxic in subcutaneous rat implant models.72 No evidence of local acidification-induced inflammation was observed when poly(1-PHE-6) degraded in vivo.

Moreover, previous work has shown that the physical, chemical and degradation properties of PEU polymers are influenced by specific amino acids and diol lengths in the monomers (i.e., increasing chain diol length in the monomer increases the amount of flexible segment in the polymer structures, which thus results in reduced elastic modulus values and increased elongation at break).227 In vitro degradation studies showed that

PEUs degrade faster than PLLA, losing approximately 5 wt% after 112 days at physiologically temperature in PBS. Careful tuning of polymer composition affords optimization of physical, chemical and degradation properties of PEUs for specific applications. Leucine (abbreviated as LEU) is a naturally occurring branched-chain

α-amino acid. In this study, we report the synthesis, characterization, and degradation study of PEUs derived from L-leucine and linear diols of different length. Furthermore,

48 we measured the protein adsorption, in vitro behavior of human umbilical vein endothelial cells (HUVECs) and A-10 smooth muscle cells (A-10 SMCs) on the PEU thin films. Our preliminary results showed that L-leucine derived PEUs are promising candidates for vascular tissue engineering.

3.3 Experimental Section

The polymer synthesis, characterization and property testing are highlighted in the following sections. The mechanical, thermal and degradation properties of synthesized polymers are fully studied by a series of tests.

3.3.1 Materials and Cells

All the reagents and solvents were purchased from Sigma Aldrich or Alfa Aesar and used without further purification unless otherwise noted. Chloroform was dried with

CaH2 overnight and distilled before use. All reactions were performed under protection of nitrogen unless noted otherwise. A-10 smooth muscle cells (CRL-1476, ATCC, Rockville,

MD) were maintained for less than 20 passages in Dulbecco’s Modified Eagle’s Medium

(DMEM) supplemented with 10 % (v/v) fetal bovine serum (FBS), 100 units mL-1 penicillin, 100 μg mL-1 streptomycin, and 2 mmol L-1 L-glutamine (all from Invitrogen,

Carlsbad, CA). HUVECs (C2517A, Lonza, Basel, Switzerland) were maintained for less than 20 passages in endothelial growth medium kit (EGM-2 BulletKit, CC-3156 &

CC-4176, Lonza, Basel, Switzerland) supplemented with 100 units mL-1 penicillin

(Invitrogen, Carlsbad, CA). Rhodamine Phalloidin and DAPI (4′,

6-diamidino-2-phenylindole) dyes were also from Invitrogen. Cytoskeleton stabilization

49 (CS) buffer was prepared using 1,4-piperazinediethanesulfonic acid (PIPES, 0.1 M)

(Fisher Scientific, Pittsburgh, PA), ethylene glycol-bis(2-aminoethylether)-

N,N,N′,N′-tetraacetic acid (EGTA, 1 mM) (Sigma Aldrich, St. Louis, MO), and 4% (w/v) polyethylene glycol (PEG, Mw =8000) (Fisher Scientific, Pittsburgh, PA) in distilled water. The solution was buffered using 1 M hydrochloric acid and 1 M sodium hydroxide solutions as required to obtain a pH of 6.9.

3.3.2 Instrumentation

1H and 13C NMR spectra of the monomers and polymers were obtained on Varian

NMRS 500 spectrometers. Chemical shifts were reported in parts per million (δ) and

1 referenced to the chemical shifts of the residual solvent ( H NMR, DMSO-d6 (2.50 ppm);

13 C, DMSO-d6 (39.50 ppm)). The following abbreviations were used to explain the multiplicities: s=singlet, d=doublet, t=triplet, br=broad singlet, m=multiplet. FTIR spectra were recorded on a Shimadzu MIRacle 10 ATR-FTIR spectrometer. Size exclusion chromatography (SEC) analyses were performed using a TOSOH HLC-8320 gel permeation chromatograph using 10 mM LiBr in DMF as eluent at a flow rate of 0.8 mL/min. The column and detector temperatures were maintained at 50 °C. The molecular mass and mass distribution were calculated from a universal calibration based on polystyrene standards. The thermal stability of the polymers was measured using thermogravimetric analysis (TGA, TA Q500) across a temperature range of 0-600 °C at a scanning rate of 20 °C/min under nitrogen. 5% decomposition temperature data was collected. The thermal transitions of the polymers were studied by using differential scanning calorimetry (DSC, TA Q2000) from -20 to 120 °C at a scanning rate of

50 20 °C/min and dynamic mechanical analysis (DMA, TA Q800) by sweeping the temperature from -20 to 80 °C at a scanning rate of 3 °C/min and a frequency of 1 Hz.

The glass transition temperature was determined as the midpoint in the second heating cycle of DSC and the maximum of tan δ in DMA.

3.3.3 Synthesis of di-p-toluenesulfonic acid salts of bis-L-leucine diester monomer

Di-p-toluenesulfonic acid salts of bis-L-leucine diester monomers were prepared according to previous published methods,72,227 as shown in Scheme 3.1. In brief, to a 2 L three-neck round-bottom flask equipped with a Dean-Stark apparatus and a magnetic stirrer was added with diol (0.34 mol, 1.0 equiv), α-amino acids (0.78 mol，2.3 equiv), p-toluenesulfonic acid (0.81 mol，2.4 equiv), and toluene (1 L). The reaction mixture was refluxed for 16 h until no more water was produced. Then, it was cooled down to room temperature and the toluene was evaporated under reduced pressure. The resulting

L-leucine-based diesters were recrystallized four times from water. The yield was about

75-90%. The chemical structures were confirmed via 1H NMR, 13C NMR, Fourier transform infrared (FT-IR) spectroscopy (Figure 3.1, Figure 3.2, and Figure 3.3).

Scheme 3.1. A two-step general synthetic route of the L-leucine-based poly(ester urea)s with diols of different chain length (n=3, P(1-LEU-6); n=4, P(1-LEU-8); n=5,

P(1-LEU-10); n=6, P(1-LEU-12)).a 51 aThe general two-step synthetic route of L-leucine-based poly(ester urea)s (PEU) with diols of different length. In the first step, 1 e.q. of diol is condensed with 2.3 e.q. of amino acids to yield the monomers. During this step, the amino acids are protonated with p-toluene sulphonic acid to prevent amidation reactions. In the second step, interfacial polycondensation of monomers with triphosgene is conducted yielding PEU polymers.

Di-p-toluenesulfonic Acid Salt of Bis-L-leucine-1,6-diester. 1H NMR (500 MHz,

DMSO- d6): 0.89 (m, 12H) 1.33 (m, 4H) 1.57-1.62 (m, 8H) 1.71 (m, 2H) 2.28 (s, 6H)

2.50 (m, DMSO) 3.30 (s, H2O) 3.95 (t, 2H) 4.12-4.16 (m, 4H) 7.09-7.11 (d, 4H)

13 7.46-7.48 (d, 4H) 8.24 (s, 6H). C NMR (500 MHz, DMSO-d6): 21.22, 22.37, 22.58,

24.26, 25.20, 28.22, 39.68-40.35 (DMSO-d6), 51.09, 65.97, 125.93, 128.51, 138.20,

145.91, 170.36. FT-IR (cm-1): 1740 [-C(CO)-O-].

Di-p-toluenesulfonic Acid Salt of Bis-L-leucine-1,8-diester. 1H NMR (500 MHz,

DMSO- d6): 0.89 (m, 12H) 1.28 (m, 4H) 1.56-1.63 (m, 8H) 1.71 (m, 2H) 2.27 (s, 6H)

2.50 (m, DMSO) 3.30 (s, H2O) 3.96 (t, 2H) 4.13-4.15 (m, 4H) 7.08-7.11 (d, 4H)

13 7.45-7.48 (d, 4H) 8.24 (s, 6H). C NMR (500 MHz, DMSO-d6): 21.22, 22.37, 22.58,

24.26, 25.58, 28.32, 28.89, 39.68-40.35 (DMSO-d6), 51.09, 66.06, 125.93, 128.51,

138.18, 145.94, 170.38. FT-IR (cm-1): 1740 [-C(CO)-O-].

Di-p-toluenesulfonic Acid Salt of Bis-L-leucine-1,10-diester. 1H NMR (500 MHz,

DMSO- d6): 0.89 (m, 12H) 1.25 (m, 4H) 1.55-1.62 (m, 8H) 1.71 (m, 2H) 2.27 (s, 6H)

2.50 (m, DMSO) 3.30 (s, H2O) 3.96 (t, 2H) 4.13-4.17 (m, 4H) 7.08-7.11 (d, 4H)

13 7.45-7.47 (d, 4H) 8.23 (s, 6H). C NMR (500 MHz, DMSO-d6): 21.22, 22.36, 22.58,

52 24.26, 25.67, 28.34, 29.00, 29.30, 39.68-40.35 (DMSO-d6), 51.09, 66.09, 125.94, 128.50,

138.17, 145.94, 170.38. FT-IR (cm-1): 1740 [-C(CO)-O-].

Di-p-toluenesulfonic Acid Salt of Bis-L-leucine-1,12-diester. 1H NMR (500 MHz,

DMSO-d6): 0.89 (m, 12H) 1.24 (m, 4H) 1.58-1.66 (m, 8H) 1.71 (m, 2H) 2.28 (s, 6H) 2.50

(m, DMSO) 3.30 (s, H2O) 3.96 (t, 2H) 4.13-4.17 (m, 4H) 7.09-7.12 (d, 4H) 7.45-7.48 (d,

13 4H) 8.24 (s, 6H). C NMR (500 MHz, DMSO-d6): 21.22, 22.36, 22.57, 24.26, 25.66,

28.34, 29.01, 29.39, 39.68-40.35 (DMSO-d6), 51.09, 66.09, 125.94, 128.50, 138.16,

145.96, 170.38. FT-IR (cm-1): 1740 [-C(CO)-O-].

1 Figure 3.1. H NMR (DMSO-d6) of di-p-toluenesulfonic acid salts of bis-L-leucine- diester monomers. 1H NMR spectra of the four monomers showed differences in intensity at 1.24-1.33 ppm that corresponds to the methylene units in the respective diol chains.

53

13 Figure 3.2. C NMR (DMSO-d6) of di-p-toluenesulfonic acid salts of bis-L-leucine- diester monomers.

Figure 3.3. FT-IR of di-p-toluenesulfonic acid salts of bis-L-leucine-diester monomers.

54 3.3.4 Synthesis of L-leucine-based poly(ester urea)s using interfacial polymerization

The PEU polymers were synthesized using interfacial polymerization as depicted in Scheme 3.1. Di-p-toluenesulfonic acid salt of bis-L-leucine diester monomers (0.038 mol, 1.0 equiv), sodium carbonate anhydrate (0.080 mol, 2.1 equiv), and distilled water

(400 mL) were added into a 2 L three-neck round-bottom flask equipped with an overhead mechanical stirrer and a thermometer. The mixture was then heated with a warm water bath at 50 °C for 30 mins. The water bath was then removed and replaced with an ice-salt bath. When the solution temperature cooled to ~ 0 °C, additional sodium carbonate (0.040 mol, 1.05 equiv) dissolved in 150 mL of distilled water was added to the mixture. Several minutes later, prepared triphosgene solution (0.013 mol, 1.05/3 equiv.) dissolved in 100 mL of distilled chloroform was added to the reaction system as quickly as possible (< 5 s) with fast mechanical stirring. The reaction was allowed to proceed for

30 min and then additional aliquots of triphosgene (0.003 mol, 0.25/3 equiv.) dissolved in

30 mL of distilled chloroform were added slowly over 30 min. After the addition was complete, stirring was continued for 2 h. Later, the organic phase was precipitated into hot water, filtered and dried in vacuum to yield a white solid. In total, four different PEUs were synthesized. The yield was ~ 85-95%. The PEU polymers are named according to the first three letters of the amino acids and the number of carbons in the diols used. For example, P(1-LEU-6) indicates poly(ester urea) synthesized from L-leucine and 1,6- hexanediol. The polymers were then further characterized by 1H NMR (Figure 3.4), 13C

NMR (Figure 3.5), Fourier transform infrared (FT-IR) spectroscopy (Figure 3.6), size exclusion chromatography (SEC), thermogravimetric analysis (TGA) (Figure 3.7), differential scanning calorimetry (DSC) (Figure 3.8) and dynamic mechanical analysis

55 (DMA) (Figure 3.9). The molecular mass and thermal properties of the polymers are summarized in Table 3.1.

Synthesis of 1,6-Hexanediol-L-leucine-Based Poly(ester urea) (P(1-LEU-6). 1H

NMR (500 MHz, DMSO-d6): 0.83-0.88 (m, 12H), 1.30 (m, 4H), 1.39-1.45 (m, 4H),

1.52-1.55 (t, 4H), 1.58-1.65 (m, 2H), 2.50 (DMSO), 3.95-4.04 (m, 4H), 4.10-4.15 (m,

13 2H), 6.29-6.31 (d, 2H). C NMR (500 MHz, DMSO-d6): 22.10, 23.07, 24.71, 25.30,

-1 28.41, 39.51-40.52 (DMSO-d6), 41.44, 51.49, 64.49, 157.51, 173.84. FT-IR (cm ): 1734

[-C(CO)-O-], 1647, 1548 [-NH-C(O)-NH-], 3370 [-NH-C(O)-NH-].

Synthesis of 1,8-Octanediol-L-leucine-Based Poly(ester urea) (P(1-LEU-8). 1H

NMR (500 MHz, DMSO-d6): 0.83-0.87 (m, 12H), 1.25 (m, 4H), 1.41-1.44 (m, 4H),

1.52-1.54 (t, 4H), 1.58-1.62 (m, 2H), 2.50 (DMSO), 3.36 (H2O), 3.96-4.02 (m, 4H),

13 4.11-4.12 (m, 2H), 6.27-6.29 (d, 2H). C NMR (500 MHz, DMSO-d6): 22.11, 23.06,

24.72, 25.64, 28.51, 28.94, 39.52-40.52 (DMSO-d6), 41.47, 51.50, 64.56, 157.50, 173.83.

FT-IR (cm-1): 1734 [-C(CO)-O-], 1635, 1548 [-NH-C(O)-NH-], 3355 [-NH-C(O)-NH-].

Synthesis of 1,10-Decanediol-L-leucine-Based Poly(ester urea) (P(1-LEU-10). 1H

NMR (500 MHz, DMSO-d6): 0.83-0.90 (m, 12H), 1.24 (m, 4H), 1.41-1.44 (m, 4H),

1.51-1.54 (t, 4H), 1.58-1.62 (m, 2H), 2.50 (DMSO), 3.28 (H2O), 3.97-4.01 (m, 4H),

13 4.11-4.14 (m, 2H), 6.26-6.28 (d, 2H). C NMR (500 MHz, DMSO-d6): 22.11, 23.06,

24.72, 25.70, 28.52, 29.04, 29.29, 39.17-40.84 (DMSO-d6), 41.43, 51.51, 64.57, 157.50,

173.84. FT-IR (cm-1): 1740 [-C(CO)-O-], 1640, 1555 [-NH-C(O)-NH-], 3355

[-NH-C(O)-NH-].

Synthesis of 1,12-Dodecanediol-L-leucine-Based Poly(ester urea) (P(1-LEU-12).

1 H NMR (500 MHz, DMSO-d6): 0.83-0.89 (m, 12H), 1.23 (m, 4H), 1.39-1.47 (m, 4H),

56 1.51-1.52 (t, 4H), 1.60-1.63 (m, 2H), 2.50 (DMSO), 3.29 (H2O), 3.97-4.01 (m, 4H),

13 4.09-4.13 (m, 2H), 6.28-6.30 (d, 2H). C NMR (500 MHz, DMSO-d6): 22.18, 22.78,

24.77, 25.75, 28.47, 29.09, 29.37, 39.17-40.84 (DMSO-d6), 42.43, 51.66, 65.37, 157.01,

174.63. FT-IR (cm-1): 1734 [-C(CO)-O-], 1635, 1555 [-NH-C(O)-NH-], 3355

[-NH-C(O)-NH-].

3.3.5 Mechanical property measurements

To determine the elastic properties of the PEU polymers, tensile tests were performed using an Instron machine (Instron 5567 universal testing machine with temperature control chamber). Thin films of each polymer were fabricated using vacuum melt-compression (TMP Technical Machine Products Corp.). Each large film was cut using an ASTM stamp into small rectangular samples with dimensions of 40 mm in length, 5 mm in width, and 0.5 mm in thickness for testing. The gauge length and crosshead speed were set as 20 mm and 5 mm/min respectively. The tensile tests were studied both at room temperature (20 ± 1 °C) and physiological temperature (37 ± 1 °C).

Stress-strain data was reported using the Instron Bluehill software. The data was plotted using Origin 8.1 and elastic modulus values were calculated as the slope of the tangent line of the data curve prior to the yield point. Results presented are average values for three individual measurements (n=3).

3.3.6 In vitro degradation tests

The in vitro hydrolytic degradation measurements were performed using ASTM standard F1635-11 “in vitro degradation testing of hydrolytically degradable polymer

57 resins and fabricated forms for surgical implants” (ASTM International, West

Conshohocken, Pa, USA, 2004). The degradation studies were conducted on regular samples prepared using melt-compression molding with dimension size of 20 mm×10 mm×5 mm at 37 °C in pH 7.4 sodium phosphate buffer (PBS). The samples were soaked in glass vials containing 20 mL of PBS buffer. At specified intervals, the samples were removed from the PBS buffer and rinsed thoroughly with distilled water, dried for several days to constant weight, weighed, and tested. The mass percent changes and molecular mass changes were recorded. The surface morphology changes during degradation were evaluated using field emission SEM (JSM-7401F, JEOL, Peabody, MA). The results are the average values of three individual measurements for each sample at each time point

(n=3). Error bars represent one standard deviation from the mean.

3.3.7 Water uptake study

Water uptake was determined to evaluate the in vitro hydrolytic degradation rate of PEU polymers (Figure 3.11b and Table 3.3). Regular samples prepared using vacuum melt-compression molding with dimension size of 20mm×10mm×5mm were used for water uptake studies. The samples were weighed (W0) and then immersed in PBS buffer for 1 week. Subsequently, the samples were taken out of the PBS buffer, wiped dry with tissue paper, and weighed again immediately (Wt). The percentage of water uptake was calculated using the following equation:

Water uptake (%) = (Wt - W0)/W0×100%

58 3.3.8 Quartz crystal microbalance (QCM)

A Q-sense E4 operator from Biolin Scientific AB was used to study the protein adsorption property of leucine based PEUs. PEU polymers were spin-coated on the

SiO2-coated crystal sensor X301 (5 MHz resonant frequency) at 2000 rpm for 1 min, and the acceleration time was 10 s. The thickness of the thin films was measured using spectroscopic ellipsometry as described in Table 3.4. All experiments were performed at

20 ± 0.1 °C using a flow rate of 0.15 mL/min. The concentration of protein solutions was

1 mg/mL. In a typical experiment, the polymer-coated QCM sensors were pre-soaked in

PBS buffer overnight to reach a hydrated equilibrium. Afterwards, the baseline was established over a 20 min interval. The protein solution was applied to the QCM sensors in a continuous flow until a plateau was reached. Subsequently, continuous flow of PRP in PBS was applied for 30 min. After that, the sensors were rinsed thoroughly with PBS buffer for 20 min. The normalized frequency of the third overtone was used for analysis.

Based on Sauerbrey model,229 the frequency shift was converted into the mass change per area according to the following equation:

m - f n where Δm is the adsorbed mass on the surface, C is the mass sensitivity constant which is

17.7 ng·cm−2·Hz-1, Δf is the resonance frequency change at the nth harmonic, and n is the overtone number.

3.3.9 Cell attachment and spreading studies

Human umbilical vein endothelial cells (C2517A, Lonza, Basel, Switzerland) and

A-10 smooth muscle cells (CRL-1476, ATCC, Rockville, MD) between passage 10 and 59 14 were used to evaluate cell attachment and spreading on PEU thin films. Blank glass coverslips studied as positive controls and glass coverslips with spin-coated PEU polymers were placed in a 12 well plate, sterilized with ethylene oxide and then wet with corresponding cell media for 4 h. HUVECs or A-10 smooth muscle cells were evenly seeded to each sample at a cell density of 6500 cells/cm2 and cultured for 48 h in a 37 °C,

5% CO2 incubator. Cells were then fixed by 3.7 % paraformaldehyde in cytoskeleton stabilization (CS) buffer for 10 min and permeabilized with 0.5% TritonX-100 for 9 min.

Excess formaldehyde was quenched with 0.05% sodium borohydride in PBS. 5% donkey serum in PBS was then added and incubated for 20 mins to block the non-specific binding activity. The actin filaments of the cytoskeleton were then stained with rhodamine phalloidin (1:200 dilution in PBS) for 1 h. After three time rinse with PBS, the nucleus was stained with DAPI (1:1000 dilution in PBS) for 40 mins and washed four times with PBS before taking fluorescent pictures. The samples were viewed with IX 81 microscope (Olympus, Center Valley, PA) with 10×, 20× and 40× objectives. Images were analyzed with the Image J software to determine average cell area.

3.3.10 Statistics

All experiments were conducted with three replicates (n= 3). A one-way analysis of variance (ANOVA) with Tukey post hoc analysis was conducted when applicable. A significance value of p > 0.05 was set for all statistical analysis. All quantitative data are reported as the mean ± standard deviation.

60 3.4 Results and Discussion

The effect of chain length of diols in the polymer repeating structure on the mechanical, thermal and degradation properties of the materials are discussed in the follow sections.

3.4.1 Polymer synthesis and characterization

The PEU polymers were prepared from 1,6-hexanediol, 1,8-octanediol,

1,10-decanediol, 1,12-dodecanediol L-leucine based bis-ester monomers and triphosgene using interfacial polymerization according to published methods.225 L-leucine was chosen because of its bulky aliphatic side chain. Compared with amino acids with more rigid aromatic side chains (e.g. L- phenylalanine), it provides PEUs with more flexibility. The structures of the synthesized PEUs were confirmed by 1H NMR (Figure 3.4), 13C NMR

(Figure 3.5) and FT-IR spectroscopy (Figure 3.6). 1H NMR spectra of the four polymers showed differences in intensity at 1.23-1.30 ppm that corresponds to the methylene units in the respective diol chains. When comparing 13C NMR spectra of the four polymers, differences from 20 to 30 ppm correspond to the methylene units in the respective diol chains were observed. The FT-IR spectra showed the characteristic urea and ester peaks, e.g. strong N-H (urea) stretch at around 3200-3500 cm-1, symmetric N-C (amide I) stretch around 1640 cm-1, asymmetric N-C (amide II) stretch around 1550 cm-1 and strong C=O

(ester) stretch around 1740 cm-1 were observed. Taken together, the 1H NMR, 13C NMR and FT-IR results indicate PEU polymers were synthesized with high purities.

61 Table 3.1. Characterization data summary of the L-Leucine-based PEUs.

T /°C T /°C T /°C Samples M d g g w (TGA) (DSC) (DMA) P(1-LEU-12) 103,000 1.9 279 23 38

P(1-LEU-10) 135,000 1.9 273 30 46

P(1-LEU-8) 131,000 1.7 272 37 50

P(1-LEU-6) 55,000 1.7 270 42 62

1 Figure 3.4. H NMR (DMSO-d6) spectra of L-leucine-based poly(ester urea)s.

62 13 Figure 3.5. C NMR (DMSO-d6) spectra of L-leucine-based poly(ester urea)s.

Figure 3.6. FT-IR spectra of L-leucine-based poly(ester urea)s.

63 In this work, interfacial polymerization was chosen to prepare the polymers because it allows the synthesis of linear polymers with high molecular mass.254 As can be seen from Table 3.1, the molecular mass (Mw) of the polymers were generally over 100 kDa with polydispersity indices (PDI) varying from 1.7 to 1.9. For P(1-LEU-6), it was more likely to obtain Mw around 55 kDa. This was unexpected as the polymerization method was the same for all PEUs and we anticipated P(1-LEU-6) to have similar high molecular weights as other PEUs. Though repeated experiments were conducted, consistent results were obtained. The volumetric ratio of aqueous phase and organic phase during interfacial polymerization and the relative solubility of P(1-LEU-6) is believed to be responsible. Despite this difference, all the PEUs showed sufficiently high molecular mass to fabricate stable polymer films for the mechanical property and degradation tests, as well as cell culture studies.

3.4.2 Thermal properties

TGA results (Table 3.1, Figure 3.7) showed that the onset degradation temperatures of all four L-leucine based PEUs are around 270 °C with only small difference due to their similar chemical structures. For all four polymers, the degradation temperature, Td ~ 270 °C, is much higher than their glass transition temperature (23 to

42 °C ). The DSC and DMA results (Table 3.1, Figure 3.8, Figure 3.9 and Appendix

Figure 1) showed that the PEUs generally have larger values of Tg than most current widely studied biodegradable polymers (e.g. PCL, ~ -60 °C). The change of carbon number in the diols from 6 to 12 led to a noticeable Tg difference (42 °C vs. 23 °C). With increasing the length of diols from 1, 6-hexanediol to 1, 12-dodecanediol, the glass

64 transition temperature of resulting PEUs decreased from 42 °C to 23 °C, as longer diol chains provides additional chain flexibility to the backbone of polymers and thus results

227 in lower Tg value. A copolymerization strategy between mixtures of the monomer precursors will further tune the thermal and mechanical properties of the polymers based on the respective composition. Taken together, the TGA, DSC and DMA data demonstrated that the leucine based PEUs can be melt processed to fabricate coatings or scaffolds for tissue engineering applications or other devices without degradation at a high temperature.

Figure 3.7. TGA analysis of L-leucine-based poly(ester urea)s.

65 Figure 3.8. DSC trace of L-leucine-based poly(ester urea)s.

Figure 3.9. DMA results of L-leucine-based poly(ester urea)s. 66 3.4.3 Mechanical properties

To further characterize the PEU polymers and evaluate their suitability for use in vascular tissue engineering applications, the mechanical properties of each PEU polymer were measured by tensile testing (Instron). Measurements were conducted under two distinct conditions: room temperature (20 ± 1 °C) and physiological temperature. The elastic modulus (E’) of each polymer was obtained from the stress-strain curve generated from the tensile testing. E’ was calculated as the slope of the initial linear region of the stress-strain curve prior to the yielding point (Figure 3.10). Mechanical property dada was summarized in Table 3.2.

The elastic moduli are statistically different from each other, indicating that all four polymers have unique mechanical properties. Besides, the change of the length of diols led to a noticeable mechanical properties difference. With increasing the length of diols from 1, 6-hexanediol to 1, 12-dodecanediol, the elastic modulus of PEUs decreases from 1.2 GPa to 12 MPa. The possible explanation is, in PEU polymer structures, the ester part is considered as a soft segment while the urea part is considered as a hard segment. Increasing the chain length of diols will increase the relative amount of soft segments in the polymer, and thus leads to decrease in elastic modulus and increase of elongation at break. When the testing temperatures were increased from room temperature (20 ± 1 °C) to physiologic temperature (37 ± 1 °C), the mechanical property of all four polymers changed significantly in the meantime. The elastic modulus dropped significantly and the elongations at break increased dramatically as the temperature shift approached or exceeded Tg. More importantly, except P(1-LEU-6), the elastic moduli of all other three PEUs fell within the range of native blood vessels (i.e. 0.16 MPa to 12

67 MPa),255-257 which is what we expected, since the close the mechanical properties of graft materials come to that of native blood vessels, the less the chance of graft failure due to mechanical property mismatch.

Figure 3.10. Stress-strain plots of L-leucine based PEUs measured at room temperature

(20 ± 1 °C) (a, b)) and physiological temperature (37 ± 1 °C) (c, d) using an Instron 3365 universal materials testing machine. The gauge length was set as 20 mm and the crosshead speed was set at 5 mm min-1. The testing samples were with dimensions of 40 mm×5 mm×0.5 mm. The elastic moduli of the polymers were calculated as the slope of the tangent line of the data curve prior to yield point. Results presented are average values for three individual measurements (n=3). Tensile testing results (i.e., elastic modulus, elongation at break, ultimate tensile stress) are summarized in Table 3.2. 68 Table 3.2. Mechanical properties of L-leucine-based PEUs with diols of different length.

P(1-LEU-12) P(1-LEU-10) P(1-LEU-8) P(1-LEU-6)

E’/MPa 12.0 ± 0.4 134 ± 9 651 ± 58 1208 ± 66

R.T. Strain/% 672 ± 20 343 ± 14 283 ± 32 2.1 ± 0.4a

Stress/MPa 0.5 ± 0.1 10.8 ± 1.2 16.4 ± 2.0 13.7 ± 2.5a

E’/MPa 0.8 ± 0.3 1.6 ± 0.4 5.6 ± 1.8 31 ± 9

37 °C Strain/% >2000b >2000b >2000b 364 ± 61

Stress/MPa 0.06 ± 0.02 0.2 ± 0.1 0.5 ± 0.1 6.8 ± 0.1 aP(1-LEU-6) are brittle, elongation break occurred at the clamps of Instron. bSample elongation reached the limit of the tensile test machine and did not break.

3.4.4 In vitro degradation studies

The in vitro hydrolytic degradation of the leucine based PEUs were carried out at

37 °C in PBS buffer (pH = 7.4). Figure 3.11(a) shows that the weight losses of the polymers during degradation over 24 weeks were minimal, which means the in vitro hydrolytic degradation of leucine based PEUs is generally slow. Though slow, compared to the widely studied biodegradable polymers, e.g. PLLA, PEUs showed faster degradation with tunable rates. For example, it takes approximately 413 days for PLLA to degrade 0.5 wt % in PBS solution (pH = 7.4) at 37 °C and 595 days to degrade 5 wt %,258,259 while it takes 105 days for the leucine based PEUs to degrade 0.5 or 5 wt % depending on the length of diols in polymer structures. When comparing hydrolytic degradation of the four leucine based PEUs with diols of different length in polymer structure, P(1-LEU-6) exhibited the fastest degradation rate followed by P(1-LEU-12),

P(1-LEU-8) and P(1-LEU-10). The hydrolytic degradation rate is believed directly 69 correlating to the extent of water uptake: the faster the extent of water uptake, the higher the polymer degradation rate.227 Water uptake study indicated P(1-LEU-6) has the highest extent of water uptake at the same time point (Figure 3.11(b),Table 3.3), followed by

P(1-LEU-12), P(1-LEU-8), and P(1-LEU-10), which explained the differences of degradation rate with four PEUs. However, due to limited extents of water uptake for all

PEU polymers, PEUs generally shows slow hydrolytic degradation in vitro. This could be a drawback of this kind of materials used for vascular tissue engineering since fast material degradation coordinating to the neotissue remolding is very important. However, the situation is, when implanted in vivo, there is enzymatic degradation of the polymers in addition to hydrolytic degradation due to the ester bonds in the polymer structure. The overall degradation of the PEUs might be greatly accelerated in vivo. On the other hand, the buffer pH was monitored during the degradation study. No significant pH drop was observed over 24 weeks (Figure 3.11(c)), indicating the acidic and basic degradation products have a buffering effect that will not cause local acidification, which is another highly desirable advantage of this kind of materials over other extensively studied polymers as vascular scaffold materials. Surface morphology changes during degradation were studied by SEM. Figure 3.11(d) show the surface morphology of PEU films before and after 4, 8, 16, 24 weeks of degradation, respectively. The results are consistent with a surface erosion degradation mechanism as we proposed for the phenylalanine based

PEUs in our previous work.227 At the beginning of the experiment, the surface is relatively flat, and the initial surface morphology is induced by the melt-compression mold. After 4 weeks hydrolytic degradation, a porous structure on the film surface was

70 observed, indicating hydrolytic attack at the surface is occurring. After 8, 16 and 24 weeks, the severity and frequency of surface porosity and cracks increased with the time.

Table 3.3. Water uptake of L-leucine-based poly(ester urea)s with diols of different length

after being soaked in PBS buffer at 37 °C for one week.

Samples Water uptake (%)

P(1-LEU-12) 9.7 ± 0.4

P(1-LEU-10) 7.6 ± 0.2

P(1-LEU-8) 8.2 ± 0.3

P(1-LEU-6) 16.4 ± 0.5

71 Figure 3.11. In vitro degradation tests were performed according to ASTM standard

F1635-11: “In vitro degradation testing of hydrolytically degradable polymer resins and fabricated forms for surgical implants”. The dimensions of testing samples were of 20 mm×10 mm×5 mm. Results presented are the average values for three individual measurements (n=3). (a) percentage weight loss with respect to incubation time. (b) water uptake of different PEU polymers after soaking in PBS at 37 °C for a week. (c) pH value of PBS buffer of different PEU polymers during degradation. (d) SEM images of PEU films degraded in PBS at 37 °C for up to 24 weeks (scale bar = 30 μm).

72 3.4.5 Protein adsorption

Protein and PRP adsorption property of the leucine based PEUs were monitored by QCM. A decrease in frequency during the experiment indicates an increase in mass due to adsorption on the surface of the QCM sensor. The increase of mass directly correlates to protein adsorption.260-262 When a protein solution was applied, adsorption occurred rapidly on the PEU coated QCM sensors, yielding a significant negative frequency shift compared to the PBS baseline (Figure 3.12). Amount of adsorbed protein was calculated from frequency shift based on the Sauerbrey Model,229 as reported in

Table 3.4. The data shows PEU polymer has a stronger adsorption to fibrinogen, about three times of that of BSA. Following the protein adsorption, PRP was subsequently applied. Quantitative data calculated on the basis of Sauerbrey Model shows there is only tiny amount of PRP adsorption on the fibrinogen pre-adsorbed PEU film (~56 ng·cm-2,

1/7 of the amount adsorbed on BSA pre-adsorbed PEU film), indicating that pre-absorbed fibrinogen on the PEU film can block the adsorption of PRP for somehow. As platelet adhesion and activation are believed to contribute to surface-induced thrombosis, which is also considered as one of the main factors that leads to the failure of vascular grafts,

PEU materials pretreated with fibrinogen will likely block the PRP adhesion to the polymer surface, decreasing the risk of thrombus formation and thus increase the chance of vascular graft success.

73 Figure 3.12. Protein adsorption properties of PEU monitored by QCM-d. The experiment contains four processes: (i) baseline in PBS buffer; (ii) adsorption of proteins; (iii) adsorption of PRP; (iv) PBS buffer washing. The shift in QCM frequency corresponds to adsorption on the polymer surface. A decrease in frequency indicates a mass increase due to adsorption, and the extent of decrease is directly proportional to the amount of increased mass. Results presented are the average values for three individual measurements (n= 3). The data are summarized in Table 3.4.

Table 3.4. Summary of protein and PRP adsorption on PEU surface.

Film Protein PRP Δf Δf Samples thickness 1 2 adsorption adsorption (- Hz) (- Hz) (nm) (ng·cm-2) (ng·cm-2) PEU 56 44 ± 6 66 ± 8 257 ± 38 386 ± 46 (BSA treated) PEU 54 146 ± 4 10 ± 2 861 ± 25 56 ± 13 (Fibrinogen treated)

74 3.4.6 Cell culture studies

In order to further evaluate to what extent the leucine based PEU materials support vascular cell adhesion, cell attachment and spreading studies were performed on

PEU thin films with human umbilical vein endothelial cells (HUVECs) and A-10 smooth muscle cells (SMCs). A-10 cells, a putative vascular SMC line isolated from rat thoracic aorta, were used here for no special reason, just because it is a commonly used model of vascular smooth muscle cells and they are available in the lab. Following culture in vitro, cells were fixed and stained for nuclei and F-actin to examine cellular morphology and adhesion. After 48 h incubation, both HUVECs and A-10 SMCs were found to behave in a similar fashion: spread and attached on all PEU films regardless of the polymer and cell types used (Figure 3.13).

75

Figure 3.13. A-10 smooth muscle cells (A-10 SMCs) and human umbilical vein endothelial cells (HUVECs) attachment and spreading on PEU thin films (glass coverslips were studied as positive control). Cell attachment and spreading are evident for all polymers up to 48 h culture time (Cell seeding density: 6500 cm-2; Red: F-actin stained by rhodamine phalloidin; Blue: nucleus stained by DAPI).

76 Quantification of cell spread area revealed that cells spread comparably with those cultured on control glass coverslips, which means cells put down attachments and stretched out on the PEU materials as they normally would on glass coverslips.

Obviously, the L-leucine based PEUs support to promote adhesion and spreading of

HUVEC endothelial cells and A-10 smooth muscle cells. This finding is significant since cell adhesion and spreading are the first events that dictate the subsequent cellular responses such as proliferation, migration and matrix deposition.254 It is important that the leucine based PEUs are able to enhance these initial events.

3.5 Conclusion

Four PEUs derived from amino acid L-leucine were synthesized using interfacial polymerization with high yields. The chemical structures, thermal and mechanical properties and in vitro degradation of the resulting polymers were characterized.

Leucine based PEUs show a wider tunable range of thermal, mechanical and degradation properties than most commonly used degradable biomaterials. The tensile test results demonstrated that three of the four PEUs showed elastic moduli that fell within the range of native blood vessels. The in vitro hydrolytic degradation study indicated that the PEU polymers are degradable and the degradation rates are faster than the published values of many currently wide studied degradable polymers, such as PLLA, PCL. More importantly, there is no significant pH decrease during the degradation process. The protein adsorption study showed this kind of materials had PRP blocking effect after pre-adsorbing fibrinogen. Vascular cells, such as A-10 SMCs and HUVECs, were found

77 to be spreading and attached on the PEU thin film. In summary, the leucine based PEUs are promising candidates for vascular tissue engineering applications.

3.6 Acknowledgements

The authors are thankful for financial support from the Akron Functional

Materials Center. Drs. Fei Lin, Jinjun Zhou and Mary Beth Wade are acknowledged for useful discussions and comments on the manuscript. Additional thanks to Prof. Fayez F.

Safadi and Douglas C. Crowder from the Northeast Ohio Medical University (NEOMED) for the donation of PRP, and Prof. Hossein Tavana in Biomedical Engineering at the

University of Akron for the source of HUVECs.

78 CHAPTER IV

PILOT RAT STUDY OF 1 MM INNER DIAMETER (ID) VASCULAR GRAFT

USING ELECTROSPUN POLY(ESTER UREA) NANOFIBERS

This work is not published yet

Yaohua Gao, Tai Yi, Toshi Shinoka , Darrel H. Reneker, Christopher Breuer, Matthew L.

Becker, in preparation, 2016

4.1 Outline

An off-the-shelf, small diameter tissue engineered vascular graft (TEVG) would be transformative to surgeons in multiple subspecialties. Herein we report the results of a small diameter (ID≈1 mm) vascular graft constructed from resorbable, amino acid-based poly(ester urea) (PEU). Electrospun PEU grafts of two different wall thicknesses (type A:

250 μm; type B: 350 μm) were implanted as abdominal infra-renal aortic grafts in a severe compromised immune deficient (SCID) mouse model and evaluated for vessel remodeling over 1 year. Significantly, the small diameter TEVG did not rupture or lead to acute thrombogenic events in the intervals tested. The pilot TEVG in vivo showed long term patency and extensive tissue remodeling with type A grafts. Extensive tissue remodeling in type A grafts lead to the development of well-circumscribed neovessels

79 with an endothelial inner lining, a neointima containing smooth muscle cells. However, due to slow degradation of the PEU scaffold materials in vivo, the grafts remained after 1 year. The type B grafts, which have 350 μm thick walls, experienced occlusion over the 1 year interval due to intimal hyperplasia. This study affords significant findings that will guide the design of future generations of small diameter vascular grafts.

4.2 Introduction

Cardiovascular diseases resulting from arteriosclerosis are the leading cause of death globally.263 In the United States, more than 500,000 coronary bypass surgeries are performed each year.118,264 Autologous grafts from the saphenous vein or the internal mammary artery are the gold standard for arterial bypass surgery.265,266 However, the procedures have drawbacks including limited tissue availability, the need for additional surgeries, donor site morbidity, and a ∼30% 10-year failure rate.63 Synthetic vascular grafts, such as those made of expanded polytetrafluoroethylene (ePTFE) and woven or knitted polyethylene terephthalate (PET, Dacron) have been clinically available for quite some time. While clearly innovative solutions, the clinic utility of these synthetic material has been limited to vascular grafts (> 5 mm ID) in areas where the blood flow rate is high and resistance is low. When applied historically to small diameter (<5 mm ID) vessel replacements, e.g. coronary bypass surgery, both ePTFE and Dacron show high failure rates due to acute thrombus formation, and chronic anastomotic (excessive tissue formation in adjacent arterial tissue) and/or intimal (excessive tissue ingrowth through the graft wall) hyperplasia.251,267-269 This significantly reduces the availability of these approaches to most patient population who need them. Therefore, development of

80 clinically acceptable small diameter vascular grafts as an alternative to autologous artery or vein substitutes is of significant clinical interest.

Tissue engineered vascular grafts (TEVGs) with bioresorbable scaffolds is considered as one of the most promising approaches to address clinical challenges in small diameter vessels. Bioresorbable vascular grafts are analogous to biostable vascular grafts, such as those made of ePTFE or Dacron except one major difference: the materials used are degradable and allow the neoartery to remodel in vivo.251,256,264 Ideally, the implanted graft should progressively degrade in a timeframe that allows compensation by extracellular matrix (ECM) deposition and remodeling associated with the natural arterial regeneration process.164 This approach combines the advantages of a synthetic graft (e.g. availability, manufacturing, sterilization, storage, etc.) and the excellent long term performance of natural tissues (e.g. diminished risk of immune rejection, optimal biological and structural properties, etc.).195

Over many years, substantial work by many groups utilizing synthetic degradable polymers, including poly(lactic acid) (PLA),161,237,270 poly(glycolic acid) (PGA),192,271-273 poly(caprolactone) (PCL),164,183,235,274 poly(glycerol sebacate) (PGS),169-171,275 poly(ester amide)s,222,254,276,277 polyurethanes,195,278,279 natural polymers (e.g. collagen,134,280 silk fibroin281-283), and their copolymers or blends,196,284-287 have been reported for tissue engineered small diameter vascular grafts. The Breuer and Shinoka lab has developed the first tissue engineered small diameter vascular graft to be used in humans.288-290 This vascular graft, which is created by seeding autologous bone marrow derived mononuclear cells (BMMCs) onto a biodegradable PLLA/PCLA tubular scaffold, forms a living vascular conduit with the ability to grow, repair and remodel. The first clinical trial

81 evaluating the use of this TEVG in humans followed.291-293 Initial results demonstrated an excellent safety profile for the TEVG, again with the additional benefit of growth potential for use in children where somatic overgrowth is a significant barrier to progress in the field. Moving forward, a cell-free TEVG for arterial circulation constructed from

PLCL and reinforced by PLA fiber mesh was fabricated to enhance cell migration into the scaffolds.294 The process of vessel remodeling with implanting these grafts in a mouse abdominal aorta model during a 12 month period demonstrated organized neotissue formation. However, due to the uncoordinated scaffold degradation time with new vessel remodeling and the acidic scaffold degradation byproducts, most grafts experienced aneurysmal change and prolonged inflammation. In addition, intimal hyperplasia and graft rupture resulting from mechanical mismatch between implanted grafts and native vessels, acute clotting, and limited tissue regeneration are believed to be responsible for the graft failures.251,264 New biodegradable polymers with tunable degradation rate and mechanical properties that can match the remodeling process of a bioresorbable vascular graft in situ are very desirable.

We have explored a series of amino acid based poly(ester urea)s (PEUs), that showed tunable degradation with non-toxic byproducts.72,223-228 The mechanical property results, in vitro and in vivo studies showed these materials might be suitable candidates for vascular tissue engineering.223 In this study, PEU derived from L-leucine and

1,10-decanediol, named as Poly(1-LEU-10), was used to fabricate porous grafts with very small diameter (ID≈1 mm). Electrospinning was used to fabricate the 1 mm inner diameter grafts as it affords the reliable and consistent generation of fibrous structure that resembles extracellular matrix (ECM) in the native vessel wall with no residual solvent. 82 The structures also support cell infiltration and cellularization of the grafts.3,295-297 Small

(1 mm) diameter graft tubes with two different wall thicknesses (i.e. type A: 250 μm; type

B: 350 μm) were fabricated. Biomechanical properties of the grafts were studied in vitro.

The grafts were then studied in vivo in a SCID mouse abdominal aorta replacement model for long term evaluation. This study aims to develop a cell free resorbable small diameter vascular graft based on electrospun PEU and asses its efficiency in vivo.

4.3 Experimental section

Fabrication of the small diameter vascular grafts using electrospinning, biomechanical property and biological activity testing of the engineered grafts are highlighted in the following section.

4.3.1 Materials

Unless listed otherwise, all chemical solvents and reagents were purchased from

Sigma-Aldrich or Alfa Aesar and used as received. Cell culture and studies were conducted following previous published paper.223

4.3.2 Polymer synthesis and characterization

The poly(ester urea) monomer and polymer were prepared following previous

223 1 methods, as shown in Figure 4.1. Poly(1-LEU-10): H NMR (500 MHz, DMSO-d6):

0.83-0.90 (m, 12H) 1.24 (m, 4H) 1.41-1.44 (m, 4H) 1.51-1.54 (t, 4H) 1.58-1.62 (m, 2H)

13 2.50 (DMSO) 3.28 (H2O) 3.97-4.01 (m, 4H) 4.11-4.14 (m, 2H) 6.26-6.28 (d, 2H). C

NMR (500 MHz, DMSO-d6): 22.11, 23.06, 24.72, 25.70, 28.52, 29.04, 29.29,

83 -1 39.17-40.84 (DMSO-d6), 41.43, 51.51, 64.57, 157.50, 173.84. FT-IR (cm ): 1740

[-C(CO)-O-], 1640, 1555 [-NH-C(O)-NH-], 3355 [-NH-C(O)-NH-]; Mn = 71 kDa, Mw =

135 kDa. Td = 273 °C , Tg = 30 °C . The characterization data summary of polymer molecular mass and thermal properties is listed in Table 4.1.

4.3.3 Graft fabrication

Vascular grafts were fabricated by electrospinning using a 10 wt% PEU solution in hexafluoroisopropanol (HFIP). The electrospinning set-up included a syringe pump, a high voltage supply, and a rotating mandrel. A 10 kV positive voltage was applied to the polymer solution by the power supply. The polymer solution was drawn through a 23 gauge blunt tip needle at a constant flow rate of 1 mL/h. Polymer fibers were then collected on a grounded rotating mandrel mounted on a homemade stand. The collecting mandrel was a stainless steel rod with approximately 1 mm diameter. The collecting mandrel was pre-coated with sugar solution. The distance between the syringe tip and the mandrel was set as 15 cm and the mandrel rotation rate was 100 rpm. To remove the graft from the mandrel, the graft together with the mandrel was soaked in deionized water for one hour. When the thin layer of sugar was dissolved by water, the graft can be easily removed from the mandrel by gently pulling it from one direction. The obtained grafts were then freeze-dried and stored in clean glass vials for future use. Prior to implantation, the grafts were sterilized by ethylene oxide (ETO) for 24 hours. Two types of PEU grafts with two different wall thicknesses were prepared (type A: 250 μm; type B: 350 μm).

84 4.3.4 Graft characterization

The electrospun grafts were characterized using field emission scanning electron microscopy (FE-SEM; JSM-7401F, JEOL Ltd., Japan). Characterization included determining the average fiber diameter and average pore area. For each sample, ten SEM images were analyzed, and at least 50 fibers chosen randomly from across the image were manually measured on each image and analyzed using ImageJ software (NIH USA,

2008). Pore areas were also measured by a subjective approximation of surface pores from the SEM images (at least 20 measurements per image). Results are given as mean ± standard deviation. For all the measurements made from the SEM images, calibration of the ImageJ software was done with the scale bar on each image.

4.3.5 Biomechanical evaluations

The biomechanical evaluations of the electrospun grafts provide important information for surgical implantation. In this section, tensile property, suture retention strength and burst pressure strength testing are included.

4.3.5.1 Tensile properties

Uniaxial tensile testing of electrospun grafts was performed on six 1 mm inner diameter tubular specimens from six different electrospun grafts using an Instron 5567 universal tensile testing machine. After soaking the specimens in PBS for 24 h at 37 °C, tensile properties were measured by clamping a 20 mm long graft in the tensile-testing machine and pulling the samples until failure. The gauge length was set as 10 mm, and the crosshead speed was set at 10 mm/min. Stress-strain data were reported using the

85 Instron Bluehill software. The data were plotted using Origin 8.1 and the ultimate tensile strength, modulus, and strain at break were calculated. Results presented are average values for three individual measurements.

4.3.5.2 Suture retention strength

Suture retention testing was performed on six 1 mm inner diameter tubular specimens from six different electrospun grafts according to the procedure described in

Section 8.8 of the American National Standards Institute (ANSI)/Association for the

Advancement of Medical Instrumentation (AAMI) ANSI/AAMI VP20:1994 entitled

“ ardiovascular Implants-Vascular Graft Prostheses”:122 After soaking the grafts in PBS for 24 h at 37 °C, one end of the graft was fixed to the stage clamp of the uniaxial tensile testing machine (Instron 5567, USA) and the other end was connected to another clamp by a loop of a common suture material (5-0 Prolene, Ethicon Inc.) placed 2 mm from the edge of the free end of the graft. The gauge length was set as 20 mm, and the crosshead speed was set at 150 mm/min until the suture ripped or the graft failed. Suture retention strength, which was defined as fracture strength, was recorded in Newton using the

Instron Bluehill software.

4.3.5.3 Burst pressure strength

The burst pressure strength for the electrospun grafts was measured by increasing the pressure within the tubular vascular graft until failure occurred (in our case, the pressure level reached the limitation of the machine before graft failed). Luer-lock needle adapters with matching size of the testing grafts were inserted and fixed by superglue to

86 both ends of the grafts. A pressure transducer catheter which connected to computer was attached to one end of the grafts via the luer-lock needle adapters. A 100 mL pressure syringe was attached to the other end of the grafts. The pressure was gradually increased until reaching the limitation of the machine and the pressure change was recorded on computer.

4.3.6 Biological activity evaluations

For the cell culture studies, PEU nanofibers of identical dimensions were electrospun onto glass coverslips to form two-dimensional structures. The PEU nanofiber covered glass coverslips were then placed into 12-well plates, sterilized by ETO for 24 h, and pre-soaked for 4 h in cell culture medium prior to cell seeding. Blank glass coverslips were studied as positive control. A-10 smooth muscle cells (A-10 SMCs) and human umbilical vein endothelial cells (HUVECs) between passage 10 and 14 were used and seeded directly on the surface of the glass coverslips at a density of 2 × 104 per well. The cell-seeded coverslips were incubated for 4 h to allow cells to adhere to the nanofibers before adding additional cell culture medium to the culture plate. Samples of separate studies were all done in triplicate to assure reproducibility of the results. A-10 SMCs were cultured with Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with

10% fetal bovine serum (FBS) and 1% penicillin and streptomycin. HUVECs were cultured with EGM™-2 Bulletkit™ (Lonza) with basal medium, growth factors, cytokines, and supplements special for endothelial cells. Both cell types were cultured at

37 °C in a humidified incubator containing 5% CO2 for scheduled time. The cell culture medium was changed every 48 hours.

87 4.3.6.1 Cell viability and proliferation study

Cell viability and proliferation were evaluated after 1, 3 and 7 days of cell seeding using PrestoBlue assay. Upon entering a living cell, PrestoBlue® reagent is reduced from resazurin, a blue compound with no intrinsic fluorescent value, to resorufin which is red in color and highly fluorescent. Cell proliferation was assessed by the intensity of red color obtained, which was directly proportional to the metabolic activity of the cell population. At scheduled time points (day 1, 3, and 7), cell culture medium was removed.

Cell seeded coverslips were transferred to empty 12-well culture plates and refilled by 1 mL of fresh cell medium containing 10% v/v of PrestoBlue. After 0.5 to 2 h of incubation at 37 °C, 3 × 100 μL of medium was taken from each well to a 96-well plate and analyzed for fluorescence measurement. The fluorescence intensity was measured on a SynergyTM

MX plate reader from BioTek at an excitation wavelength of 560 nm and an emission wavelength of 590 nm. The observed fluorescence intensity was then converted to cell numbers according to established calibration curves.

4.3.6.2 Cell attachment and spreading study

To study cell attachment and spreading on the scaffold material, A-10 SMCs and

HUVECs with cell density of 78 mm-2 were seeded directly on the surface of the PEU nanofiber covered glass coverslips in 12-well culture plates and were cultured for 48 h before fixation and immunostaining. For immunostaining studies, cells were first fixed by

3.7% paraformaldehyde in CS buffer for 10 min on dry block and then permeabilized with 0.5% TritonX-100 for 9 min. Excess formaldehyde was quenched with 0.05%

88 sodium borohydride in PBS. 5% donkey serum in CS buffer was then added and the well plate was incubated at room temperature for 20 mins to block the non-specific binding activity. The actin filaments of the cytoskeleton were then stained with rhodamine phalloidin (1:200 dilution in PBS) for 1 h. After three time rinse with PBS, the nucleus was stained with DAPI (1:1000 dilution in PBS) for 20 mins and washed four times with

PBS. Coverslips were mounted on microscope slides with mounting medium for fluorescence (Vector Laboratories Inc. Burlingame, CA) and sealed with nail enamel upon drying. Fluorescent pictures were taken using IX 81 microscope (Olympus, Center

Valley, PA) with 10×, 20× and 40× objectives. ImageJ software was used to determine average cell spreading areas.

4.3.7 Animal study

All animals received humane care in compliance with the National Institutes of

Health (NIH) Guide for the Care and Use of Laboratory Animals. The Institutional

Animal Care and Use Committee (IACUC) at Nationwide hildren’s Hospital approved the use of animals and all procedures described in this study. 8-week old female SCID/Bg mice were purchased from Taconic Biosciences (Hudson, NY).

4.3.7.1 Graft implantation

Five Poly(1-LEU-10) grafts were implanted as infra-renal aortic interposition conduits using microsurgical technique. The mice were anesthetized using ketamine xylazine cocktail with ketoprofen as pre-anesthesia analgesic. The hair in the surgical area was removed by a shaver and then disinfected by betadine and alcohol pads. A

89 midline laparotomy incision from below the xyphoid to the suprapubic region was made, and a self-retaining retractor inserted. The intestines were wrapped in saline-moistened gauze and retracted. The infrarenal aorta and inferior vena cava were bluntly defined.

Microsurgery was performed using an operating microscope with zoom magnification.

The aorta was separated from the inferior vena cava and vascular control was obtained with microvascular clamps and then the infra-renal aorta was transected. A

Poly(1-LEU-10) aortic interposition graft was implanted with proximal and distal end-to-end anastomoses using a sterile 10-0 monofilament suture on tapered needles. Any hemorrhage was controlled by applying topical absorbable sterile hemostatic agents

(Surgicel). The intestines were returned to the abdominal cavity. The abdominal musculature and skin were closed in two layers using 6-0 prolene suture. The length of the procedure was 35 minutes. Motrin water was provided for 48 hours after surgery.

Animals were followed for 9 weeks and 12 months after implantation. Post-operatively, no drugs such as anti-platelet or anti-coagulant agents were used.

4.3.7.2 Ultrasound

Serial ultrasonography (Vevo Visualsonics 770; Visualsonics, Toronto, ON,

Canada) was used to monitor grafts after implantation. Prior to ultrasonography, mice were anesthetized with 1.5% inhaled isoflurane. Graft luminal diameter was determined sonographically at the indicated time points after implantation and patency was determined by measuring flow velocity proximal and distal to the graft.

90 4.3.7.3 Contrast-enhanced micro-CT angiography

Under anesthesia, in vivo micro-CT angiography was performed with the GE eXplore Locus in vivo micro-CT scanner (GE Healthcare, Milwaukee, WI, USA).

Micro-CT data were acquired with an x-ray source of 70 kVp tube voltage, 32 mA tube current, 4×4 detector binning model, 16 milliseconds exposure per frame, 70 gain, and 20 offset for contrast-enhanced CT acquisitions. One minute prior to acquisition, animals were given an intra-jugular 0.3 cc bolus of Ultravist (370 mgI/ml, Bayer Healthcare,

Wayne, NJ). A single frame of 220 projections for 42 seconds of continuous x-ray exposure was used. Volumetric microCT images were reconstructed in a 360 × 185 × 505 format with voxel dimensions of 98.4 ×98.4 × 98.4 μm3 using a Feldkamp algorithm with calibrated Hounsfield units (HU). Micro-CT data was transferred to the Advanced

Workstation (version 4.4; GE Healthcare) for further reconstruction and quantitative analysis. Sites of anastomosis were approximated. Measurements of graft length, inner luminal diameter, and graft volume were performed. Similar measurements were performed on adjacent aortas in mice implanted with grafts as well as in controls having undergone sham operation.

4.3.7.4 Histology and immunohistochemistry

Grafts harvested at 12 months were fixed in 4% paraformaldehyde (PFA) and embedded in paraffin. Five-micron thick sections were then stained using standardized technique for hematoxylin and eosin (H&E). Identification of the endothelium and smooth muscle cells was done by immunohistochemical staining of the paraffin-imbedded explant sections with anti-CD31 (1:50, Abcam), alpha smooth muscle

91 actin (a-SMA) (Dako, Carpinteria, CA). Antibody binding for CD31 and aSMA was detected using biotinylated anti IgG (1:200, Vector, Burlingame, CA). This was followed by binding of streptavidin-horse radish peroxidase (HRP) and color development with

3,3-diaminobenzidine (DAB).

4.3.8 Statistical analysis

Results are expressed as mean ± standard deviation. The statistical significance of differences among time points was analyzed using one-way ANOVA. A probability value of less than 0.05 was considered significant.

4.4 Results and discussion

The polymer synthesis, scaffold characterization and long term performance evaluation of the engineered grafts in an abdominal aortic mouse model are discussed in the following sections.

4.4.1 Polymer synthesis and characterization

The PEU polymer was synthesized from the amino acid L-leucine, 1,

10-decanediol, and triphosgene using an interfacial polymerization as described in

Scheme 4.1.72,225 The chemical structures of the monomer and polymer were confirmed by 1H NMR and 13C NMR spectroscopy (Figure 3.5 and 3.6). The molecular mass, molecular mass distribution and thermal properties of the polymer were studied by SEC,

DSC (Figure 3.7) and TGA (Figure 3.8). The data summery is listed in Table 4.1. In this study, high molecular mass PEUs (Mw > 100 kDa) with good fiber forming property were

92 synthesized using interfacial polymerization. This is of great significance since sufficiently high molecular mass capable of fiber or film forming is required for practical applications of PEU polymers as biomaterial scaffolds. Moreover, the degradation temperature (Td, 273 °C) of the Poly(1-LEU-10) material is much higher than its glass transition temperature (Tg, 30 °C), indicating that this material can be melt processed with limited impact of thermal degradation. These characteristics allow high temperature processing techniques such as molding and melt processing to be used to process the PEU polymers in addition to electrospinning.

Scheme 4.1. The two-step synthetic route of L-leucine-based poly(ester urea)s

(Poly(1-LEU-10)).

Table 4.1. Characterization data summary for the amino acid-based poly(ester urea).

Samples Mw Td (°C) Tg (°C) Poly(1-LEU-10) 135,000 1.9 273 30

4.4.2 Scaffold characterization

Electrospinning conditions (solution concentration, flow rate, voltage levels, and distance between the needle tip and the mandrel, etc.) were optimized in order to obtain uniform bead-free nanofibrous morphology before collecting on rotating mandrel for

93 graft fabrication. Graft scaffolds obtained were cut into 1 mm thick cross-sections and imaged on a field emission SEM (FE-SEM; JSM-7401F, JEOL Ltd., Japan). Fiber diameters at the outer surface and wall thickness were measured from high and low magnification SEM images. Figure 4.1(a) shows a gross appearance of the whole graft tube. The grafts fabricated were generally about 3 cm in length and approximately 1 mm in inner diameter. Figure 4.1(b) shows tilted view of the graft tube at low magnification

SEM. Under low magnification we can see the graft has nice round tubular structure.

When zooming into higher magnification at the surface we can see the nice fiber structure that appears consistent (Figure 4.1(c)), and at the edge we can see the cross sections of the fibers interconnecting across the thickness of the graft wall (Figure 4.1(d)). The averaged fiber diameter and pore area counted by ImageJ were 422 ± 33 nm and 10 ± 4

μm2, which are good for cell adhesion and blood vessel regeneration. Additionally, the wall thickness of the graft tubes, as determined from SEM images, was found proportionally increasing with the electrospinning collecting time, indicating that the wall thickness can be finely tuned by adjusting the electrospinning collecting time. The wall thicknesses of the tubes fabricated at different electrospinning collecting times were as follows: 151 ± 27 μm (t = 60 mins), 204 ± 18 μm (t = 90 mins), 255 ± 18 μm (t = 120 mins), 305 ± 15 μm (t = 150 mins) and 348 ± 17 μm (t = 180 mins).

94 Figure 4.1. (a) The gross appearance and SEM images of small diameter electrospun PEU grafts, (b) entire (×40), (c) surface (×2500), and (d) cross-sectional (×2500) morphologies. Based on the SEM image analysis, the average fiber diameter and pore size are 420 ± 20 nm and 10 ± 4 μm2, respectively. These findings demonstrate the feasibility of using electrospinning to fabricate porous small diameter vascular grafts, which are good for cell adhesion and blood vessel regeneration.

4.4.3 Biomechanical properties of scaffolds

The tensile properties, suture retention strength, and burst pressure were measured on all PEU electrospun tubes to ensure that they possess significant biomechanical properties to function as vascular grafts. Physical properties of the Poly(1-LEU-10) electrospun grafts was summerized in Table 4.2.

The tensile properties of the electrospun tube scaffolds were studied by uniaxial tensile testing of the whole graft using an Instron 5567 universal tensile testing machine.

Based on stress-strain curve (Figure 4.2), the ultimate tensile strength (UTS), elongation 95 at break, and elastic modulus of the grafts were obtained. The graft scaffolds showed averaged elastic modulus of 1.8 ± 0.1 MPa, ultimate tensile strength of 1.7 ± 0.2 MPa and elongation at break of 598 ± 26 %. Here, we noticed that the elastic modulus of the PEU grafts fell within the range of the native blood vessels (0.16-12 MPa).255-257 This is of great significance, since the close the mechanical properties of grafts come to that of native blood vessels, the less the chance of graft failure due to mechanical property mismatch.251 In addition, this matching of mechanical properties may aid in reducing compliance mismatch as well.122

Suture retention strength is essential to evaluate the material for resisting the tension during implantation and it directly relates to the success of the graft implantation procedure. Results analysis as determined by the ultimate tensile strength test (Figure 4.3) demonstrated the electrospun PEU grafts show sufficiently high enough suture retention strength (8.7 ± 0.4 N). Compared to suture strength of native artery (nonviable porcine femoral artery, nvPFA, 2.31-3.51 N ) and commonly used vascular graft material (ePTFE,

4.91-6.67 N ) as referred from previous published papers,122 the electrospun PEU grafts showed more than adequate strength for suturing during implantation. Also, there are other previous works reported it is generally accepted to be greater than 2.0 N.298

Burst pressure identified as the maximum pressure that the scaffolds could endure before failure is a crucial factor to determine whether the scaffold material is strong enough to endure physiologic forces and avoid blood leakage. In our case, the limitation of the burst pressure testing machine is 1000 mmHg. As the pressure inside the grafts with continuous water flow increased gradually until it reached the limitation of the machine, the PEU electrospun grafts did not break even after we held the pressure at

96 1000 mmHg for 30 mins and no leakage was observed either. This result demonstrates that the PEU grafts possess excellent physical strength and can be developed as substitutes for native blood vessels since the blood pressure is generally less than 200 mm

Hg for human beings.

Table 4.2. Physical properties summery of Poly(1-LEU-10) electrospun grafts.

Image analysis Tensile testing (n=6) Suture Burst PEU Ultimate retention pressure Wall Fiber Pore Elastic Elongation grafts tensile strength (N) (mm thickness diameter area Modulus at break strength (n=6) Hg) (μm) (nm) (μm2) (MPa) (%) (MPa) Type 254 ± 20 422 ± 33 10 ± 4 8.7 ± 0.4 > 1000 A 1.7 ± 0.2 1.8 ± 0.1 598 ± 26 Type 350 ± 18 417 ± 28 10 ± 4 12.0 ± 1.3 > 1000 B

Figure 4.2. Stress-strain curve of PEU whole graft in wet condition from uniaxial tensile testing. The tensile properties of the PEU grafts were measured using an Instron 3365 universal materials testing machine. The gauge length was 10 mm and the crosshead 97 speed was set at 10 mmmin-1. The elastic modulus of the grafts, ultimate tensile stress

(UTS) and elongation at break (%) were obtained from the stress-strain curve. Results presented are average values for six individual measurements (n=6).

Figure 4.3. Suture retention strength of PEU whole grafts test with commercial 5-0

Prolene sutures.

4.4.4 In vitro cell study results

There is little evidence regarding the ability of PEUs to support vascular cell attachment and proliferation, which is a first requirement for their application in vascular tissue engineering. In order to evaluate to what extent the current PEU materials support the attachment and spreading of vascular cells, A-10 SMCs and HUVECs were seeded on positive control glass coverslips and electrospun PEU covered glass coverslips, both of which are two-dimensional (2D) surfaces. After being cultured for up to 48 h, labeling for

F-actin and DNA were used to examine cellular morphology. As shown in Figure 4.4(a), 98 both A-10 SMCs and HUVECs were well attached and spread on the 2D surface with abundant and aligned F-actin expression. Quantification of cell spread area revealed that cells spread comparably with those cultured on control glass coverslips (Figure 4.4(b)), which means cells put down attachments and stretched out on the PEU electrospun nanofibers as they normally would on glass coverslips. This result obviously suggests that the PEU nanofibers are able to support vascular cell adhesion and spreading in vitro.

Since cell adhesion and spreading are the first events that dictate the subsequent cellular responses such as proliferation, migration and matrix deposition,254 it is important that the

PEU nanofibers were able to promote these initial events.

99

Figure 4.4. A-10 smooth muscle cells (A-10 SMCs) and human umbilical vein cells

(HUVECs) attachment and spreading on PEU electrospun nanofibers (cell seed density:

78 mm-2; red: F-actin stained by rhodamine phalloidin; blue: nucleus stained by DAPI).

(a) A flat and spread morphology is clear observed for both cell type up to 48 h culture time, indicating the PEU nanofibers are able to support adhesion of A-10 SMCs and

HUVECs in vitro. (b) Quantification of cell spread area revealed that cells spread comparably with those cultured on control glass coverslips, which means cells put down attachments and stretched out on the PEU electrospun nanofibers as they normally would on glass coverslips. 100 The growth of vascular cells on the nanofibrous scaffolds is another critical issue for their clinical applications. The proliferation of A-10 SMCs and HUVECs on electrospun PEU nanofiber in vitro can provide initial confirmation of the utility of the scaffolds. The growth profiles of A-10 SMCs and HUVECs cultured on the positive control glass coverslips and electrospun PEU covered glass coverslips were measured over a seven-day time course. As shown in Figure 4.5, the vascular cells continued to increase in number over the time interval examined on both positive controls and electrospun PEU at similar proliferation rates, indicating that the PEU nanofibers are able to support vascular cell proliferation without producing toxic effects for at least 7 days in vitro.

Figure 4.5. Cell proliferation of A-10 smooth muscle cells (A-10 SMCs) and human umbilical vein cells (HUVECs) cultured in direct contact with electrospun PEU nanofibers after 1, 3 and 7 days of cell seeding, as determined by PrestoBlue assay. Blank 101 glass coverslips are used as positive controls. All experimental groups with PEU electrospun nanofibers showed similar proliferation rates of the A-10 SMCs and

HUVECs to positive control at day 1, 3 and 7, indicating the PEU nanofibers are able to support proliferation of vascular cells in vitro.

4.4.5 Survival of the animals

The survival rate of mice with Poly(1-LEU-10) graft implantation at 24 hours after surgery was 100%. The survival rate at 12 months post-operation was 80% (4/5).

One mouse died at 3 months after graft implantation due to a thymus tumor (determined by necropsy) and was not graft related. No long term graft related complications such as graft rupture or aneurysmal dilatation were observed.

4.4.6 Ultrasound and micro-CT

In vivo ultrasound was performed at 5 weeks, 9 weeks and 12 months. All

Poly(1-LEU-10) grafts demonstrated luminal patency at each time point without evidence of aneurysmal dilatation or stenosis (Figure 4.6(b) and 4.6(c)). The inner diameter of the graft lumen decreased from the 5 week time point to the 12 month time point post-implantation as a consequence of neovessel regeneration. Micro-CT angiography showed that aneurysmal dilatation and stenosis were absent in Poly(1-LEU-10) type A grafts 12 months after graft implantation (Figure 4.7(a) and 4.7(b)), while all type B grafts experienced occlusion (Figure 4.7(c) and 4.7(d)).

102 Figure 4.6. (a) Intraoperative photograph demonstrating Poly(1-LEU-10) vascular grafts during surgical implantation. (b) Serial doppler ultrasound examinations were performed on all implanted grafts. All grafts remained patent to the experimental end point according to the ultrasound tests. (c) Graft inner diameter change was calculated by

Image J software.

103

Figure 4.7. In vivo micro computed tomography (CT) angiography was performed at 12 months (a, b, c, d). All type A grafts (250 μm) showed long term patency at the time point

12 months while all type B grafts (350 μm) demonstrated occlusion.

4.4.7 Histological assessment

Histological assessment was further performed on all harvested Poly(1-LEU-10) grafts after 12 months (2 of type A grafts and 2 of type B grafts) to evaluate the neovessel remodeling process. Endothelial cells are the predominant cells in the lumen of the blood vessel walls and are significant for the structural and functional integrity of the new blood vessel. In this study, endothelial cells, which were defined by immunohistochemical staining, showed presence of CD31 (a marker for endothelial cells), lined in the lumen of type A grafts after graft implantation for 12 months. This result indicated confluent endothelium layers were formed in the type A graft lumen that can mimic native aorta

(Figure 4.8(b) and 4.8(e)). Smooth muscle cells were defined by immunohistochemical smooth muscle actin (aSMA) staining and were observed in the tunica media which infiltrated and replaced the Poly(1-LEU-10) type A grafts (Figure 4.8(c) and 4.8(f)).

When examining the immunohistochemical staining results of type B grafts, 104 well-organized endothelium and smooth muscle layers as observed in type A grafts were absent. The significantly different new tissue remodeling could explain the long term patency of type A grafts and occlusion of type B grafts after 12 months implantation.

However, for vascular grafts that only varied in wall thickness while maintaining the same material composition, elastic mechanical properties, and structure morphology, it is interesting to observe such vastly different results in the animal model study. Future work will include further exploration into the wall thickness effects by testing compliance properties of the two different graft types.

Figure 4.8. H&E image of harvested Poly(1-LEU-10) grafts at 12 months after implantation (a, d, h, k); Endothelial layer of graft lumen stained by CD31 markers (b, e, i, l); Smooth muscle cells stained by immunohistochemical smooth muscle actin (aSMA)

(c, f, j, m).

105 4.5 Conclusion

This manuscript described a cell free small diameter (ID≈1 mm) vascular graft engineered from electrospun resorbable poly(ester urea) (PEU). Long term performance of two types of PEU grafts of different wall thickness (type A: 250 μm; type B: 350 μm) were evaluated in an abdominal infra-renal aortic interposition mouse model over 1 year.

Extensive tissue remodeling was observed in all type A grafts, which lead to the development of neovessels that mimics native arteries. However, the type B grafts experienced occlusion over the 1 year interval due to intimal hyperplasia. These results suggest the poly(ester urea) may be a promising material for cell-free tissue engineered small diameter vascular grafts, but further investigations will be required to assess the effect of compliance on the long term performance of the grafts.

4.6 Acknowledgments

The authors are grateful for financial support from the Akron Functional Materials

Center and the Austen Bioinnovation Institute in Akron. Drs. Fei Lin, Erin P. Childers,

Mary Beth Wade are acknowledged for useful discussions and comments on the manuscript. Additional thanks to Dr. Yong Ung Lee from the Nationwide Children’s

Hospital for the grafts compliance testing and Prof. Hossein Tavana in Biomedical

Engineering at the University of Akron for the source of HUVECs.

106 CHAPTER V

NEW SUSTAINED RELEASE STRATEGY OF RECOMBINANT HUMAN

GROWTH HORMONE (rhGH) FROM BIORESORBABLE POLY(ESTER UREA)

NANOFIBERS

This work is not published yet

Yaohua Gao, Adam Land, Joshua Bundy, Gina Policastro, Todd Ritzman, Matthew L.

Becker, in preparation, 2016

5.1 Outline

Recombinant human growth hormone (rhGH) for protein therapeutics is in great demand. However, rhGH has a short half-life and the product requires frequent subcutaneous injection leading to poor patient compliance. Currently, it is challenging to develop effective rhGH sustained delivery systems that can exceed a month in duration.

In this study, a new sustained release strategy of rhGH was developed by encapsulating sugar glass-stabilized rhGH in electrospun, bioresorbable poly(ester urea) (PEU) nanofibers. The protein was found to be randomly dispersed throughout the electrospun fibers in an aggregate form, and sustained rhGH release with modest burst release was observed for at least 6 weeks as confirmed by BCA protein assay. These results clearly

107 suggest the feasibility of this system as a long-term sustained release strategy for rhGH

and its potential use as an implantable delivery scaffold for therapeutics.

5.2 Introduction

Protein therapeutics possess advantages over small-molecule techniques such as

high target specificity, low interference with normal biological processes.299-302 Human

growth hormone (hGH) has an unique role in promoting longitudinal bone growth and is

widely used for the clinical treatment of pediatric short stature caused by growth

hormone deficiency, growth failure from Turner syndrome, or chronic renal failure.303-311

However, as a protein drug, the current rhGH product requires frequent subcutaneous

injection (i.e. three times a week or daily) as a consequence of its short half-life, which

leads to poor patient compliance, high dose, and increased cost.312-315 Therefore, a

sustained-release rhGH formulation would not only provide improved patient compliance,

but also alleviate the costly burden associated with frequent injections. The development

of such a system has been attempted extensively using various strategies, including

fusion of stabilizing peptides,316-318 crystal formulation,319 encapsulation in

microspheres320-326 or hydrogels327-331, and PEGylation332-335. Among them, the most

investigated method is encapsulation of rhGH into biodegradable poly (lactic

acid-co-glycolic acid) (PLGA) microspheres. The best known rhGH sustained delivery

system, Nutropin Depot®, which is composed of PLGA microsphere, was approved by

the US FDA in 1999 as a monthly product.319,325,326 However, this product was

withdrawn from the market in 2004 as a result of several drawbacks, including low

108 loading efficiency, high burst release, protein denaturation during microsphere preparation, and inflammation from acidic PLGA degradation byproducts.325,326 So far, it is still quite a challenge to develop effective rhGH delivery system that lasts longer than a month.

Electrospun nanofibers have attracted increasing interest in controlled delivery of bioactive molecules, such as proteins209-211,214, growth factors111,212,214,215, and genes113,336,337. As controlled delivery carriers, electrospun nanofibers offer many advantages including:201,202 1) high drug loading efficiency, 2) enhanced drug diffusion to surrounding medium, and 3) improved solubility of various bioactive molecules.

Furthermore, sustained release can be achieved by properly modulating the biodegradability, hydrophilicity or hydrophobicity properties of polymers, as well as the sizes and porosities of the nanofibers.204 In addition, to decrease the amount of bioactive molecules administrated and the non-target delivery toxicity, local delivery can be achieved by using the electrospun nanofibers as an implantable device. Despite these advantages, problems associated with electropun nanofibers are the loss of biomolecule bioactivity during electrospinning preparation and execution, and the adverse inflammation reactions to surrounding tissues caused by acidic degradation of the polymer nanofibers. Therefore, stabilizing the bioactive molecules prior to electrospinning and using a polymer that does not produce acidic degradation byproducts, are necessary to make effective sustained delivery system for protein drugs like rhGH.

Recently, a novel sugar glass nanoparticle system for stabilizing proteins in drug delivery systems was reported by Giri et al.114 This sugar-glass technology yields excellent protein protection from process-related stresses (e.g. organic solvent exposure)

109 with little or no denaturation, very good encapsulation efficiency, and burst-free sustained release for essentially any protein and polymer system of interest. Additionally, we have also explored a series of amino acid based poly(ester urea)s (PEUs), that showed tunable degradation with non-acidic byproducts.72,223-227 We therefore hypothesized that the sugar glass nanoparticle stabilized rhGH encapsulated in the PEU electrospun nanofibers could provide a new sustained release strategy for rhGH. PCL is one of the mostly studied degradable polymers that have been used for several human clinical applications.128 In this work, rhGH delivery system based on PCL electrospun nanofibers was also prepared and studied as a comparison.

5.3 Experimental section

PEU polymer synthesis, sugar glass nanoparticle stabilized rhGH fabrication, blend electrospinning of sugar nanoparticle stabilized rhGH with PEU polymer, and in vitro protein release study are highlighted in the following sections.

5.3.1 Materials

1,8-Octanediol (98%, Sigma-Aldrich), p-Toluenesulfonic acid (99%, Fisher

Scientific), Triphosgene (98%, Alfa Aesar), Sodium carbonate (NaCO3, 99%, Fisher

Scientific), Dioctyl sulfosuccinate (AOT, 96%, Alfa Aesar), D-(+)-trehalose dehydrate

(trehalose, 99%, Sigma-Aldrich), PCL (poly (ɛ-caprolactone), 150 kDa, Scientific polymer products, Inc., Ontario, NY) were used as received. All solvents of HPLC grade, e.g., 2, 2, 4-trimethylpentane (isooctane), 1,1,1,3,3,3-Hexafluoro -2-propanol (HFIP), and analytical grade such as dichloromethane (DCM), dimethylformamide (DMF),

110 1,4-Dioxane and ethanol (EtOH) were purchased from Sigma-Aldrich (St Louis, MO) and used without further purification. Recombinant human growth hormone (rhGH) was kindly donated by Dr. Todd Ritzman from the Akron hildren’s Hospital (Akron, OH).

BCA assays (bicinchoninic acid) were obtained from Pierce Biotechnology (Rockford,

IL). Rhodamine B was purchased from Sigma- Aldrich (St Louis, MO).

5.3.2 Polymer Synthesis and Characterization

The poly(ester urea) monomer and polymer were prepared following previous

223 1 methods. Poly(1-LEU-8): H NMR (500 MHz, DMSO-d6): 0.83-0.87 (m, 12H), 1.25

(m, 4H), 1.41-1.44 (m, 4H), 1.52-1.54 (t, 4H), 1.58-1.62 (m, 2H), 2.50 (DMSO), 3.36

13 (H2O), 3.96-4.02 (m, 4H), 4.11-4.12 (m, 2H), 6.27-6.29 (d, 2H). C NMR (500 MHz,

DMSO-d6): 22.11, 23.06, 24.72, 25.64, 28.51, 28.94, 39.52-40.52 (DMSO-d6), 41.47,

51.50, 64.56, 157.50, 173.83. FT-IR (cm-1): 1734 [-C(CO)-O-], 1635, 1548

[-NH-C(O)-NH-], 3355 [-NH-C(O)-NH-]; Mn = 77 kDa, Mw = 131 kDa. Td = 272 °C , Tg

= 37 °C .

5.3.3 SGnPs preparation and characterization

SGnPs were prepared from inverse micelles of AOT in isooctane according to a published procedure with minor modification.114 Generally, AOT (2.13 g, 0.0048 mol) was dissolved in 12 mL of isooctane in a 25 mL centrifuged tube to produce a 0.4 M solution. An aqueous solution containing protein with trehalose was then added dropwise while vortexing. The water:surfactant mole ratio, w = [H2O]/[AOT], was set to 15. The mass ratio of protein to trehalose was maintained at 1:200 (0.5 mg protein in 100 mg of

111 trehalose). After addition of the aqueous solution, the mixture was continuously vortexed for 2 minutes until a clear suspension was observed. The resulting inverse micelles were obtained by flash-freezing the clear suspension with liquid nitrogen. The frozen micelles were then lyophilized under vacuum for 3 days, followed by washing 5 times with isooctane. The resulting nanoparticles were re-suspended in isooctane and stored in a desiccator at -20 °C until use. Rhodamine B loaded SGnPs were prepared using a similar method. Here, sugar glass nanoparticles encapsulated with rhGH are named as rhGH-SGnP, and sugar nanoparticles encapsulated with fluorescent rhodamine B dyes are abbreviated as RB-SGnP. The morphology and size distribution of the nanoparticles were characterized by transmission electron microscopy (Philips TECNAI TEM) and dynamic light scattering (Brookhaven light scattering spectrometer, BI-200SM).

5.3.4 Electrospun fiber mat fabrication and characterization

Plain PEU and PCL nanofibers: plain PEU and PCL nanofibers without protein or fluorescent dye loading were prepared by electrospinning on a 10 wt.% Poly(1-LEU-8) polymer solution in HFIP and a 12 wt.% PCL polymer solution in DCM:DMF = 4:1 (v/v) separately.

PEU nanofibers loaded with rhGH-SGnPs or RB-SGnPs: To fabricate growth hormone or fluorescent dye encapsulated PEU fibers, Poly(1-LEU-8) was initially dissolved in 1,4-dioxane at a concentration of 12 wt.%. Growth hormone or fluorescent dye was added into the PEU polymer solution in the form of sugar glass nanoparticles.

The rhGH-SGnPs or RB-SGnPs suspension in isooctane was centrifuged at 300 g for 2 minutes. After discarding the supernatant isooctane, a mixture of 1,4-dioxane and EtOH

112 was added to the precipitate. The reconstituted rhGH-SGnP or RB-SBnPs suspension was then transferred to the prepared PEU polymer solution. The resulting SGnPs-polymer suspension was vortexed to distribute the protein or dye nanoparticles uniformly throughout the polymer solution. The final polymer concentration was 7 wt.% in

1,4-dioxane:EtOH = 2:1 (v/v) for electrospinning and the loading level of growth hormone or fluorescent dye nanoparticles in the nanofibers was 1.6 wt.%.

PCL loaded with rhGH-SGnPs or RB-SGnPs: Growth hormone or fluorescent dye loaded PCL nanofibers were prepared similarly to PEU nanofibers, except a mixture solvent of DCM:EtOH = 3:1 (v/v) was used for electrospinning. The loading level of growth hormone and dye nanoparticles in the PCL nanofibers was controlled the same as the PEU nanofibers (i.e., 1.6 wt.%).

The electrospinning set-up included a syringe pump, a high voltage supply, and a piece of aluminum foil. A positive high voltage (10 kV) was applied to the polymer solution by the power supply, and the mixture solution was delivered through a 21 gauge blunt tip syringe needle at a constant flow rate of 3 mL/h to produce fine polymer nanofibers. The fibers were collected on a piece of grounded aluminum foil. The collecting distance between the syringe needle tip and the aluminum foil was 15 cm and the fiber collection was continued until all polymer solution was consumed. The obtained polymer nanofibers were carefully peeled off from the aluminum foil and further dried for at least 12 hours under vacuum to remove trace residue of solvent. The vacuum dried fiber mats were stored in a desiccator for future use. The fiber diameter, morphology and distribution of protein/dye nanoparticles within the fibers were studied by scanning

113 electron microscopy (JEOLJSM-7401F SEM) and fluorescent microscopy (OLYMPUS

IX 81).

5.3.5 In vitro protein release study

Approximately 90 mg of electrospun nanofibers encapsulated with protein

(rhGH-SGnPs) was soaked in 10 mL of PBS in a 20 mL glass vial. The glass vial was incubated at 37 °C in the presence of 5% carbon dioxide with mild stirring. At various time points, 0.15 mL of supernatant was retrieved from the vial followed by an equal addition of fresh medium. The released protein was quantified using a Micro BCA

(bicinchoninic acid) Protein Assay Kit (Pierce, USA) according to the manufacturer protocol. Absorbance at 562 nm was measured on a SynergyTM MX plate reader from

BioTek. The protein release was calculated using an established standard curve of rhGH ranging from 0 to 250 μg mL. All release experiments were conducted in triplicate.

5.4 Results and discussion

The sugar glass nanoparticle characterizations, protein distribution within the electrospun nanofibers and sustained protein release from the nanofibers for up to 6 weeks in vitro are discussed in the following sections.

5.4.1 Sugar glass protein nanoparticle

Sugar glass protein nanoparticles were prepared from an inverse micelle of AOT according to a previous published method,114 as shown in Figure 5.1(a). According to the literature, the [water]/[surfactant] mole ratio, w = [H2O]/[AOT], plays an important role

114 in determining the equilibrium micelle size. The [water]/[surfactant] mole ratio can be varied from 10 to 15 and the mass ratio of protein to sugar (i.e. trehalose) can be tuned from 1:500 to 1:200 to provide sufficient coating of sugar-glass around the protein without adversely effecting performance. In this study, to maximize rhGH encapsulation while maintaining the stabilization effects, the [water]/[surfactant] mole ratio and the mass ratio of protein to sugar used were 15 and 1:200, respectively. The choice of w and mass ratio of protein to sugar resulted in typical particle sizes distributed from 64 ± 13 nm, as measured from TEM (Figure 5.1(c)) and 81 ± 20 nm as determined by DLS

(Figure 5.1(d)). The size of the protein nanoparticles determined by DLS is somewhat larger than that measured by TEM, which is because the surfactant chains surrounding the nanoparticles are extended in the solution state. The freeze-dried nanoparticles can be reintroduced into organic solvents and polymer solutions as a stable suspension. Figure

5.1(b) showed that the sugar glass stabilized rhGH nanoparticles can be homogeneously suspended in isooctane and a 12 wt.% PCL polymer solution in DCM.

115

Figure 5.1. (a) Schematic presentation of a biomolecule encapsulated in a SGnP. (b) rhGH-SGnPs suspended in isooctane. (c) Representative TEM image of rhGH-SGnP. (d)

Particle size distribution of rhGH-SGnP from DLS.

5.4.2 Electrospun fiber mat fabrication and characterization

The electrospinning parameters were optimized by conducting a series of systematic studies on the effects of flow rate, polymer/protein concentration, applied voltage and jet stability on the size and morphology of the resulting polymer nanofibers.

To fabricate plain nanofibers without protein encapsulation, stable polymer jets yielding desired fiber morphologies can be easily obtained by electrospinning a 10 wt.% PEU in

HFIP or a 12 wt% PCL in DCM:DMF (4:1 v/v). However, to fabricate protein/dye nanoparticles encapsulated corresponding polymer nanofibers, the optimized parameter

116 was found to be 7 wt.% PEU polymer solution plus 1.6 wt.% protein/dye nanoparticles in

1,4-dioxane:EtOH = 2:1 (v/v) and 7 wt.% PCL polymer solution plus 1.6 wt.% protein/dye nanoparticles in DCM:EtOH = 3:1 (v/v). Dioxane and EtOH was used in the mixture solvent instead of HFIP or DMF for the protein loaded fiber electrospinning is because the sugar glass nanoparticles are effective at acting as a barrier between the protein and organic solvent if the solvent polarity index is less than 5.114 Dioxane, DCM and EtOH have a polarity index of 4.8, 3.4 and 4.3, respectively. The solvent mixture of

Dioxane:EtOH = 2:1 (v/v) and DCM:EtOH =3:1 (v/v) will have a polarity index less than

5, which is favorable for protein bioactivity reservation during the electrospinning process. The fiber size, morphology, and sugar glass protein nanoparticle distribution within the fibers were analyzed using SEM. Generally, plain nanofibers without protein encapsulation, showed randomly oriented fibers with uniform size distribution in the range of 422 ± 33 nm for PEU and 581 ± 30 nm for PCL with smooth surface morphology (Figure 5.2(a) and 5.2(d)). However, the protein-encapsulated fibers showed much rougher surface morphology and less uniform size as a consequence of protein nanoparticle aggregation within the fibers (Figure 5.2(b), PEU, 542 ± 128 nm; Figure

5.2(e), PCL, 360 ± 118 nm). Besides, when incorporating the sugar stabilized protein nanoparticles into the polymer system, the electrospinning fiber jet became less stable, and this may cause the less uniform fiber diameters too.

To study the protein nanoparticle distribution within the polymer fibers, fluorescent dye sugar glass nanoparticles (RB-SGnPs) were prepared and introduced into the corresponding polymer nanofibers. As confirmed by the fluorescent microscopy

(Figure 5.2(c) and 5.2(f)), the fluorescent RB-SGnPs were found to be randomly

117 dispersed throughout the nanofibers in an aggregate form. Furthermore, if we compare the two different polymer systems, we found that the aggregation of protein nanoparticles was much less within the PEU nanofibers than within PCL for unknown reason. The aggregation of protein nanoparticles in the polymer nanofibers may increase the chance of burst release when separation occurs between the polymer and protein-bearing phases.

Figure 5.2. SEM image of polymer nanofibers (a) plain PEU, (b) rhGH-SGnP loaded

PEU, (c) RB-SGnP loaded PEU, (d) plain PCL, (e) rhGH-SGnP loaded PCL, (f)

RB-SGnP loaded PCL.

5.4.3 In vitro release study

The in vitro protein release from both rhGH-SGnP loaded PEU and PCL nanofibers was quantified using a BCA (bicinchoninic acid) protein assay. The cumulative rhGH release profile is shown in Figure 5.3. Sustained release of rhGH from

118 both PEU and PCL nanofiber delivery systems was observed for up to 6 weeks with a modest burst release within the first 24 hours (i.e., 7-10 % for PEU and 13-15 % for PCL).

The long-term relatively slow and steady protein release is expected as a consequence of the slow degradation rates of PEU and PCL.204,223,338,339 The initial burst release is generally attributed to the nanoparticles located on the surface of the fibers. These nanoparticles have some direct exposure to the medium, as opposed to being shielded by a significant amount of polymer, so they are expected to be released from the fiber matrix rapidly. This behavior is illustrated in the inset of Figure 5.3. Additionally, aggregation of the protein nanoparticles within the polymer fibers contributes to the initial burst release as well. We observed that the rhGH-SGnPs loaded PEU nanofibers which had less protein nanoparticle aggregation demonstrated lower burst release. Uncontrolled initial burst release is usually considered detrimental to sustained protein release. It is good to observe a reduced burst release in our PEU delivery system compared to PCL. In addition to the reduced burst release, another advantage of the PEU nanofiber delivery system is that the degradation of PEU does not cause local acidosis or adverse inflammatory reactions to surrounding tissues.72,226 Therefore, the electrospun PEU delivery system also has a great potential to be used as an implantable bioactive scaffold for local rhGH delivery.

119 Figure 5.3. Sustained release profile of rhGH-SGnPs from PEU and PCL electrospun nanofibers within 6 weeks in vitro (insert: schematic presentation of rhGH-SGnPs distribution across the fiber diameter).

5.5 Conclusion

Sugar glass nanoparticle stablized rhGH was successfully incorporated into resorbable poly(ester urea) (PEU) nanofibers through electrospinning. The protein was found to be randomly dispersed throughout the polymer fibers with some aggregations. A sustained release of rhGH from the electrospun PEU nanofiber mat for up to 6 weeks was observed. Work is ongoing in evaluating the bioactivity of rhGH after encapsulation and release.

120 5.6 Acknowledgements

The authors thank Prof. Marcus Cicerone for help and discussion with sugar protein nanoparticle preparation, Dr. Jinjun Zhou for TEM analysis and Dr. Hui Li for

DLS testing. This work was supported by the Akron Functional Materials Center and the

Austen Bioinnovation Institute in Akron.

121 REFERENCES

(1) Agarwal, S.; Wendorff, J. H.; Greiner, A. Polymer 2008, 49, 5603.

(2) Bhardwaj, N.; Kundu, S. C. Biotechnology Advances 2010, 28, 325.

(3) Pham, Q. P.; Sharma, U.; Mikos, A. G. Tissue engineering 2006, 12, 1197.

(4) Darrell, H. R.; Iksoo, C. Nanotechnology 1996, 7, 216.

(5) Ke, P.; Jiao, X.-N.; Ge, X.-H.; Xiao, W.-M.; Yu, B. RSC Advances 2014, 4, 39704.

(6) Reneker, D. H.; Yarin, A. L.; Fong, H.; Koombhongse, S. J. Appl. Phys. 2000, 87, 4531.

(7) Li, D.; Xia, Y. Advanced Materials 2004, 16, 1151.

(8) Xu, C. Y.; Inai, R.; Kotaki, M.; Ramakrishna, S. Biomaterials 2004, 25, 877.

(9) Frenot, A.; Chronakis, I. S. Curr. Opin. Colloid Interface Sci. 2003, 8, 64.

(10) Deitzel, J. M.; Kleinmeyer, J.; Harris, D.; Beck Tan, N. C. Polymer 2001, 42, 261.

(11) Gupta, P.; Elkins, C.; Long, T. E.; Wilkes, G. L. Polymer 2005, 46, 4799.

(12) Tan, S. H.; Inai, R.; Kotaki, M.; Ramakrishna, S. Polymer 2005, 46, 6128.

(13) Koski, A.; Yim, K.; Shivkumar, S. Mater. Lett. 2004, 58, 493.

(14) Zong, X.; Kim, K.; Fang, D.; Ran, S.; Hsiao, B. S.; Chu, B. Polymer 2002, 43, 4403.

(15) Fong, H.; Chun, I.; Reneker, D. H. Polymer 1999, 40, 4585.

(16) Zhang, C.; Yuan, X.; Wu, L.; Han, Y.; Sheng, J. Eur. Polym. J. 2005, 41, 423.

(17) Zuo, W.; Zhu, M.; Yang, W.; Yu, H.; Chen, Y.; Zhang, Y. Polymer Engineering & Science 2005, 45, 704.

122 (18) Jiang, H.; Fang, D.; Hsiao, B. S.; Chu, B.; Chen, W. Biomacromolecules 2004, 5, 326.

(19) Kim, B.; Park, H.; Lee, S.-H.; Sigmund, W. M. Mater. Lett. 2005, 59, 829.

(20) Hayati, I.; Bailey, A. I.; Tadros, T. F. J. Colloid Interface Sci. 1987, 117, 205.

(21) Mit-uppatham, C.; Nithitanakul, M.; Supaphol, P. Macromol. Chem. Phys. 2004, 205, 2327.

(22) Demir, M. M.; Yilgor, I.; Yilgor, E.; Erman, B. Polymer 2002, 43, 3303.

(23) Baker, S. C.; Atkin, N.; Gunning, P. A.; Granville, N.; Wilson, K.; Wilson, D.; Southgate, J. Biomaterials 2006, 27, 3136.

(24) Catalani, L. H.; Collins, G.; Jaffe, M. Macromolecules 2007, 40, 1693.

(25) Fennessey, S. F.; Farris, R. J. Polymer 2004, 45, 4217.

(26) Buchko, C. J.; Chen, L. C.; Shen, Y.; Martin, D. C. Polymer 1999, 40, 7397.

(27) Beachley, V.; Wen, X. Materials science & engineering. C, Materials for biological applications 2009, 29, 663.

(28) Kidoaki, S.; Kwon, I. K.; Matsuda, T. Biomaterials 2005, 26, 37.

(29) Lyons, J.; Li, C.; Ko, F. Polymer 2004, 45, 7597.

(30) Yuan, X.; Zhang, Y.; Dong, C.; Sheng, J. Polym. Int. 2004, 53, 1704.

(31) Wannatong, L.; Sirivat, A.; Supaphol, P. Polym. Int. 2004, 53, 1851.

(32) Cui, W.; Li, X.; Zhou, S.; Weng, J. J. Appl. Polym. Sci. 2007, 103, 3105.

(33) Nottelet, B.; Pektok, E.; Mandracchia, D.; Tille, J. C.; Walpoth, B.; Gurny, R.; Möller, M. J. Biomed. Mater. Res., Part A 2009, 89A, 865.

(34) Yarin, A. L.; Koombhongse, S.; Reneker, D. H. J. Appl. Phys. 2001, 90, 4836.

(35) Mo, X. M.; Xu, C. Y.; Kotaki, M.; Ramakrishna, S. Biomaterials 2004, 25, 1883.

(36) Heikkilä, P.; Harlin, A. Eur. Polym. J. 2008, 44, 3067.

(37) Tong, H.-W.; Wang, M. Journal of Macromolecular Science, Part B 2011, 50, 1535.

(38) Geng, X.; Kwon, O.-H.; Jang, J. Biomaterials 2005, 26, 5427.

123 (39) Heydarkhan-Hagvall, S.; Schenke-Layland, K.; Dhanasopon, A. P.; Rofail, F.; Smith, H.; Wu, B. M.; Shemin, R.; Beygui, R. E.; MacLellan, W. R. Biomaterials 2008, 29, 2907.

(40) Li, W.-J.; Laurencin, C. T.; Caterson, E. J.; Tuan, R. S.; Ko, F. K. Journal of Biomedical Materials Research 2002, 60, 613.

(41) Wang, X.; Ding, B.; Li, B. Materials Today 2013, 16, 229.

(42) Heikkilä, P.; Taipale, A.; Lehtimäki, M.; Harlin, A. Polymer Engineering & Science 2008, 48, 1168.

(43) Hsiao, H.-Y.; Huang, C.-M.; Liu, Y.-Y.; Kuo, Y.-C.; Chen, H. J. Appl. Polym. Sci. 2012, 124, 4904.

(44) Podgórski, A.; Bałazy, A.; Gradoń, L. Chem. Eng. Sci. 2006, 61, 6804.

(45) Scholten, E.; Bromberg, L.; Rutledge, G. C.; Hatton, T. A. ACS Applied Materials & Interfaces 2011, 3, 3902.

(46) Shin, C.; Chase, G. G. J. Dispersion Sci. Technol. 2006, 27, 5.

(47) Uyar, T.; Havelund, R.; Nur, Y.; Balan, A.; Hacaloglu, J.; Toppare, L.; Besenbacher, F.; Kingshott, P. J. Membr. Sci. 2010, 365, 409.

(48) Zhang, Q.; Welch, J.; Park, H.; Wu, C.-Y.; Sigmund, W.; Marijnissen, J. C. M. J. Aerosol Sci. 2010, 41, 230.

(49) Jianrong, C.; Yuqing, M.; Nongyue, H.; Xiaohua, W.; Sijiao, L. Biotechnology Advances 2004, 22, 505.

(50) Norris, I. D.; Shaker, M. M.; Ko, F. K.; MacDiarmid, A. G. Synth. Met. 2000, 114, 109.

(51) Wang, X.; Drew, C.; Lee, S.-H.; Senecal, K. J.; Kumar, J.; Samuelson, L. A. Nano Lett. 2002, 2, 1273.

(52) Wang, X.; Kim, Y.-G.; Drew, C.; Ku, B.-C.; Kumar, J.; Samuelson, L. A. Nano Lett. 2004, 4, 331.

(53) Faccini, M.; Vaquero, C.; Amantia, D. Journal of Nanomaterials 2012, 2012, 9.

(54) Lee, S.; Obendorf, S. K. Text. Res. J. 2007, 77, 696.

(55) Ramakrishna, S.; Fujihara, K.; Teo, W.-E.; Yong, T.; Ma, Z.; Ramaseshan, R. Materials Today 2006, 9, 40.

124 (56) Vitchuli, N.; Shi, Q.; Nowak, J.; McCord, M.; Bourham, M.; Zhang, X. J. Appl. Polym. Sci. 2010, 116, 2181.

(57) Catto, V.; Far; #xe8; , S.; Freddi, G.; Tanzi, M. C. ISRN Vascular Medicine 2014, 2014, 27.

(58) Hasan, A.; Memic, A.; Annabi, N.; Hossain, M.; Paul, A.; Dokmeci, M. R.; Dehghani, F.; Khademhosseini, A. Acta Biomater. 2014, 10, 11.

(59)Ji, W.; Sun, Y.; Yang, F.; van den Beucken, J. J. J. P.; Fan, M.; Chen, Z.; Jansen, J. A. Pharm. Res. 2011, 28, 1259.

(60) Liu, H.; Ding, X.; Zhou, G.; Li, P.; Wei, X.; Fan, Y. Journal of Nanomaterials 2013, 2013, 11.

(61) Vasita, R.; Katti, D. S. International Journal of Nanomedicine 2006, 1, 15.

(62) Serbo, J. V.; Gerecht, S. Stem Cell Research & Therapy 2013, 4, 8.

(63) Li, S.; Sengupta, D.; Chien, S. Wiley Interdiscip. Rev. Syst. Biol. Med. 2014, 6, 61.

(64) Moroni, L.; de Wijn, J. R.; van Blitterswijk, C. A. Biomaterials 2006, 27, 974.

(65) Tamayol, A.; Akbari, M.; Annabi, N.; Paul, A.; Khademhosseini, A.; Juncker, D. Biotechnology Advances 2013, 31, 669.

(66) Stevens, M. M.; George, J. H. Science 2005, 310, 1135.

(67) Mikos, A. G.; Bao, Y.; Cima, L. G.; Ingber, D. E.; Vacanti, J. P.; Langer, R. Journal of Biomedical Materials Research 1993, 27, 183.

(68) Mikos, A. G.; Sarakinos, G.; Leite, S. M.; Vacant, J. P.; Langer, R. Biomaterials 1993, 14, 323.

(69) Lu, L.; Peter, S. J.; D. Lyman, M.; Lai, H.-L.; Leite, S. M.; Tamada, J. A.; Uyama, S.; Vacanti, J. P.; Robert, L.; Mikos, A. G. Biomaterials 2000, 21, 1837.

(70) Park, H. J.; Lee, O. J.; Lee, M. C.; Moon, B. M.; Ju, H. W.; Lee, J. m.; Kim, J.-H.; Kim, D. W.; Park, C. H. Int. J. Biol. Macromol. 2015, 78, 215.

(71) Policastro, G. M.; Lin, F.; Smith Callahan, L. A.; Esterle, A.; Graham, M.; Sloan Stakleff, K.; Becker, M. L. Biomacromolecules 2015, 16, 1358.

(72)Stakleff, K. S.; Lin, F.; Smith Callahan, L. A.; Wade, M. B.; Esterle, A.; Miller, J.; Graham, M.; Becker, M. L. Acta Biomater. 2013, 9, 5132.

125 (73) Keskar, V.; Marion, N. W.; Mao, J. J.; Gemeinhart, R. A. Tissue Engineering. Part A 2009, 15, 1695.

(74) Mikos, A. G.; Temenoff, J. S. Electronic Journal of Biotechnology 2000, 3, 114.

(75) Mooney, D. J.; Baldwin, D. F.; Suh, N. P.; Vacanti, J. P.; Langer, R. Biomaterials 1996, 17, 1417.

(76) Wachiralarpphaithoon, C.; Iwasaki, Y.; Akiyoshi, K. Biomaterials 2007, 28, 984.

(77) Curodeau, A.; Sachs, E.; Caldarise, S. Journal of Biomedical Materials Research 2000, 53, 525.

(78) Hollister, S. J. Nat. Mater. 2005, 4, 518.

(79) Seitz, H.; Rieder, W.; Irsen, S.; Leukers, B.; Tille, C. Journal of Biomedical Materials Research Part B: Applied Biomaterials 2005, 74B, 782.

(80) Wu, C.; Luo, Y.; Cuniberti, G.; Xiao, Y.; Gelinsky, M. Acta Biomater. 2011, 7, 2644.

(81) Causa, F.; Netti, P. A.; Ambrosio, L. Biomaterials 2007, 28, 5093.

(82) Holder, W. D.; Gruber, H. E.; Moore, A. L.; Culberson, C. R.; Anderson, W.; Burg, K. J. L.; Mooney, D. J. Journal of Biomedical Materials Research 1998, 41, 412.

(83) Kiechel, M. A.; Schauer, C. L. Carbohydr. Polym. 2013, 95, 123.

(84) Li, X.; Feng, Q.; Liu, X.; Dong, W.; Cui, F. Biomaterials 2006, 27, 1917.

(85) Li, X.; Huang, Y.; Zheng, L.; Liu, H.; Niu, X.; Huang, J.; Zhao, F.; Fan, Y. J. Biomed. Mater. Res., Part A 2014, 102, 1092.

(86) Li, X.; van Blitterswijk, C. A.; Feng, Q.; Cui, F.; Watari, F. Biomaterials 2008, 29, 3306.

(87) Putthanarat, S.; Eby, R. K.; Kataphinan, W.; Jones, S.; Naik, R.; Reneker, D. H.; Farmer, B. L. Polymer 2006, 47, 5630.

(88) Rnjak-Kovacina, J.; Wise, S. G.; Li, Z.; Maitz, P. K. M.; Young, C. J.; Wang, Y.; Weiss, A. S. Acta Biomater. 2012, 8, 3714.

(89) Yang, L.; Fitié, C. F. C.; van der Werf, K. O.; Bennink, M. L.; Dijkstra, P. J.; Feijen, J. Biomaterials 2008, 29, 955.

(90) Zhang, Y.; Ouyang, H.; Lim, C. T.; Ramakrishna, S.; Huang, Z.-M. Journal of Biomedical Materials Research Part B: Applied Biomaterials 2005, 72B, 156.

126 (91) Kim, K.; Luu, Y. K.; Chang, C.; Fang, D.; Hsiao, B. S.; Chu, B.; Hadjiargyrou, M. J. Controlled Release 2004, 98, 47.

(92) Li, W.-J.; Danielson, K. G.; Alexander, P. G.; Tuan, R. S. J. Biomed. Mater. Res., Part A 2003, 67A, 1105.

(93) Luu, Y. K.; Kim, K.; Hsiao, B. S.; Chu, B.; Hadjiargyrou, M. J. Controlled Release 2003, 89, 341.

(94) Uematsu, K.; Hattori, K.; Ishimoto, Y.; Yamauchi, J.; Habata, T.; Takakura, Y.; Ohgushi, H.; Fukuchi, T.; Sato, M. Biomaterials 2005, 26, 4273.

(95) Yang, F.; Murugan, R.; Wang, S.; Ramakrishna, S. Biomaterials 2005, 26, 2603.

(96)He, W.; Ma, Z.; Teo, W. E.; Dong, Y. X.; Robless, P. A.; Lim, T. C.; Ramakrishna, S. J. Biomed. Mater. Res., Part A 2009, 90A, 205.

(97) Zhang, X.; Wang, X.; Keshav, V.; Wang, X.; Johanas, J. T.; Leisk, G. G.; Kaplan, D. L. Biomaterials 2009, 30, 3213.

(98) Wise, S. G.; Byrom, M. J.; Waterhouse, A.; Bannon, P. G.; Ng, M. K. C.; Weiss, A. S. Acta Biomater. 2011, 7, 295.

(99) Lee, C. H.; Shin, H. J.; Cho, I. H.; Kang, Y.-M.; Kim, I. A.; Park, K.-D.; Shin, J.-W. Biomaterials 2005, 26, 1261.

(100) Schnell, E.; Klinkhammer, K.; Balzer, S.; Brook, G.; Klee, D.; Dalton, P.; Mey, J. Biomaterials 2007, 28, 3012.

(101) Zheng, J.; Kontoveros, D.; Lin, F.; Hua, G.; Reneker, D. H.; Becker, M. L.; Willits, R. K. Biomacromolecules 2015, 16, 357.

(102) Cui, W.; Li, X.; Zhou, S.; Weng, J. J. Biomed. Mater. Res., Part A 2007, 82A, 831.

(103) Zhang, Y.; Venugopal, J. R.; El-Turki, A.; Ramakrishna, S.; Su, B.; Lim, C. T. Biomaterials 2008, 29, 4314.

(104) Hu, X.; Liu, S.; Zhou, G.; Huang, Y.; Xie, Z.; Jing, X. J. Controlled Release 2014, 185, 12.

(105) Williams, G. R.; Chatterton, N. P.; Nazir, T.; Yu, D.-G.; Zhu, L.-M.; Branford-White, C. J. Therapeutic Delivery 2012, 3, 515.

(106) Zamani, M.; Prabhakaran, M. P.; Ramakrishna, S. International Journal of Nanomedicine 2013, 8, 2997.

127 (107) Kenawy, E.-R.; Bowlin, G. L.; Mansfield, K.; Layman, J.; Simpson, D. G.; Sanders, E. H.; Wnek, G. E. J. Controlled Release 2002, 81, 57.

(108) Jiang, H.; Hu, Y.; Li, Y.; Zhao, P.; Zhu, K.; Chen, W. J. Controlled Release 2005, 108, 237.

(109) Liao, I. C.; Chew, S. Y.; Leong, K. W. Nanomedicine 2006, 1, 465.

(110) Liu, J.-J.; Wang, C.-Y.; Wang, J.-G.; Ruan, H.-J.; Fan, C.-Y. J. Biomed. Mater. Res., Part A 2011, 96A, 13.

(111) Sahoo, S.; Ang, L. T.; Goh, J. C.-H.; Toh, S.-L. J. Biomed. Mater. Res., Part A 2010, 93A, 1539.

(112) Nie, H.; Ho, M.-L.; Wang, C.-K.; Wang, C.-H.; Fu, Y.-C. Biomaterials 2009, 30, 892.

(113) Nie, H.; Wang, C.-H. J. Controlled Release 2007, 120, 111.

(114) Giri, J.; Li, W.-J.; Tuan, R. S.; Cicerone, M. T. Advanced Materials 2011, 23, 4861.

(115) Seifu, D. G.; Purnama, A.; Mequanint, K.; Mantovani, D. Nat Rev Cardiol 2013, 10, 410.

(116) Teebken, O. E.; Haverich, A. European Journal of Vascular and Endovascular Surgery 2002, 23, 475.

(117) Levenson, J. W.; Skerrett, P. J.; Gaziano, J. M. Preventive Cardiology 2002, 5, 188.

(118) Diodato, M.; Chedrawy, E. G. Surgery Research and Practice 2014, 2014, 6.

(119) Kim, B.-S.; Mooney, D. J. Trends Biotechnol. 1998, 16, 224.

(120) Zhang, W. J.; Liu, W.; Cui, L.; Cao, Y. Journal of Cellular and Molecular Medicine 2007, 11, 945.

(121) Mano, J. F.; Silva, G. A.; Azevedo, H. S.; Malafaya, P. B.; Sousa, R. A.; Silva, S. S.; Boesel, L. F.; Oliveira, J. M.; Santos, T. C.; Marques, A. P.; Neves, N. M.; Reis, R. L. Journal of The Royal Society Interface 2007, 4, 999.

(122) McClure, M. J.; Sell, S. A.; Simpson, D. G.; Walpoth, B. H.; Bowlin, G. L. Acta Biomater. 2010, 6, 2422.

(123) Thomas, L. V.; Lekshmi, V.; Nair, P. D. International Journal of Cardiology 2013, 167, 1091.

128 (124) Yow, K. H.; Ingram, J.; Korossis, S. A.; Ingham, E.; Homer-Vanniasinkam, S. British Journal of Surgery 2006, 93, 652.

(125) Lee, S. J.; Liu, J.; Oh, S. H.; Soker, S.; Atala, A.; Yoo, J. J. Biomaterials 2008, 29, 2891.

(126) Tillman, B. W.; Yazdani, S. K.; Lee, S. J.; Geary, R. L.; Atala, A.; Yoo, J. J. Biomaterials 2009, 30, 583.

(127) Malafaya, P. B.; Silva, G. A.; Reis, R. L. Adv. Drug Delivery Rev. 2007, 59, 207.

(128) Couet, F.; Rajan, N.; Mantovani, D. Macromol. Biosci. 2007, 7, 701.

(129) Berglund, J. D.; Mohseni, M. M.; Nerem, R. M.; Sambanis, A. Biomaterials 2003, 24, 1241.

(130) Marelli, B.; Achilli, M.; Alessandrino, A.; Freddi, G.; Tanzi, M. C.; Farè, S.; Mantovani, D. Macromol. Biosci. 2012, 12, 1566.

(131) Matthews, J. A.; Wnek, G. E.; Simpson, D. G.; Bowlin, G. L. Biomacromolecules 2002, 3, 232.

(132) Pankajakshan, D.; Agrawal, D. K. Can. J. Physiol. Pharmacol. 2010, 88, 855.

(133) Weinberg, C.; Bell, E. Science 1986, 231, 397.

(134) Boccafoschi, F.; Habermehl, J.; Vesentini, S.; Mantovani, D. Biomaterials 2005, 26, 7410.

(135) Girton, T. S.; Oegema, T. R.; Grassl, E. D.; Isenberg, B. C.; Tranquillo, R. T. Journal of Biomechanical Engineering 2000, 122, 216.

(136) Qi, J.; Chi, L.; Wang, J.; Sumanasinghe, R.; Wall, M.; Tsuzaki, M.; Banes, A. J. Exp. Cell Res. 2009, 315, 1990.

(137) Zhou, J.; Cao, C.; Ma, X.; Lin, J. Int. J. Biol. Macromol. 2010, 47, 514.

(138) Buttafoco, L.; Kolkman, N. G.; Engbers-Buijtenhuijs, P.; Poot, A. A.; Dijkstra, P. J.; Vermes, I.; Feijen, J. Biomaterials 2006, 27, 724.

(139) Cummings, C. L.; Gawlitta, D.; Nerem, R. M.; Stegemann, J. P. Biomaterials 2004, 25, 3699.

(140) Arafat, M. T.; Tronci, G.; Yin, J.; Wood, D. J.; Russell, S. J. Polymer 2015, 77, 102.

129 (141) Takitoh, T.; Bessho, M.; Hirose, M.; Ohgushi, H.; Mori, H.; Hara, M. J. Biosci. Bioeng. 2015, 119, 217.

(142) Tom, S.; Monika, u.; Pavla, S.; Martina, V. n.; Zbyn?k, S.; rka, R. g.; Margit, a.; Radek, S. e.; Marie Hub?lek, K. o. Biomed. Mater. 2015, 10, 065008.

(143) Mosesson, M. W.; Siebenlist, K. R.; Meh, D. A. Ann. N. Y. Acad. Sci. 2001, 936, 11.

(144) Jockenhoevel, S.; Zund, G.; Hoerstrup, S. P.; Chalabi, K.; Sachweh, J. S.; Demircan, L.; Messmer, B. J.; Turina, M. European Journal of Cardio-Thoracic Surgery 2001, 19, 424.

(145) Ye, Q.; Zünd, G.; Benedikt, P.; Jockenhoevel, S.; Hoerstrup, S. P.; Sakyama, S.; Hubbell, J. A.; Turina, M. European Journal of Cardio-Thoracic Surgery 2000, 17, 587.

(146) Long, J. L.; Tranquillo, R. T. Matrix Biol. 2003, 22, 339.

(147) Neidert, M. R.; Lee, E. S.; Oegema, T. R.; Tranquillo, R. T. Biomaterials 2002, 23, 3717.

(148) Grassl, E. D.; Oegema, T. R.; Tranquillo, R. T. Journal of Biomedical Materials Research 2002, 60, 607.

(149) Swartz, D. D.; Russell, J. A.; Andreadis, S. T. American Journal of Physiology - Heart and Circulatory Physiology 2005, 288, H1451.

(150) Badylak, S. F.; Gilbert, T. W. Seminars in Immunology 2008, 20, 109.

(151) Schmidt, C. E.; Baier, J. M. Biomaterials 2000, 21, 2215.

(152) Gilbert, T. W.; Sellaro, T. L.; Badylak, S. F. Biomaterials 2006, 27, 3675.

(153) Mirsadraee, S.; Wilcox, H. E.; Watterson, K. G.; Kearney, J. N.; Hunt, J.; Fisher, J.; Ingham, E. J. Surg. Res. 2007, 143, 407.

(154) Rathore, A.; Cleary, M.; Naito, Y.; Rocco, K.; Breuer, C. Wiley Interdisciplinary Reviews: Nanomedicine and Nanobiotechnology 2012, 4, 257.

(155) Song, J. J.; Ott, H. C. Trends in Molecular Medicine 2011, 17, 424.

(156) Badylak, S. F.; Freytes, D. O.; Gilbert, T. W. Acta Biomater. 2009, 5, 1.

(157) Huynh, T.; Abraham, G.; Murray, J.; Brockbank, K.; Hagen, P.-O.; Sullivan, S. Nat Biotech 1999, 17, 1083.

130 (158) Kaushal, S.; Amiel, G. E.; Guleserian, K. J.; Shapira, O. M.; Perry, T.; Sutherland, F. W.; Rabkin, E.; Moran, A. M.; Schoen, F. J.; Atala, A.; Soker, S.; Bischoff, J.; Mayer, J. E. Nat Med 2001, 7, 1035.

(159) Dahl, S. L. M.; Kypson, A. P.; Lawson, J. H.; Blum, J. L.; Strader, J. T.; Li, Y.; Manson, R. J.; Tente, W. E.; DiBernardo, L.; Hensley, M. T.; Carter, R.; Williams, T. P.; Prichard, H. L.; Dey, M. S.; Begelman, K. G.; Niklason, L. E. Science Translational Medicine 2011, 3, 68ra9.

(160) McKenna, K. A.; Hinds, M. T.; Sarao, R. C.; Wu, P.-C.; Maslen, C. L.; Glanville, R. W.; Babcock, D.; Gregory, K. W. Acta Biomater. 2012, 8, 225.

(161) Koch, S.; Flanagan, T. C.; Sachweh, J. S.; Tanios, F.; Schnoering, H.; Deichmann, T.; Ellä, V.; Kellomäki, M.; Gronloh, N.; Gries, T.; Tolba, R.; Schmitz-Rode, T.; Jockenhoevel, S. Biomaterials 2010, 31, 4731.

(162) Gilding, D. K.; Reed, A. M. Polymer 1979, 20, 1459.

(163) Garlotta, D. J. Polym. Environ. 2001, 9, 63.

(164) de Valence, S.; Tille, J.-C.; Mugnai, D.; Mrowczynski, W.; Gurny, R.; Möller, M.; Walpoth, B. H. Biomaterials 2012, 33, 38.

(165) Williams, S. F.; Martin, D. P.; Horowitz, D. M.; Peoples, O. P. Int. J. Biol. Macromol. 1999, 25, 111.

(166) Zinn, M.; Witholt, B.; Egli, T. Adv. Drug Delivery Rev. 2001, 53, 5.

(167) Chen, G.-Q.; Wu, Q. Biomaterials 2005, 26, 6565.

(168) Rai, R.; Tallawi, M.; Grigore, A.; Boccaccini, A. R. Prog. Polym. Sci. 2012, 37, 1051.

(169) Motlagh, D.; Yang, J.; Lui, K. Y.; Webb, A. R.; Ameer, G. A. Biomaterials 2006, 27, 4315.

(170) Wu, W.; Allen, R. A.; Wang, Y. Nat. Med. 2012, 18, 1148.

(171) Allen, R. A.; Wu, W.; Yao, M.; Dutta, D.; Duan, X.; Bachman, T. N.; Champion, H. C.; Stolz, D. B.; Robertson, A. M.; Kim, K.; Isenberg, J. S.; Wang, Y. Biomaterials 2014, 35, 165.

(172) Generali, M.; Dijkman, P. E.; Hoerstrup, S. P. EMJ Int Cardiol 2014, 1, 91.

(173) Gunatillake, P. A.; Adhikari, R. European cells & materials 2003, 5, 1.

(174) Martin, D. P.; Williams, S. F. Biochemical Engineering Journal 2003, 16, 97. 131 (175) Lazarus, G. S.; Daniels, J. R.; Brown, R. S.; Bladen, H. A.; Fullmer, H. M. J. Clin. Invest. 1968, 47, 2622.

(176) Han, B.; Schwab, I. R.; Madsen, T. K.; Isseroff, R. R. Cornea 2002, 21, 505.

(177) Niklason, L. E.; Gao, J.; Abbott, W. M.; Hirschi, K. K.; Houser, S.; Marini, R.; Langer, R. Science 1999, 284, 489.

(178) Ito, Y.; Kajihara, M.; Imanishi, Y. J Biomed Mater Res 1991, 25, 1325.

(179) Murphy, W. L.; Peters, M. C.; Kohn, D. H.; Mooney, D. J. Biomaterials 2000, 21, 2521.

(180) Ye, Q.; Zund, G.; Jockenhoevel, S.; Schoeberlein, A.; Hoerstrup, S. P.; Grunenfelder, J.; Benedikt, P.; Turina, M. ASAIO journal (American Society for Artificial Internal Organs : 1992) 2000, 46, 730.

(181) Whitaker, M. J.; Quirk, R. A.; Howdle, S. M.; Shakesheff, K. M. J. Pharm. Pharmacol. 2001, 53, 1427.

(182) Lee, K.; Silva, E. A.; Mooney, D. J. Journal of the Royal Society Interface 2011, 8, 153.

(183) Zheng, W.; Wang, Z.; Song, L.; Zhao, Q.; Zhang, J.; Li, D.; Wang, S.; Han, J.; Zheng, X.-L.; Yang, Z.; Kong, D. Biomaterials 2012, 33, 2880.

(184) Pitarresi, G.; Fiorica, C.; Palumbo, F. S.; Rigogliuso, S.; Ghersi, G.; Giammona, G. J. Biomed. Mater. Res., Part A 2014, 102, 1334.

(185) Zhang, H.; Jia, X.; Han, F.; Zhao, J.; Zhao, Y.; Fan, Y.; Yuan, X. Biomaterials 2013, 34, 2202.

(186) Blair, H. C.; Teitelbaum, S. L.; Grosso, L. E.; Lacey, D. L.; Tan, H. L.; McCourt, D. W.; Jeffrey, J. J. Biochem. J. 1993, 290, 873.

(187) Cordewener, F. W.; van Geffen, M. F.; Joziasse, C. A. P.; Schmitz, J. P.; Bos, R. R. M.; Rozema, F. R.; Pennings, A. J. Biomaterials 2000, 21, 2433.

(188) Hakkarainen, M.; Höglund, A.; Odelius, K.; Albertsson, A.-C. J. Am. Chem. Soc. 2007, 129, 6308.

(189) Ramp, W. K.; Lenz, L. G.; Kaysinger, K. K. Bone Miner. 1994, 24, 59.

(190) de Valence, S.; Tille, J. C.; Giliberto, J. P.; Mrowczynski, W.; Gurny, R.; Walpoth, B. H.; Möller, M. Acta Biomater. 2012, 8, 3914.

(191) Kuwabara, F.; Narita, Y.; Yamawaki-Ogata, A.; Satake, M.; Kaneko, H.; Oshima, H.; Usui, A.; Ueda, Y. Journal of Artificial Organs 2012, 15, 399. 132 (192) Roh, J. D.; Nelson, G. N.; Brennan, M. P.; Mirensky, T. L.; Yi, T.; Hazlett, T. F.; Tellides, G.; Sinusas, A. J.; Pober, J. S.; Saltzman, W. M.; Kyriakides, T. R.; Breuer, C. K. Biomaterials 2008, 29, 1454.

(193) Boland, E. D.; Pawlowski, K. J.; Barnes, C. P.; Simpson, D. G.; Wnek, G. E.; Bowlin, G. L. ACS Symposium Series 2006, 918, 188.

(194) Browning, M. B.; Dempsey, D.; Guiza, V.; Becerra, S.; Rivera, J.; Russell, B.; Höök, M.; Clubb, F.; Miller, M.; Fossum, T.; Dong, J. F.; Bergeron, A. L.; Hahn, M.; Cosgriff-Hernandez, E. Acta Biomater. 2012, 8, 1010.

(195) Soldani, G.; Losi, P.; Bernabei, M.; Burchielli, S.; Chiappino, D.; Kull, S.; Briganti, E.; Spiller, D. Biomaterials 2010, 31, 2592.

(196) Yao, Y.; Wang, J.; Cui, Y.; Xu, R.; Wang, Z.; Zhang, J.; Wang, K.; Li, Y.; Zhao, Q.; Kong, D. Acta Biomater. 2014, 10, 2739.

(197) Ryu, J. K.; Kim, H. S.; Nam, D. H. Biotechnol. Bioprocess Eng. 2012, 17, 900.

(198) Mitragotri, S.; Burke, P. A.; Langer, R. Nat Rev Drug Discov 2014, 13, 655.

(199) Lu, Y.; Yang, J.; Sega, E. The AAPS Journal 2006, 8, E466.

(200) Vermonden, T.; Censi, R.; Hennink, W. E. Chemical Reviews 2012, 112, 2853.

(201) Sill, T. J.; von Recum, H. A. Biomaterials 2008, 29, 1989.

(202) Sridhar, R.; Lakshminarayanan, R.; Madhaiyan, K.; Amutha Barathi, V.; Lim, K. H. C.; Ramakrishna, S. Chem. Soc. Rev. 2015, 44, 790.

(203) Szentivanyi, A.; Chakradeo, T.; Zernetsch, H.; Glasmacher, B. Adv. Drug Delivery Rev. 2011, 63, 209.

(204) Chew, S. Y.; Wen, J.; Yim, E. K. F.; Leong, K. W. Biomacromolecules 2005, 6, 2017.

(205) Son, Y. J.; Kim, W. J.; Yoo, H. S. Arch. Pharmacal Res. 2013, 37, 69.

(206) Li, Y.; Jiang, H.; Zhu, K. J. Mater. Sci.: Mater. Med. 2007, 19, 827.

(207) Michel, R.; Pasche, S.; Textor, M.; Castner, D. G. Langmuir 2005, 21, 12327.

(208) Wang, Y.; Annunziata, O. The Journal of Physical Chemistry B 2007, 111, 1222.

133 (209) Asadian, M.; Rashidi, A.; Majidi, M.; Mehrjoo, M.; Emami, B. A.; Tavassoli, H.; Asl, M. P.; Bonakdar, S. Journal of the Iranian Chemical Society 2015, 12, 1089.

(210) Zeng, J.; Aigner, A.; Czubayko, F.; Kissel, T.; Wendorff, J. H.; Greiner, A. Biomacromolecules 2005, 6, 1484.

(211) Yang, Y.; Li, X.; Qi, M.; Zhou, S.; Weng, J. European Journal of Pharmaceutics and Biopharmaceutics 2008, 69, 106.

(212) Schneider, A.; Wang, X. Y.; Kaplan, D. L.; Garlick, J. A.; Egles, C. Acta Biomater. 2009, 5, 2570.

(213) Jiang, H.; Hu, Y.; Zhao, P.; Li, Y.; Zhu, K. Journal of Biomedical Materials Research Part B: Applied Biomaterials 2006, 79B, 50.

(214) Li, H.; Zhao, C.; Wang, Z.; Zhang, H.; Yuan, X.; Kong, D. Journal of biomaterials science. Polymer edition 2010, 21, 803.

(215) Choi, J. S.; Choi, S. H.; Yoo, H. S. J. Mater. Chem. 2011, 21, 5258.

(216) Grafahrend, D.; Calvet, J. L.; Klinkhammer, K.; Salber, J.; Dalton, P. D.; Möller, M.; Klee, D. Biotechnol. Bioeng. 2008, 101, 609.

(217) Ambrose, C.; Clanton, T. Ann. Biomed. Eng. 2004, 32, 171.

(218) Bos, R. R. M.; Rozema, F. B.; Boering, G.; Nijenhius, A. J.; Pennings, A. J.; Verwey, A. B.; Nieuwenhuis, P.; Jansen, H. W. B. Biomaterials 1991, 12, 32.

(219) Bostman, O.; Hirvensalo, E.; Makinen, J.; Rokkanen, P. J. Bone Joint Surg. 1990, 72-B, 592.

(220) Higgins, S. P.; Solan, A. K.; Niklason, L. E. J. Biomed. Mater. Res., Part A 2003, 67A, 295.

(221) Sung, H.-J.; Meredith, C.; Johnson, C.; Galis, Z. S. Biomaterials 2004, 25, 5735.

(222) Srinath, D.; Lin, S.; Knight, D. K.; Rizkalla, A. S.; Mequanint, K. J. Tissue Eng. Regen. Med. 2014, 8, 578.

(223) Gao, Y.; Childers, E. P.; Becker, M. L. ACS Biomaterials Science & Engineering 2015, 1, 795.

(224) Li, S.; Yu, J.; Wade, M. B.; Policastro, G. M.; Becker, M. L. Biomacromolecules 2015.

(225) Lin, F.; Yu, J.; Tang, W.; Zheng, J.; Xie, S.; Becker, M. L. Macromolecules 2013, 46, 9515. 134 (226) Policastro, G. M.; Lin, F.; Smith Callahan, L. A.; Esterle, A.; Graham, M.; Sloan Stakleff, K.; Becker, M. L. Biomacromolecules 2015.

(227) Yu, J.; Lin, F.; Lin, P.; Gao, Y.; Becker, M. L. Macromolecules 2013, 47, 121.

(228) Zhou, J.; Defante, A. P.; Lin, F.; Xu, Y.; Yu, J.; Gao, Y.; Childers, E.; Dhinojwala, A.; Becker, M. L. Biomacromolecules 2014, 16, 266.

(229) Sauerbrey, G. Z. Physik 1959, 155, 206.

(230) Langer, R.; Vacanti, J. Science 1993, 260, 920.

(231) Ravi, S.; Chaikof, E. L. Regen. Med. 2009, 5, 107.

(232) Dvir, T.; Timko, B. P.; Kohane, D. S.; Langer, R. Nat. Nanotechnol. 2011, 6, 13.

(233) Tian, H.; Tang, Z.; Zhuang, X.; Chen, X.; Jing, X. Prog. Polym. Sci. 2012, 37, 237.

(234) Ji, Q.; Zhang, S.; Zhang, J.; Wang, Z.; Wang, J.; Cui, Y.; Pang, L.; Wang, S.; Kong, D.; Zhao, Q. Biomacromolecules 2013, 14, 4099.

(235) Melchiorri, A. J.; Hibino, N.; Yi, T.; Lee, Y. U.; Sugiura, T.; Tara, S.; Shinoka, T.; Breuer, C.; Fisher, J. P. Biomacromolecules 2014, 16, 437.

(236) Wang, Z.; Cui, Y.; Wang, J.; Yang, X.; Wu, Y.; Wang, K.; Gao, X.; Li, D.; Li, Y.; Zheng, X.-L.; Zhu, Y.; Kong, D.; Zhao, Q. Biomaterials 2014, 35, 5700.

(237) Wang, S.; Zhang, Y.; Wang, H.; Yin, G.; Dong, Z. Biomacromolecules 2009, 10, 2240.

(238) Dahl, S. L. M.; Koh, J.; Prabhakar, V.; Niklason, L. E. Cell Transplant. 2003, 12, 659.

(239) Wang, C.; Cen, L.; Yin, S.; Liu, Q.; Liu, W.; Cao, Y.; Cui, L. Biomaterials 2010, 31, 621.

(240) Mooney, D. J.; Mazzoni, C. L.; Breuer, C.; McNamara, K.; Hern, D.; Vacanti, J. P.; Langer, R. Biomaterials 1996, 17, 115.

(241) Shin'oka, T.; Imai, Y.; Ikada, Y. N. Engl. J. Med. 2001, 344, 532.

(242) Dargaville, B. L.; Vaquette, C.; Peng, H.; Rasoul, F.; Chau, Y. Q.; Cooper-White, J. J.; Campbell, J. H.; Whittaker, A. K. Biomacromolecules 2011, 12, 3856.

135 (243) Liu, H.; Li, X.; Niu, X.; Zhou, G.; Li, P.; Fan, Y. Biomacromolecules 2011, 12, 2914.

(244) Pitt, C. G.; Chasalow, F. I.; Hibionada, Y. M.; Klimas, D. M.; Schindler, A. J. Appl. Polym. Sci. 1981, 26, 3779.

(245) Sun, H.; Mei, L.; Song, C.; Cui, X.; Wang, P. Biomaterials 2006, 27, 1735.

(246) Amsden, B.; Wang, S.; Wyss, U. Biomacromolecules 2004, 5, 1399.

(247) Höglund, A.; Odelius, K.; Hakkarainen, M.; Albertsson, A.-C. Biomacromolecules 2007, 8, 2025.

(248) Neffe, A. T.; Tronci, G.; Alteheld, A.; Lendlein, A. Macromol. Chem. Phys. 2010, 211, 182.

(249) Pitt, C. G.; Hendren, R. W.; Schindler, A.; Woodward, S. C. J. Controlled Release 1984, 1, 3.

(250) Younes, H. M.; Bravo-Grimaldo, E.; Amsden, B. G. Biomaterials 2004, 25, 5261.

(251) Walpoth, B. H.; Bowlin, G. L. Expert Rev. Med. Devices 2005, 2, 647.

(252) Katsarava, R.; Beridze, V.; Arabuli, N.; Kharadze, D.; Chu, C. C.; Won, C. Y. J. Polym. Sci. A Polym. Chem. 1999, 37, 391.

(253) Pang, X.; Chu, C.-C. Biomaterials 2010, 31, 3745.

(254) Karimi, P.; Rizkalla, A. S.; Mequanint, K. Materials 2010, 3, 2346.

(255) Kurane, A.; Simionescu, D. T.; Vyavahare, N. R. Biomaterials 2007, 28, 2830.

(256) Sell, S. A.; McClure, M. J.; Barnes, C. P.; Knapp, D. C.; Walpoth, B. H.; Simpson, D. G.; Bowlin, G. L. Biomed. Mater. 2006, 1, 72.

(257) Teng, Z.; Tang, D.; Zheng, J.; Woodard, P. K.; Hoffman, A. H. J. Biomech. 2009, 42, 2535.

(258) Weir, N. A.; Buchanan, F. J.; Orr, J. F.; Dickson, G. R. Proc. Inst. Mech. Eng. H 2004, 218, 307.

(259) Weir, N. A.; Buchanan, F. J.; Orr, J. F.; Farrar, D. F.; Dickson, G. R. Proc. Inst. Mech. Eng. H 2004, 218, 321.

(260) Monta ez, M. I.; Hed, .; Utsel, .; Ropponen, .; Malmstr m, E.; gberg, L.; Hult, A.; Malkoch, M. Biomacromolecules 2011, 12, 2114. 136 (261) Orelma, H.; Teerinen, T.; Johansson, L.-S.; Holappa, S.; Laine, J. Biomacromolecules 2012, 13, 1051.

(262) Zhang, Y.; Luo, S.; Tang, Y.; Yu, L.; Hou, K.-Y.; Cheng, J.-P.; Zeng, X.; Wang, P. G. Anal. Chem. 2006, 78, 2001.

(263) Yusuf, S.; Reddy, S.; Ôunpuu, S.; Anand, S. Circulation 2001, 104, 2855.

(264) Sell, S. A.; Bowlin, G. L. J. Mater. Chem. 2008, 18, 260.

(265) Canver, C. C. Chest 1995, 108, 1150.

(266) Perera, G. B.; Mueller, M. P.; Kubaska, S. M.; Wilson, S. E.; Lawrence, P. F.; Fujitani, R. M. Annals of vascular surgery 2004, 18, 66.

(267) Kannan, R. Y.; Salacinski, H. J.; Edirisinghe, M. J.; Hamilton, G.; Seifalian, A. M. Biomaterials 2006, 27, 4618.

(268) Lovett, M.; Cannizzaro, C.; Daheron, L.; Messmer, B.; Vunjak-Novakovic, G.; Kaplan, D. L. Biomaterials 2007, 28, 5271.

(269) Tiwari, A.; Salacinski, H.; Seifalian, A. M.; Hamilton, G. Cardiovascular Surgery 2002, 10, 191.

(270) Tara, S.; Kurobe, H.; Maxfield, M. W.; Rocco, K. A.; Yi, T.; Naito, Y.; Breuer, C. K.; Shinoka, T. Journal of Vascular Surgery 2015, 62, 734.

(271) Hajiali, H.; Shahgasempour, S.; Naimi-Jamal, M. R.; Peirovi, H. International Journal of Nanomedicine 2011, 6, 2133.

(272) Torikai, K.; Ichikawa, H.; Hirakawa, K.; Matsumiya, G.; Kuratani, T.; Iwai, S.; Saito, A.; Kawaguchi, N.; Matsuura, N.; Sawa, Y. The Journal of Thoracic and Cardiovascular Surgery 2008, 136, 37.

(273) Wen, S. J.; Zhao, L. M.; Wang, S. G.; Li, J. X.; Chen, H. Y.; Liu, J. L.; Liu, Y.; Luo, Y.; Changizi, R. Chinese medical journal 2007, 120, 1331.

(274) Gloria, A.; Causa, F.; Russo, T.; Battista, E.; Della Moglie, R.; Zeppetelli, S.; De Santis, R.; Netti, P. A.; Ambrosio, L. Biomacromolecules 2012, 13, 3510.

(275) Lee, K. W.; Wang, Y. Journal of visualized experiments 2011, 50. e2691.

(276) Horwitz, J. A.; Shum, K. M.; Bodle, J. C.; Deng, M.; Chu, C.-C.; Reinhart-King, C. A. J. Biomed. Mater. Res., Part A 2010, 95A, 371.

(277) Knight, D. K.; Gillies, E. R.; Mequanint, K. Acta Biomater. 2014, 10, 3484.

137 (278) van der Lei, B.; Bartels, H. L.; Nieuwenhuis, P.; Wildevuur, C. R. Surgery 1985, 98, 955.

(279) Ye, S.-H.; Hong, Y.; Sakaguchi, H.; Shankarraman, V.; Luketich, S. K.; D’Amore, A.; agner, . R. ACS Applied Materials & Interfaces 2014, 6, 22796.

(280) Boccafoschi, F.; Rajan, N.; Habermehl, J.; Mantovani, D. Macromol. Biosci. 2007, 7, 719.

(281) Catto, V.; Farè, S.; Cattaneo, I.; Figliuzzi, M.; Alessandrino, A.; Freddi, G.; Remuzzi, A.; Tanzi, M. C. Materials Science and Engineering: C 2015, 54, 101.

(282) Gui, L.; Boyle, M. J.; Kamin, Y. M.; Huang, A. H.; Starcher, B. C.; Miller, C. A.; Vishnevetsky, M. J.; Niklason, L. E. Tissue Engineering. Part A 2014, 20, 1499.

(283) Lovett, M.; Eng, G.; Kluge, J. A.; Cannizzaro, C.; Vunjak-Novakovic, G.; Kaplan, D. L. Organogenesis 2010, 6, 217.

(284) Ahn, H.; Ju, Y. M.; Takahashi, H.; Williams, D. F.; Yoo, J. J.; Lee, S. J.; Okano, T.; Atala, A. Acta Biomater. 2015, 16, 14.

(285) Caves, J. M.; Kumar, V. A.; Martinez, A. W.; Kim, J.; Ripberger, C. M.; Haller, C. A.; Chaikof, E. L. Biomaterials 2010, 31, 7175.

(286) Han, J.; Lazarovici, P.; Pomerantz, C.; Chen, X.; Wei, Y.; Lelkes, P. I. Biomacromolecules 2011, 12, 399.

(287) Matsumura, G.; Nitta, N.; Matsuda, S.; Sakamoto, Y.; Isayama, N.; Yamazaki, K.; Ikada, Y. PLoS One 2012, 7, e35760.

(288) Brennan, M. P.; Dardik, A.; Hibino, N.; Roh, J. D.; Nelson, G. N.; Papademitris, X.; Shinoka, T.; Breuer, C. K. Annals of surgery 2008, 248, 370.

(289) Hibino, N.; hin’oka, T.; Matsumura, G.; Ikada, .; Kurosawa, H. The Journal of Thoracic and Cardiovascular Surgery 2005, 129, 1064.

(290) Matsumura, G.; Miyagawa-Tomita, S.; Shin'oka, T.; Ikada, Y.; Kurosawa, H. Circulation 2003, 108, 1729.

(291) Hibino, N.; McGillicuddy, E.; Matsumura, G.; Ichihara, Y.; Naito, Y.; Breuer, C.; Shinoka, T. The Journal of Thoracic and Cardiovascular Surgery 2010, 139, 431.

(292) hin’oka, T.; Matsumura, G.; Hibino, N.; Naito, .; atanabe, M.; Konuma, T.; Sakamoto, T.; Nagatsu, M.; Kurosawa, H. The Journal of Thoracic and Cardiovascular Surgery 2005, 129, 1330.

(293) Shin'oka , T.; Imai , Y.; Ikada , Y. N. Engl. J. Med. 2001, 344, 532.

138 (294) Tara, S.; Kurobe, H.; Maxfield, M. W.; Rocco, K. A.; Yi, T.; Naito, Y.; Breuer, C. K.; Shinoka, T. Journal of Vascular Surgery 2015, 62, 734.

(295) Liang, D.; Hsiao, B. S.; Chu, B. Adv. Drug Delivery Rev. 2007, 59, 1392.

(296) Ma, Z.; Kotaki, M.; Inai, R.; Ramakrishna, S. Tissue engineering 2005, 11, 101.

(297) Sell, S. A.; McClure, M. J.; Garg, K.; Wolfe, P. S.; Bowlin, G. L. Adv. Drug Delivery Rev. 2009, 61, 1007.

(298) Billiar, K.; Murray, J.; Laude, D.; Abraham, G.; Bachrach, N. Journal of Biomedical Materials Research 2001, 56, 101.

(299) Carter, P. J. Exp. Cell Res. 2011, 317, 1261.

(300) Leader, B.; Baca, Q. J.; Golan, D. E. Nat Rev Drug Discov 2008, 7, 21.

(301) Stumpp, M. T.; Binz, H. K.; Amstutz, P. Drug Discov. Today 2008, 13, 695.

(302) Walsh, G. Nat Biotech 2010, 28, 917.

(303) Johannsson, G.; Mårin, P.; Lönn, L.; Ottosson, M.; Stenlöf, K.; Björntorp, P.; Sjöström, L.; Bengtsson, B.-Å. The Journal of Clinical Endocrinology & Metabolism 1997, 82, 727.

(304) McGauley, G. A. Acta Pædiatrica 1989, 78, 70.

(305) Schlechter, N. L.; Russell, S. M.; Spencer, E. M.; Nicoll, C. S. Proceedings of the National Academy of Sciences 1986, 83, 7932.

(306) Betts, P. R.; Butler, G. E.; Donaldson, M. D. C.; Dunger, D. B.; Johnston, D. I.; Kelnar, C. J. H.; Kirk, J.; Price, D. A.; Wilton, P. Archives of Disease in Childhood 1999, 80, 221.

(307) Carroll, P. V.; Christ the members of Growth Hormone Research Society Scientific Committee, E. R.; Bengtsson, B. Å.; Carlsson, L.; Christiansen, J. S.; Clemmons, D.; Hintz, R.; Ho, K.; Laron, Z.; Sizonenko, P.; Sönksen, P. H.; Tanaka, T.; Thorner, M. The Journal of Clinical Endocrinology & Metabolism 1998, 83, 382.

(308) Fine, R.; Pyke-Grimm, K.; Nelson, P.; Boechat, M. I.; Lippe, B.; Yadin, O.; Kamil, E. Pediatr Nephrol 1991, 5, 477.

(309) Sandberg, D. E.; Brook, A. E.; Campos, S. P. Pediatrics 1994, 94, 832.

(310) Spiliotis, B. E. Therapeutics and Clinical Risk Management 2008, 4, 1177.

139 (311) Rothenbuhler, A.; Ormières, B.; Kalifa, G.; Bougnères, P. Growth Hormone & IGF Research 2015, 25, 96.

(312) Bond, W. S.; Hussar, D. A. American journal of hospital pharmacy 1991, 48, 1978.

(313) Kim, S. J.; Hahn, S. K.; Kim, M. J.; Kim, D. H.; Lee, Y. P. J. Controlled Release 2005, 104, 323.

(314) Kwak, H. H.; Shim, W. S.; Choi, M. K.; Son, M. K.; Kim, Y. J.; Yang, H. C.; Kim, T. H.; Lee, G. I.; Kim, B. M.; Kang, S. H.; Shim, C. K. J. Controlled Release 2009, 137, 160.

(315) Smith, S. L.; Hindmarsh, P. C.; Brook, C. G. Archives of Disease in Childhood 1993, 68, 91.

(316) Lee, E. N.; Kim, Y. M.; Lee, H. J.; Park, S. W.; Jung, H. Y.; Lee, J. M.; Ahn, Y. H.; Kim, J. Pharm. Res. 2005, 22, 1735.

(317) Schellenberger, V.; Wang, C.-w.; Geething, N. C.; Spink, B. J.; Campbell, A.; To, W.; Scholle, M. D.; Yin, Y.; Yao, Y.; Bogin, O.; Cleland, J. L.; Silverman, J.; Stemmer, W. P. C. Nat Biotech 2009, 27, 1186.

(318) Kim, E.-S.; Jang, D. S.; Yang, S. Y.; Lee, M. N.; Jin, K. S.; Cha, H. J.; Kim, J. K.; Sung, Y. C.; Choi, K. Y. Nanoscale 2013, 5, 4262.

(319) Govardhan, C.; Khalaf, N.; Jung, C.; Simeone, B.; Higbie, A.; Qu, S.; Chemmalil, L.; Pechenov, S.; Basu, S.; Margolin, A. Pharm. Res. 2005, 22, 1461.

(320) Salmaso, S.; Bersani, S.; Scomparin, A.; Balasso, A.; Brazzale, C.; Barattin, M.; Caliceti, P. J. Controlled Release 2014, 194, 168.

(321) Jordan, F.; Naylor, A.; Kelly, C. A.; Howdle, S. M.; Lewis, A.; Illum, L. J. Controlled Release 2010, 141, 153.

(322) Takada, S.; Yamagata, Y.; Misaki, M.; Taira, K.; Kurokawa, T. J. Controlled Release 2003, 88, 229.

(323) Johnson, O.; Jaworowicz, W.; Cleland, J.; Bailey, L.; Charnis, M.; Duenas, E.; Wu, C.; Shepard, D.; Magil, S.; Last, T.; Jones, A. S.; Putney, S. Pharm. Res. 1997, 14, 730.

(324) Cleland, J. L.; Duenas, E.; Daugherty, A.; Marian, M.; Yang, J.; Wilson, M.; Celniker, A. C.; Shahzamani, A.; Quarmby, V.; Chu, H.; Mukku, V.; Mac, A.; Roussakis, M.; Gillette, N.; Boyd, B.; Yeung, D.; Brooks, D.; Maa, Y.-F.; Hsu, C.; Jones, A. J. S. J. Controlled Release 1997, 49, 193.

140 (325) Johnson, O. L.; Cleland, J. L.; Lee, H. J.; Charnis, M.; Duenas, E.; Jaworowicz, W.; Shepard, D.; Shahzamani, A.; Jones, A. J. S.; Putney, S. D. Nat Med 1996, 2, 795.

(326) Wei, Y.; Wang, Y.; Kang, A.; Wang, W.; Ho, S. V.; Gao, J.; Ma, G.; Su, Z. Mol. Pharm. 2012, 9, 2039.

(327) Singh, N. K.; Nguyen, Q. V.; Kim, B. S.; Lee, D. S. Nanoscale 2015, 7, 3043.

(328) Seo, B.-B.; Park, M.-R.; Chun, C.; Lee, J.-Y.; Song, S.-C. Biomaterials 2011, 32, 8271.

(329) Huynh, C. T.; Kang, S. W.; Li, Y.; Kim, B. S.; Lee, D. S. Soft Matter 2011, 7, 8984.

(330) Park, M.-R.; Chun, C.; Ahn, S.-W.; Ki, M.-H.; Cho, C.-S.; Song, S.-C. J. Controlled Release 2010, 147, 359.

(331) Tae, G.; Kornfield, J. A.; Hubbell, J. A. Biomaterials 2005, 26, 5259.

(332) Kolate, A.; Baradia, D.; Patil, S.; Vhora, I.; Kore, G.; Misra, A. J. Controlled Release 2014, 192, 67.

(333) Nemirovskiy, O.; Zheng, Y. J.; Tung, D.; Korniski, B.; Settle, S.; Skepner, A.; Yates, M.; Aggarwal, P.; Sunyer, T.; Aguiar, D. J. Xenobiotica 2010, 40, 586.

(334) Touraine, P.; D'Souza, G. A.; Kourides, I.; Abs, R.; Barclay, P.; Xie, R.; Pico, A.; Torres-Vela, E.; Ekman, B. Eur. J. Endocrinol. 2009, 161, 533.

(335) Webster, R.; Xie, R.; Didier, E.; Finn, R.; Finnessy, J.; Edgington, A.; Walker, D. Xenobiotica 2008, 38, 1340.

(336) Liao, I. C.; Chen, S.; Liu, J. B.; Leong, K. W. J. Controlled Release 2009, 139, 48.

(337) Kim, H. S.; Yoo, H. S. J. Controlled Release 2010, 145, 264.

(338) Mathew, C. C.; Jeffrey, S. W.; Norman, C. B. Polymer Degradation and Performance; American Chemical Society, 2009; Vol. 1004.

(339) Valmikinathan, C. M.; Defroda, S.; Yu, X. Biomacromolecules 2009, 10, 1084.

141 APPENDIX

Figure A. 1. DSC of L-leucine-based poly(ester urea)s with different chain length of diols.

142