Poly(vinyl alcohol) and heparin hydrogels: Synthesis, structure and presentation of signalling molecules for growth factor activation

by

Anastasia Nilasaroya

A thesis submitted for the degree of Doctor of Philosophy

Graduate School of Biomedical Engineering

University of New South Wales

March 2010

COPYRIGHT STATEMENT

‘I hereby grant the University of New South Wales or its agents the right to archive and to make available my thesis or dissertation in whole or part in the University libraries in all forms of media, now or here after known, subject to the provisions of the Copyright Act 1968. I retain all proprietary rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation. I also authorise University Microfilms to use the 350 word abstract of my thesis in Dissertation Abstract International (this is applicable to doctoral theses only). I have either used no substantial portions of copyright material in my thesis or I have obtained permission to use copyright material; where permission has not been granted I have applied/will apply for a partial restriction of the digital copy of my thesis or dissertation.'

Signed Anastasia Nilasaroya

Date 11/10/2010

AUTHENTICITY STATEMENT

‘I certify that the Library deposit digital copy is a direct equivalent of the final officially approved version of my thesis. No emendation of content has occurred and if there are any minor variations in formatting, they are the result of the conversion to digital format.’

Signed Anastasia Nilasaroya

Date 11/10/2010

ORIGINALITY STATEMENT

‘I hereby declare that this submission is my own work and to the best of my knowledge it contains no materials previously published or written by another person, or substantial proportions of material which have been accepted for the award of any other degree or diploma at UNSW or any other educational institution, except where due acknowledgement is made in the thesis. Any contribution made to the research by others, with whom I have worked at UNSW or elsewhere, is explicitly acknowledged in the thesis. I also declare that the intellectual content of this thesis is the product of my own work, except to the extent that assistance from others in the project's design and conception or in style, presentation and linguistic expression is acknowledged.’

Signed Anastasia Nilasaroya

Date 11/10/2010

Abstract

he design of scaffold materials that could be used to assist tissue regeneration has Tbeen based on the extracellular matrix (ECM), in structure and in the presentation of molecular cues. Synthetic hydrogels resemble the ECM with their structure, high water content, and elasticity, but they lack bioactivity and recognition.

Therefore, the incorporation of biomolecules into synthetic hydrogels is a growing current area of research. Heparan sulfate (HS), a glycosaminoglycan present in basement membrane, has been known to bind and signal various ECM proteins.

Sustained presentation of biomolecules like HS could be achieved by covalent linking to scaffolds; however it poses a challenge on the preservation and expression of the biomolecules’ activity after incorporation.

Biosynthetic hydrogels derived from heparin (a model for HS) and poly(vinyl alcohol) (PVA) were formed by photopolymerisation. PVA and heparin were both functionalised with photopolymerisable methacrylate groups prior to crosslinking, and the effect of this modification on heparin was examined. PVA/heparin co-hydrogels were made with varying compositions and assessed in terms of their structure, strength and growth factor presentation. The activity of the co-hydrogels following incubation with platelet extract (PE) was also studied, to simulate responses that might occur when the hydrogels, as tissue engineered scaffolds, come in contact with blood products.

Heparin remained structurally intact and biologically active following methacrylate modification and UV exposure. The addition of up to 2.5 wt% of heparin

- i -

Abstract increased the hydrogel swelling capacity without compromising the strength of the resulting hydrogel network. The specific FGF-2-signalling activity of heparin in the

PVA/heparin co-hydrogels was demonstrated, with results indicating that co-hydrogels may be formulated with a minimal amount of heparin (≥0.05 wt%), thus limiting any effects on structural integrity. PE treatment of the hydrogel-bound heparin diminished its anticoagulation properties but increased the FGF-2 signalling, suggesting the heparanase activity in PE cleave at the antithrombin binding site to yield fragments that can signal cell receptors. This work has demonstrated the formation of biosynthetic co- hydrogels, capable of presenting growth factors to cells, and provides a novel insight on the molecular activation of heparin-based hydrogels upon enzymatic degradation.

- ii -

Acknowledgements

Finally I am presenting you, dear reader, with my thesis. After what has been the best and the worst time of my life, the journey has come to an end. I thank God for leading me through, and the incredible people that I met along the way.

My most unreserved thanks go to my supervisor, Penny Martens, whose constant support has made the completion of this thesis possible. Thank you for persevering with me and continuing to believe in my ability. I would like to equally thank my co- supervisor, John Whitelock, for his endless stream of ideas and for his invaluable insight into the world of GAGs.

Special thanks to Laura Poole-Warren for being my ‘unofficial’ supervisor and for her input during various stages of the project; to Ross Odell for, yes, his help in statistics and words of encouragement for the project and in teaching; and to Lynn

Ferris for always making the lab so much easier to work in.

Fellow students, current and former, provided much emotional support and help in the progression of the project. I thank Christine Chuang and Ruby Estrella for their early companion in the lab; Megan Lord for help in many things cell-related; Litania Lie and Bill Cheng for help towards the completion of my experimental work. Thank you to

Brooke Farrugia, Nicole Fong, MoonSun Jung, Andreas Permadi, Natalia Pramana and

Miriam van Kalsbeek for their friendship and the much needed coffee breaks.

At last, to those who have had to put up with me the most during this journey. My parents, with their innocent questioning on when I would finish this thesis. My sister

Stella, with her impatient questioning on the subject, while still helping me with some illustration works. And my dear Wee Yong, who has always been a pillar of strength. I thank you all for your constant love and care through and beyond this journey.

- iii -

List of Publications

Journal publications:

1. Nilasaroya A, Poole-Warren LA, Whitelock JM, Martens PJ. Structural and functional characterisation of poly(vinyl alcohol) and heparin hydrogels. Biomaterials 2008, 29, 4658–4664.

2. Martens P, Blundo J, Nilasaroya A, Odell RA, Cooper-White J, Poole-Warren LA. Effect of poly(vinyl alcohol) macromer chemistry and chain interactions on hydrogel mechanical properties. Chemistry of Materials 2007, 19, 2641–2648.

3. Rees MD, Whitelock JM, Malle E, Chuang CY, Iozzo RV, Nilasaroya A, Davies MJ. Myeloperoxidase-derived oxidants selectively disrupt the protein core of the heparan sulfate proteoglycan perlecan. Matrix Biology 2009, 29, 63–73.

4. Martens P, Grant M, Nilasaroya A, Whitelock J, Poole-Warren L. Characterisation of redox initiators for producing poly(vinyl alcohol) hydrogels. Macromolecular Symposia 2008, 266, 59–62.

Conference presentations:

1. Nilasaroya A, Poole-Warren LA, Whitelock JM and Martens P. Characterisation of Poly(Vinyl Alcohol) and Heparin Hydrogels Formed from Methacrylated Macromers. 17th Annual Conference of the Australasian Society for Biomaterials, Melbourne, Australia, 11–13 April 2007.

2. Nilasaroya A, Poole-Warren LA, Whitelock JM, Martens P. Poly(Vinyl Alcohol) and Heparin Hydrogels: Characterization and Functional Study. International Congress on Biohydrogels, Viareggio, Italy, 14–18 November 2007.

- iv -

List of Publications

3. Nilasaroya A, Poole-Warren LA, Whitelock JM, Martens P. Structural and Functional Characterisation of Poly(Vinyl Alcohol) and Heparin Copolymers. 8th World Biomaterials Congress, Amsterdam, the Netherlands, 28 May–1 June 2008.

4. Nilasaroya A, Poole-Warren LA, Whitelock JM, Martens P. (Poster) Structural and Functional Characterisation of Poly (Vinyl Alcohol) and Heparin Copolymers. Australasian Society for Biomaterials and Tissue Engineering Symposium, Canberra, Australia, 22 February 2008.

5. Nilasaroya A, Poole-Warren LA, Whitelock J, Martens P. (Poster) Poly(Vinyl Alcohol)–Heparin Hydrogels: Growth Factor Activation and Degradation. 3rd Indo-Australian Conference on Biomaterials, Implants, Tissue Engineering & Regenerative Medicine/ 19th Annual Conference of Australasian Society for Biomaterials and Tissue Engineering, Sydney, Australia, 21–23 January 2009.

6. Poole-Warren LA, Nilasaroya A, Grant M, Whitelock JM, Martens P. Biosynthetic Hydrogels as Injectable Soft-tissue Engineering Scaffolds. Tissue Engineering & Regenerative Medicine International Society – Asia Pacific Chapter Meeting, Tokyo, Japan, 3–5 December 2007.

7. Whitelock J, Davies N, Jung MS, Nilasaroya A, Chuang C, Neilsen N, Martens P, Poole-Warren L, Lyons JG, Melrose J. Heparan Sulfate as a Biosignalling Molecule for Tissue Engineering Applications. Tissue Engineering & Regenerative Medicine International Society – Asia Pacific Chapter Meeting, Tokyo, Japan, 3–5 December 2007.

8. Martens P, Nilasaroya A, Poole-Warren L, Whitelock J. Poly(Vinyl Alcohol) and Heparin Co-Polymers. International Workshop on Biomaterials for Tissue Engineering and Biotechnological Applications, Kharagpur, India, 22–24 November 2008.

9. Martens P, Nilasaroya A, Reeves A, Kundu J, Kundu SC, Whitelock J, Poole- Warren LA. Poly (Vinyl Alcohol) Hydrogels Improving Biological Performance. 22nd European Conference of Biomaterials, Lausanne, Switzerland, 7–11 September 2009.

- v -

Table of Contents

Abstract i Acknowledgements iii List of Publications iv Table of Contents vi List of Figures and Schemes x List of Tables xiii Common Abbreviations and Terminology xiv

Chapter 1 Introduction 1 Aims of Thesis 3

Chapter 2 Literature Review 5 2.1 Tissue Engineered Constructs 6 2.1.1 Molecular Interaction 7 2.1.2 Vascularisation of Constructs 9 2.2 Biosynthetic Materials 13 2.2.1 Physical Incorporation of Signalling Molecules into Scaffolds 14 2.2.2 Covalent Incorporation of Signalling Molecules into Scaffolds 17 2.2.2.1 EDC/NHS chemistry 18 2.2.2.2 Thiol modification 21 2.2.2.3 Lactose-modified chitosan 23 2.2.2.4 Methacrylate modification 24 2.3 Poly(Vinyl Alcohol) (PVA) Hydrogels 26 2.5.1 Structure and Properties 26 2.5.2 Biomedical Applications 28 2.4 Heparan Sulfate Proteoglycans 30 2.4.1 Biosynthesis 31 2.4.2 Protein Interactions 33 2.4.2.1 Interactions with growth factors 34 2.4.3 Role in Vascularisation 36

- vi -

Table of Contents

2.5 Summary 38

Chapter 3 From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment 39 3.1 Background and Aims 40 3.2 Experimental 43 3.2.1 Materials 43 3.2.2 Synthesis of PVA Macromers 44 3.2.3 Synthesis of Heparin Macromers 45 3.2.4 Characterisation of Methacrylated Macromers 45 3.2.5 Hydrogel Formation 47 3.2.6 Tensile Testing of PVA Hydrogels 48 3.2.6 Cell Growth Inhibition Assay 49

3.3 Results and Discussion 51 3.3.1 Synthesis of PVA Macromers 51 3.3.2 Effect of Macromer Attachment on the Cell Compatibility of PVA 53 3.3.3 PVA Hydrogels from GMA- and ICEMA-Synthesised Macromers 54 3.3.3.1 Hydrogel swelling and sol fraction 54 3.3.3.2 Mechanical properties 57 3.3.3.3 Cell growth inhibition 58 3.3.4 Synthesis of Heparin Macromers 61 3.3.5 UV-Polymerisation Conditions for PVA and Heparin Macromers 64 3.4 Conclusion 67

Chapter 4 Methacrylate–Modified Heparin for Hydrogel Incorporation 68 4.1 Background and Aims 69 4.2 Experimental 71 4.2.1 Materials 71 4.2.2 Size Exclusion Chromatography 72 4.2.3 Clotting Time Assay 73 4.2.4 BaF3 Cell Proliferation Assay 73 4.3 Results and Discussion 75 4.3.1 Structural Analysis of Methacrylated Heparin 75

- vii -

Table of Contents

4.3.2 Antithrombin III Activation 77 4.3.3 Cellular Signalling of FGF-2 79 4.4 Conclusion 83 Acknowledgements 84

Chapter 5 Structural and Functional Characterisation of Poly(Vinyl Alcohol)/Heparin Co-Hydrogels 85 5.1 Background and Aims 86 5.2 Experimental 89 5.2.1 Materials 89 5.2.2 Hydrogel Formation 89 5.2.3 Hydrogel Characterisation 90 5.2.3.1 Swelling study and mass loss analysis 90 5.2.3.2 Composition of co-hydrogel mass loss 91 5.2.3.3 Final composition of co-hydrogels at equilibrium 92 5.2.3.4 Mechanical testing 93 5.2.4 Bioactivity of PVA/Heparin Co-Hydrogels 93 5.2.4.1 Antithrombin III activation 93 5.2.4.2 Cellular signalling of FGF-2 94 5.3 Results and Discussion 94 5.2.1 Hydrogel Characterisation 94 5.3.1.1 Swelling study and mass loss analysis 94 5.3.1.2 Equilibrium mass loss and co-hydrogel compositions 98 5.3.1.3 Mechanical properties 101 5.3.2 Bioactivity of PVA/Heparin Co-Hydrogels 103 5.3.2.1 Anticoagulation property 104 5.3.2.2 Growth factor activation 108 5.4 Conclusion 112 Acknowledgements 113

Chapter 6 Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity 114 6.1 Background and Aims 115

- viii -

Table of Contents

6.2 Experimental 117 6.2.1 Materials 117 6.2.2 Extraction of Heparanase from Platelets 118 6.2.3 Heparanase Activity of PE on Heparin 120 6.2.4 Effect of Molecular Weight on Heparin Activity 121 6.2.5 Enzymatic Degradation Profile of Heparin from Hydrogels 122 6.3 Results and Discussion 123 6.3.1 Heparanase Activity of the Platelet Extract 123 6.3.1.1 Amount of heparin degradation following PE treatment 124 6.3.1.2 Molecular weight of PE-treated heparin 127 6.3.2 Effect of Molecular Weight on Heparin Activity 129 6.3.3 Enzymatic Degradation Profile of Heparin from the Hydrogels 136 6.4 Conclusion 141 Acknowledgements 142

Chapter 7 Conclusions and Future Work 143 Summary of Contributions 146 Recommendations for Future Work 147

Bibliography 150

- ix -

List of Figures and Schemes

List of Figures and Schemes

Figure 2.1 The general strategy for growth factor delivery from degradable matrices. . 16

Figure 2.2 Covalently bound GAGs facilitate growth factor binding onto scaffold ...... 18

Figure 2.3 Formation of PVA hydrogels by physical and covalent crosslinking ...... 27

Figure 2.4 The basic structure of a heparan sulfate proteoglycan...... 32

Figure 2.5 The release of HS-bound growth factors by enzymatic cleavage of HS chains

...... 37

Figure 3.1 Photocrosslinking of polymers carrying multiple pendant functional

groups ...... 41

1 Figure 3.2 H NMR of PVA-methacrylate in D2O ...... 46

1 Figure 3.3 H NMR of heparin-methacrylate in D2O ...... 47

Figure 3.4 Hydrogel specimen for immersed tensile testing...... 49

Figure 3.5 L929 cell compatibility of PVA macromers ...... 53

Figure 3.6 L929 cell compatibility to the PVA hydrogel extracts ...... 59

Figure 3.7 Methacrylate group attachment as a function of reaction time with glycidyl

methacrylate ...... 63

Figure 3.8 Photopolymerisation efficiency as a function of UV intensity and UV

exposure time ...... 65

Figure 4.1 Molecular weight distribution profiles of hep-MA and heparin standards .... 76

Figure 4.2 Illustration of the binding of proteins to specific sequences on heparin

chain ...... 77

Figure 4.3 Plasma clotting time as a function of heparin and hep-MA concentrations .. 78

- x -

List of Figures and Schemes

Figure 4.4 Illustration of a BaF3 cell and its cell surface receptors ...... 80

Figure 4.5 BaF3 cell proliferation as a function of heparin and FGF-2 concentrations . 82

Figure 4.6 Comparison of FGF-2 signalling activity of heparin and hep-MA ...... 83

Figure 5.1 Comparison of heparin loss from PVA/heparin 17.5/2.5 hydrogels when

heparin was covalently linked and when physically blended ...... 95

Figure 5.2 Swelling and mass loss profiles of PVA, heparin, and their copolymer

hydrogels...... 96

Figure 5.3 PVA, heparin and PVA/heparin hydrogels at their equilibrium swelling ..... 97

Figure 5.4 Comparison of hydrogel mass loss and their compositions ...... 99

Figure 5.5 Representative stress vs. strain curves for PVA and PVA/heparin

hydrogels...... 102

Figure 5.6 Illustration of the accessibility of ATIII binding sites in the PVA/heparin co-

hydrogels...... 105

Figure 5.7 Anticoagulant activity of PVA and PVA/heparin hydrogels ...... 106

Figure 5.8 Illustration of the accessibility heparin on PVA/heparin co-hydrogels for the

signalling of FGF-2 to its cell receptor ...... 109

Figure 5.9 BaF3 cell proliferation on PVA and PVA/heparin hydrogels ...... 110

Figure 5.10 The increase in BaF3 cell proliferation with increasing heparin content in

the PVA/heparin co-hydrogels ...... 111

Figure 6.1 Content of platelet extract (PE) as determined by ELISA ...... 124

Figure 6.2 Heparin degradation profiles at pH 5.1 (A) and 7.4 (B) ...... 126

Figure 6.3 Heparin degradation by platelet extract at pH 5.1 and 7.4 ...... 128

Figure 6.4 The effect of heating on the anticoagulation property of heparin ...... 130

Figure 6.5 The effect of heating on the FGF-2 signalling activity of heparin ...... 131

- xi -

List of Figures and Schemes

Figure 6.6 Changes in the anticoagulant activity of heparin, after an incubation period

with PE ...... 132

Figure 6.7 The effect of heparin molecular weight on the cellular signalling of

FGF-2 ...... 134

Figure 6.8 Changes in the FGF-2 cellular signalling activity of heparin, after an

incubation period with PE ...... 135

Figure 6.9 Anticoagulant activity of PVA/heparin 19/1 hydrogel supernatants ...... 138

Figure 6.10 FGF-2 signalling activity of PVA/heparin 19/1 hydrogel supernatants .... 140

Scheme 2.1 Crosslinking mechanisms for covalent biomolecule incorporation ...... 20

Scheme 2.2 Poly(vinyl alcohol) (PVA) ...... 26

Scheme 2.3 The predominant repeating disaccharides of heparan sulfate (HS) and

heparin ...... 33

Scheme 3.1 Synthesis of poly(vinyl alcohol) (PVA) macromer ...... 52

Scheme 3.2 Synthesis of heparin macromer ...... 62

Scheme 4.1 The specific heparin pentasaccharide sequence for ATIII binding ...... 69

Scheme 6.1 The β-glycosidic linkage on heparan sulfate that is cleaved by

heparanase...... 133

- xii -

Tables

List of Tables

Table 2.1. Strategies for producing vascularised tissue engineered constructs ...... 10

Table 3.1 Efficiency of PVA-methacrylate synthesis by reaction with glycidyl

methacrylate...... 52

Table 3.2 Properties of PVA hydrogels from GMA- and ICEMA- synthesised

macromers...... 56

Table 5.1 Equilibrium swelling, mass loss and tensile properties of PVA, heparin and

PVA/heparin hydrogels ...... 97

Table 5.2 Nominal and equilibrium compositions of PVA, heparin and PVA/heparin

hydrogels...... 101

Table 5.3 Tensile properties of PVA and PVA/heparin hydrogels ...... 103

Table 5.4 Average equilibrium heparin content in the PVA/heparin co-hydrogels ...... 107

Table 6.1 Molecular weight estimate of platelet extract (PE)-treated heparin ...... 129

- xiii -

Common Abbreviations and Terminology

Common Abbreviations and Terminology

aPTT activated partial thromboplastin time

DMMB 1,9-dimethylmethylene blue

FGF-2 fibroblast growth factor-2

GAG glycosaminoglycan

GMA glycidyl methacrylate

GPC gel permeation chromatography hep- or /hep heparin- or /heparin

HS heparan sulfate

HSPG heparan sulfate proteoglycan

ICEMA 2-isocyanatoethyl methacrylate

IL-3 interleukin-3

(-)MA (-)methacrylate

Macromer polymer chain modified with crosslinkable pendant groups

® MTS CellTiter 96 AQueous One Solution Reagent ([3-(4,5-dimethylthiazol-2-

yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium,

inner salt

MW molecular weight

NMR nuclear magnetic resonance

PE platelet extract

PVA poly(vinyl alcohol)

RU repeating unit

- xiv -

Common Abbreviations and Terminology

SEC size exclusion chromatography

Sol fraction soluble fraction of a hydrogel, consisting of polymer chains that are not

incorporated into the hydrogel network during the hydrogel-crosslinking

process

UV ultraviolet

- xv - Chapter 1

Introduction

ollowing trauma, the body regulates the regeneration of the injured tissue by Frecruiting cells responsible for the formation of connective tissue and blood vessels. In the case of organ failure or massive loss of tissue, organ transplant and tissue grafts are the most commonly used methods to replace the function of the damaged tissue. The limited number of organ donors has seen the increase in demand for an artificial system that is capable of mimicking the function of the damaged organ [1-4].

Tissue engineering has emerged to address this issue, with the focus of designing biomaterials to support the healing, regeneration, and replacement of function of the damaged tissue. It has been an ongoing challenge to engineer materials that express the inherent functionalities specific to their applications [5]. In order to serve their purpose, these biomaterials need to be able to interact with biological systems. The incorporation of cell-adhesion peptides such as fibronectin have been shown to encourage cell attachment and growth on biomaterials, however more sophisticated molecular cues are needed for the in situ regulation of growth factors and cells during wound healing. The natural extracellular matrix (ECM) contains such molecules, including the glycosaminoglycans (GAGs), and provides structural support for tissue regeneration.

- 1 -

Chapter 1 – Introduction

Hydrogels, the crosslinked form of hydrophilic polymers, have attracted interest for application in soft-tissue repair due to their high water content and elasticity, properties that resemble the ECM. Hydrogels from natural sources such as collagen offer the advantage of having an inherent biological recognition, however this can be hindered by reproducibility issues often faced with compounds extracted from biological tissues [6]. In recent studies, synthetic hydrogels have been coated, combined and covalently modified with molecular cues to provide the resulting scaffold with biofunctionality [5, 7-8]. A significant effort has been made to encourage vascularisation into scaffolds, as tissues cannot grow more than a few millimetres thick without proper channels for nutrients and waste transport [4].

The formation of blood vessels in vivo is mediated by growth factors such as fibroblast growth factors (FGFs) and vascular endothelial growth factors (VEGF).

Heparan sulfate (HS) proteoglycans, molecules that are widely distributed on cell surfaces and in the ECM, bind these angiogenic growth factors through their HS GAGs chains and have been found to regulate their activity. The binding of these factors to

HS, and to the structurally similar heparin, not only protects them from proteolytic degradation but also enhances interactions with their cell-surface receptors. For this reason, heparin/HS have been incorporated as co-factors in the encapsulation and delivery of growth factors from polymeric scaffolds. Heparin-crosslinked hydrogels have been analysed for the binding and controlled release of growth factors [8-10] and have been explored to support the growth of various cell types [11-12], however the specific function of the heparin on cell growth has not been well-characterised.

- 2 -

Chapter 1 – Introduction

Aims of Thesis

This thesis describes the incorporation of heparan sulfate as a bioactive molecule into a synthetic hydrogels, in order to facilitate hydrogel interactions with cells and protein following implantation. The overall objective of the current research was to investigate the formation of hydrogels derived from heparan sulfate, with poly(vinyl alcohol) (PVA) as structural support, and to examine the capability of the hydrogels to enhance growth factor signalling to cells, with a potential for applications in wound healing. Hydrogels were formed by the covalent crosslinking of PVA and heparin chains, using the photopolymerisation technique. The specific aims of this work were:

(1) To functionalise PVA and heparin with photopolymerisable pendant groups of

similar reactivity, which will enable the crosslinking of PVA and heparin;

(2) To investigate heparin bioactivity following the attachment of pendant groups. As

heparin activity is structure-dependent, it is important that the chemically-

modified heparin preserve its activity prior to incorporation to PVA;

(3) To characterise the structure and strength of the formed PVA and heparin co-

hydrogels, specifically, the effect of copolymerisation;

(4) To investigate the ability of the co-hydrogels to present growth factors to

surrounding cells; and

(5) To study the response of the co-hydrogels to enzymatic conditions, in order to

predict what might happen in vivo when the co-hydrogels are exposed to these

factors following implantation.

These specific aims were addressed in separate Chapters in the thesis. Literature on the area of tissue engineering, identifying the need for scaffolds to mimic the properties of natural extracellular matrix for tissue regeneration, was reviewed in

- 3 -

Chapter 1 – Introduction

Chapter 2. Subsequent Chapters focused on the specific aims of the thesis, along with more detailed review of relevant literature.

In Chapter 3, the methods used for the synthesis of PVA and heparin macromers were discussed. Two synthesis routes were considered and the cell compatibility of the functional groups was investigated.

Chapter 4 focused on the structure and bioactivity of the heparin, following methacrylate modification. The anticoagulant and growth factor-signalling activities of methacrylated heparin were assessed and compared to those of non-modified heparin.

In Chapter 5, the formation of PVA and heparin co-hydrogels was investigated.

The effect of adding heparin on the structure and stability of the resulting co-hydrogels were characterised, while the ability of the co-hydrogels to present their growth factor signalling activity to surrounding cells was confirmed.

Finally Chapter 6 investigated the degradation of heparin by its natural enzyme, heparanase, extracted from human platelets. The specific activity of the heparanase on heparin was investigated, before the enzyme was applied to the hydrogels in order to simulate the release of heparin in vivo. The activity of heparin has been known to be structure-specific, therefore the effect of size on heparin activity, incorporation and modification were also discussed.

Conclusions of the thesis and recommendations for future work are summarised in Chapter 7.

- 4 -

Chapter 2

Literature Review

xtensive research has been done in developing various biomaterials for tissue Eregeneration. Progress in polymer fabrication technologies, in particular, provided more options in the way biomaterials could be made. Different polymers are often combined as composites to combine their properties and potentially tailor them to meet specific criteria. The incorporation of glycosaminoglycans, the biopolymers present in the extracellular matrix and on cell surfaces, into TE constructs has been increasingly demonstrated due to their ability to promote cell interactions and growth factor signalling. This chapter reviews existing work in biomaterials development for use as tissue engineered constructs, and addresses aspects that are yet to be explored, which became the platform behind the research in this present work. The methods used for the combination of synthetic and biologically-sourced polymers are discussed, along with the polymers used in this study.

- 5 -

Chapter 2 – Literature Review

2.1 TISSUE ENGINEERED CONSTRUCTS

The field of tissue engineering has grown immensely since its early conception, moving closer towards clinical applications in regenerative medicine. This interdisciplinary field combines the principles of engineering and life sciences to create functional substitutes that can be used to facilitate the restoration of the structure and function of a damaged tissue [1-2, 13]. The general approach in tissue engineering involves transplanting the appropriate cells onto a biomaterial scaffold, culturing the cells to encourage attachment and growth, before implanting the cell-scaffold unit into the desired site in the patient’s body. The potential impact of tissue engineering in the future is extensive; engineered tissue could reduce the need for organ replacement and may ultimately eliminate the need for organ transplants [4, 13].

The design criteria of biomaterial scaffolds are influenced by the intended application, and several key requirements that must be met include application-specific biocompatibility, the ability to support selective cell adhesion, as well as structural and mechanical properties matching those of the tissue to be regenerated [14]. Different strategies have been applied to engineer different tissue types. For example, in supporting the regeneration of a soft tissue, a scaffold should maintain its structural integrity during the initial tissue formation, promote vascularisation of the developing tissue, and subsequently degrade to be replaced by new tissue. A rapid and high level of vascularisation is required for the survival of most cell types following transplantation.

In addition, the ability of the scaffold to induce the migration, growth and differentiation of cells, and to finally integrate with the native tissue, is critical for the functional repair of the tissue [15]. The choice of materials and scaffold forming mechanism is crucial in fulfilling the physical property, mass transport and biological interaction requirements of the scaffold [16].

- 6 -

Chapter 2 – Literature Review

2.1.1 Molecular Interaction

The development of biomaterial scaffolds has been increasingly aimed at improving their properties to better mimic the natural extracellular matrix (ECM), as the

ECM and its components play a central role in wound healing and tissue regeneration

[5-6]. The natural ECM is a highly hydrated network that surrounds cells and comprises molecular signals. Insoluble structural macromolecules such as collagen, fibrin and elastin, provide the necessary architecture and scaffold for the cells, while complex polysaccharides or glycosaminoglycans (GAG) keep the structure hydrated. The ECM is also made up of transient components such as growth factors, diffusible molecules that can be released or activated upon appropriate stimuli [17]. Cells differentiate, proliferate, migrate or perform other specific functions in response to the bi-directional molecular interactions with these ECM compounds [7, 18]. Tissue regeneration in vivo involves these molecular signals, both from the wound site and from healthy surrounding tissues. As an analogue to the ECM, a scaffold should be able to selectively interact with the adhesion domains and growth factor receptors expressed by target cells that are required to repair the damaged tissue.

Both biologically derived and synthetic polymers have been extensively researched for use as biomaterial scaffolds. Polymers from natural sources have the advantage of being able to closely simulate the native cellular conditions, including the presentation of receptor binding ligands, however there might be a variation in composition between batches and also lack of physical strength [6, 19]. Synthetic biomaterials, on the other hand, allow greater control in tailoring the material properties such as mechanical stability. An obvious choice for biologically sourced scaffold material is collagen, which is a major structural component of the natural ECM, found in all major tissues that require both strength and flexibility. Type I collagen accounts

- 7 -

Chapter 2 – Literature Review for more than 90% of all fibrous proteins, and due to the abundance in source, it has been used extensively in the development of biomedical materials, many of which had been approved for commercial use [20]. The primary structure of collagen I also presents attachment sites for cells [21]. Collagen has been frequently crosslinked to form hydrogels to maintain their mechanical properties and stability during implantation

[20, 22].

As for synthetic materials, resorbable polyesters of lactic acid and glycolic acid, and their copolymer (PLGA), are the most widely used scaffold materials in tissue engineering, and have been used for over twenty years in a variety of medical implants, including sutures and bone fixation devices. They are relatively low in toxicity, with an extensive US Food and Drug Administration (FDA) approval history. PLGA has been used in the controlled delivery of growth factors, which were incorporated into the polymer either by simple mixing before processing [23-24] or by pre-encapsulation in

PLGA microspheres [25-27]. The rate of growth factor delivery is typically coupled to the polymer degradation rate, which, along with the mechanical properties of the copolymer, is regulated by the ratio of lactide to glycolide and the degree of polymerisation [23, 28]. These polyesters are hydrophobic but degrade by hydrolysis into lactic and glycolic acids, which are eventually resorbed. However, these acidic degradation products have been reported to cause adverse tissue reactions [29].

Hydrophilic polymers, by their chemical nature, show promise for use in biological applications and have been crosslinked to provide structure and strength [30].

Hydrogels, the insoluble network of hydrophilic polymers, can be formed by physical or chemical crosslinks [31]. The type and degree of crosslinking among the polymer chains ultimately dictate the properties of the hydrogels. Hydrogels have tissue-like properties due to their high swellability, making them an attractive alternative scaffold

- 8 -

Chapter 2 – Literature Review material to PLGA. The relatively mild processing conditions of hydrogels may enable in situ formation, with reported studies including the development of injectable hydrogel systems and transdermal photopolymerisation [6, 16]. Synthetic hydrogels lack the molecular recognition possessed by their biologically-sourced counterpart such as collagen. Hybrid materials have been developed to achieve this molecular recognition, preserving the bulk properties of synthetic materials while using biopolymers that resemble proteins in vivo at the interface of the scaffolds and cells. In recent studies, natural materials have been combined with synthetic polymers for structural support, while providing the scaffold with biofunctionality [5, 7, 32].

2.1.2 Vascularisation of Constructs

Hydrogels, as well as porous scaffolds, have been investigated for use as cell delivery matrices since both types of structure allow for nutrient diffusion into the network. Even so, the delivery of nutrients and oxygen by diffusion alone is inadequate for tissues that are more than a few hundred micrometres in size. The lack of perfusion causes cells in the centre of these constructs to undergo apoptosis, leading to the necrosis of surrounding tissue [13, 33]. Creating a vascular network capable of facilitating nutrients and waste transport within the scaffold has proven to be a challenge, and the absence of this vascular network has been a major limiting factor on the size of bioengineered tissue in vitro [34].

Various approaches have been used in an attempt to create new vasculature in TE constructs. Fundamental design requirements are that the scaffold has the appropriate soluble and insoluble biological signals, adequate pore/mesh size, combined with degradability to allow migration and proliferation of endothelial cells or vascular progenitor cells. A challenge associated with using biosynthetic hydrogels in this

- 9 -

Chapter 2 – Literature Review context is that unless they are degradable, cell migration and tube formation within the gels is unlikely. Many studies have reported tube formation in collagen gels [35-36], but formation of vascular structures in synthetic hydrogel constructs is rare. Given that the above essential design requirements are fulfilled, vascularisation of constructs may be produced via a range of approaches which are summarised in Table 2.1. These include loading and subsequent release of angiogenic growth factors for both in vitro and in vivo vascularisation, in vitro seeding of hydrogels with endothelial and progenitor cells, and developing pre-vascularised constructs in vivo.

Table 2.1. Strategies for producing vascularised tissue engineered constructs. Strategy Description References 1. Angiogenic growth Growth factors (GF) incorporated into 3D factors delivery matrices for delivery. Rate of GF delivery controlled by: a. Pre-encapsulation of GF for sustained release [37-39] and to protect GF from denaturation b. Use of HS/heparin crosslinked matrices to [8, 40-41] increase affinity of GF to the matrices for sustained release c. Controlling the release of heparin, as co-factor [42-43] in the delivery of GF 2. In vitro seeding Formation of vascular-like structures in hydrogel [35-36, 44] using endothelial or matrices derived from collagen or basement progenitor cells membrane proteins (Matrigel) Seeding of cells on matrices in vitro followed by [24, 45] immediate transplantation; using the matrices and host as bioreactors for vascular formation 3. In vivo Implanted GF-loaded matrices encouraged vascularisation vascularisation in tissues surrounding the implant a. Multiple growth factor delivery from PLGA [46] matrices for vessel maturation b. Use of heparin-crosslinked hydrogels (natural [8, 10, 47- and synthetic) to enhance GF activity in 48] promoting neovascularisation HS-crosslinked and GF-loaded collagen matrices encouraged sustained vascularisation throughout [49] the hydrogel structure.

- 10 -

Chapter 2 – Literature Review

The discovery that growth factors alone can stimulate development of new blood vessels promoted broad experimentation using factors such as vascular endothelial growth factor (VEGF), platelet derived growth factor (PDGF), and fibroblast growth factor (FGF). Pharmacological studies have been performed extensively on therapeutical administration of these angiogenic factors to induce vessel formation and improve tissue perfusion [33, 50]. However, high doses or repeat injections are required to achieve significant response because of the short protein half lives and rapid diffusion out of target tissue, causing concerns for potential side effects at distant sites.

To control a localised and sustained delivery of growth factors to a target site, scaffold materials have been designed to either incorporate or interact with these factors. Growth factors regulate different stages of vascularisation, and thus degradable matrices such as PLGA have been used to deliver multiple growth factors in succession.

In these matrices, initial growth factor delivery was controlled by the degradation rate of the bulk structure, while delayed release was achieved by pre-encapsulating another factor prior to scaffold processing [46, 51]. When implanted subcutaneously in rats, the dual-release scaffold induced higher vessel density and the growth of more mature vessels compared to the response from scaffolds containing either factors alone [46].

Most of the work in developing hydrogels for growth factor delivery has focused on improving the incorporation efficiency and controlling the release of these factors

[52]. The angiogenic potential of these growth factor-carrying matrices has mostly been evaluated in terms of their capability to support endothelial cell growth and to encourage vascularisation of surrounding tissue. Biologically-sourced hydrogel components such as alginate and heparin-sepharose beads have been used in the incorporation of growth factors into synthetic matrices, due to their ability to interact ionically or specifically with the factors [53]. These interactions resulted in more

- 11 -

Chapter 2 – Literature Review sustained release of the growth factors from the matrix, while at the same time protected the factors from denaturation. When calcium-crosslinked alginate beads were used to pre-encapsulate VEGF prior to incorporation in PLGA matrices, the effect was not only the improvement in incorporation efficiency but also the observed increase in VEGF activity in stimulating endothelial growth in vitro [37-38].

The affinity of growth factors to GAG molecules, the natural components of the

ECM, have led to the development of growth factor release vehicles built of GAG chains, using synthetic polymers as hydrophilic crosslinkers [54-55]. Heparin, in particular, provides an analogue of the heparan sulfate, molecules that have been known to regulate blood vessel formation in vivo [56]. Heparinised collagen matrices have been shown to support the attachment and proliferation of human umbilical vein endothelial cells (HUVEC) in vitro [40, 57], making them potential materials for EC seeding for synthetic vascular grafts. When HS-modified collagen matrices were implanted, the presence of HS alone was shown to induce transient vascularisation at the periphery of the matrices, while pre-loading the matrix with basic fibroblast growth factors (FGF-2) was shown to encourage capillary infiltrations [49]. The structures remained vascularised during a 10-week implantation period. Similar matrices containing VEGF were also shown to increase the capillary formation in the chorioallantoic membrane of chicken embryo, and increase vascularisation of surrounding tissue after a 14-day implantation in rats [41, 47].

- 12 -

Chapter 2 – Literature Review

2.2 BIOSYNTHETIC MATERIALS

The use of synthetic materials as biomaterial scaffolds is appealing due to the ability to control their properties and reproducibility. While synthetic hydrogels such as poly(vinyl alcohol) (PVA), poly(ethylene oxide) (PEO), and poly(ethylene glycol)

(PEG) are hydrophilic and have been found to be low in toxicity, they are relatively resistant to protein adsorption and cell attachment [58]. Hydrophobic scaffolds have been shown to encourage better protein deposition. The hydrophobic poly(lactic acid)

(PLA) has been grafted as side chains onto PVA backbone and shown to improve the adhesion of valve interstitial cells to the hydrogels [58].

Fibronectin (Fn) is a well-known cell adhesion molecule, with a cell-binding domain consisting of a tripeptide sequence of Arg-Gly-Asp (RGD) [59]. Fn, as well as its RGD derivative, has been covalently incorporated into polymers to create cell- binding surfaces, and found to support the attachment and spread of many cell types

[59-61]. It has been reported however that the presence of these cell-adhesion molecules reduced the production of ECM by the attached cells [62]. While the incorporation of adhesion peptides facilitiate initial cell attachment, further cellular processes such as proliferation and vessel formation requires more sophisticated molecular cues, capable of cellular signalling of growth factors and cytokines. The inclusion of growth factor- binding GAG molecules is scaffolds have been demonstrated to control the incorporation and release of growth factors from the scaffolds [10, 41, 63]. The following section reviews various growth factor and GAG incorporation techniques into collagen and synthetic scaffolds. Physical and covalent incorporations of these signalling molecules are discussed, as well as the effect of GAG on the presentation, release or signalling of the growth factors by the scaffolds.

- 13 -

Chapter 2 – Literature Review

2.2.1 Physical Incorporation of Signalling Molecules into Scaffolds

Growth factors are rapidly degraded in vivo following injection or ingestion. A study has shown that intravenously administered FGF-2 was rapidly cleared from the blood, predominantly by the liver, and could only be maintained at a constant level in blood with continuous intravenous infusion [53, 64]. To administer growth factors locally over a long period of time and in their active form, research had been done on the use of degradable polymeric matrices as sustained delivery vehicles. In this application, the degradability of a scaffold thus becomes an important consideration, as it would control the growth factor release.

Initial attempts at incorporating growth factors into degradable matrices have involved the use of hydrophobic polymers using the solvent casting technique [53]. This technique involved mixing the growth factors and the polymer in an organic solvent, casting into a mould, quenching, before applying vacuum to remove the solvent [65-66].

The high temperature drop and organic solvents used in this process have presented limitations to maintain the biological activity of the released factors. Mooney et al. developed a gas foaming/ particulate leaching technique which avoids the use of those harsh processing conditions, to create a PLGA scaffold with an interconnecting porous structure [67-69]. In the study, vascular endothelial growth factors (VEGF) was mixed with the PLGA prior to processing. The released VEGF was found to maintain 90% bioactivity, however the incorporation efficiency of the VEGF into the scaffolds was less than 30% [23].

A matrix capable of delivering multiple growth factors has also been developed, based on the fact that tissue regeneration is governed by the synchronised work of a number of growth factors. The synthesis of the matrix combined the principle of micro- encapsulation with the gas foaming/ particulate leaching technique [51]. A growth

- 14 -

Chapter 2 – Literature Review factor (PDGF) was pre-encapsulated in PLGA microspheres, while another factor

(VEGF) was added to the mixture of PLGA particles and the microspheres during the scaffold formation process [46]. The system allowed a two-phase delivery of VEGF and

PDGF; the first, to stimulate endothelial vessel formation, and the second, to recruit smooth muscle cells that will stabilise the newly formed vessels. By first releasing

VEGF, endothelial vessel formation was stimulated, and the subsequent release of

PDGF initiated the recruitment of smooth muscle cells to stabilise the newly formed vessels. After 4 weeks subcutaneous implantation in rats, the dual-release scaffold was shown to induce higher vessel density and growth of more mature vessels compared to the responses from scaffolds containing either factors alone. Human microvascular endothelial cells (HMVEC) transplanted on VEGF-containing PLGA matrices were able to form human-derived vessels within 7 days of implantation in mice; the combination of transplanted EC and VEGF resulted in a significant increase in density of vessels within the scaffold [24].

The incorporation efficiency of VEGF into PLGA has been shown to improve when VEGF was pre-encapsulated in calcium-crosslinked alginate hydrogel beads prior to polymer processing [37-38], or when alginate was crosslinked into the matrix [24].

There was a noted increase in the activity of growth factors released from the alginate beads, which might have acted as a protective shield from surrounding environment.

The incorporation efficiency of the growth factors into the calcium alginate microspheres has been shown to increase further when the factors were incubated with heparin-sepharose beads prior to encapsulation [37, 53]. Heparin binds various growth factors and has been shown to protect them from heat and acid inactivation. Heparin was also shown to slow the rate of FGF-2 clearance from the body when administered intravenously together with the growth factor [64]. The binding of growth factors to

- 15 -

Chapter 2 – Literature Review heparinised beads has been shown to provide a more sustained release, while at the same time protected these factors against denaturation. The mitogenic potential of FGF-

2 released from this system, assayed by measuring its ability to stimulate DNA synthesis from cultured mammalian cells, had been found to be preserved [53]. A study on VEGF encapsulated in these alginate beads showed that the sustained release of

VEGF enhanced the stimulation of endothelial cell growth in vitro compared to when the same amount of VEGF was introduced directly to the cell culture [37].

More recently, heparin has been investigated as a co-release agent for growth factors. The release kinetics of heparin has been studied from composite alginate/hydroxyapatite microspheres, as a parameter that could be used to control the delivery of bound growth factors. The inclusion of hydroxyapatite was found to increase the encapsulation efficiency but also increased the rate of heparin release [43]. The co- incorporation of heparin and fibroblast growth factors (FGF) into a chitosan hydrogels has been shown to have an incorporation efficiency of 75%, and when implanted in a mouse model chitosan hydrogels that contained heparin induced higher neovascularisation that those containing FGF only [42]. The general strategy for the physical incorporation of growth factors, with or without co-factors such as alginate or heparin, is depicted in Figure 2.1.

GF2

Polymer + + solution GF1

microspheres Biodegradable scaffold

Figure 2.1 The general strategy for growth factor delivery from degradable matrices. Growth factors are either mixed directly into polymer solution (GF1) or pre-encapsulated in microspheres (GF2). GF2 can be co-delivered with GAG molecules. The order of release will be GF1, as a result of degradation of bulk scaffold, followed by GF2 from the slower degrading microspheres.

- 16 -

Chapter 2 – Literature Review

2.2.2 Covalent Incorporation of Signalling Molecules into Scaffolds

Composites of collagen and GAG have long been investigated for use as artificial tissues, in particular as skin grafts, mainly because they are natural components of the

ECM that can be degraded by extracellular enzymes or hydrolytic scission into non- toxic products [19, 70-71]. The degradation rate of collagen can be controlled by crosslinking, however heavily crosslinked collagen is a stiff and brittle material. The addition of GAGs to collagen has been shown to yield composite matrices that are more resistant to collagenase degradation, providing a way to control the degradation without excessive crosslinking, and that had higher elasticity [19, 72]. Several different chemistries have been employed to covalently link GAG to collagen and synthetic polymers, and also to crosslink the GAG to form hydrogel scaffolds. Carboxyl and hydroxyl groups on the polysaccharide backbone have generally been the targeted sites for the attachment of chemical crosslinkers [63, 73-74].

It is only in more recent studies that the biological functionality of GAG in the scaffolds was investigated closely. Growth factors must undergo various processes in order for them to be incorporated into scaffolds for controlled delivery, which might affect or even denature its functionality. GAGs have been known to regulate these proteins in various biological processes in vivo, are more resistant to temperature changes and have been shown to retain their activity following chemical modifications

[75-76]. Therefore, by covalently incorporating the GAG molecules to a scaffold, continuous presentation of the growth factors could be expected, for longer time periods than the time it takes for them to denature in vivo.

Figure 2.2 illustrates the strategy for covalently linking GAGs into a scaffold, depicting the capability of covalently-bound GAG molecules to retain growth factors within or on the surface of the scaffold by providing specific binding sites for the

- 17 -

Chapter 2 – Literature Review growth factors to bind to. The following section will discuss the synthesis methods that have been used to modify GAG and similar polysaccharides for covalent incorporation.

*

growth factors GAG-incorporated scaffold

specific, high affinity binding

GAG

scaffold covalent linkage

Figure 2.2 Covalently bound GAGs facilitate growth factor binding onto scaffold. Growth factors are introduced to the scaffold (*) by injection or incubation and retained by specific binding to GAGs.

2.2.2.1 EDC/NHS chemistry

Crosslinking of collagen has been done to increase its resistance to proteolytic degradation and to mask immunogenicity to implanted collagen-rich xenograft materials. Different crosslinking methods to form collagen hydrogels, matrices or films have been reported. Among the first studies on the formulation of a collagen-GAG matrix was the work by Burke et al., which demonstrated the applicability of a collagen-chondroitin sulfate matrix as an artificial skin for the treatment of burn injury

[70]. Composites of collagen and GAG have been prepared by precipitating both components at low pH and crosslinking the ionic molecules by a dehydrothermal (DHT) treatment, or by using glutaraldehyde or cyanamide as crosslinkers [22, 72]. Unreacted glutaraldehyde (GA) remaining in the cosslinked matrix had been shown to be locally toxic to the surrounding tissue [77-79]. When proper extraction procedures to remove these toxic residues were applied, GA-crosslinked collagen showed low cell growth

- 18 -

Chapter 2 – Literature Review inhibition in vitro. However, cytotoxicity was still apparent when the matrix was subcutaneously implanted in rats, especially as GA-containing fragments were enzymatically released from the collagen matrix [80].

Olde Damink et al. explored the use of water soluble 1-ethyl-3-(3-dimethyl aminopropyl) carbodiimide (EDC) and N-hydroxysuccinimide (NHS) to crosslink collagen [22]. The resulting collagen matrix was found to be non-cytotoxic in vitro, and able to support the formation of new collagen in vivo while only inducing cellular reactions that were similarly found in normal wound healing [80]. The group of van

Kuppevelt further adapted this method for crosslinking chondroitin sulfate (CS) with collagen [81]. CS was incorporated throughout the matrix whereas when using the above method (DHT), CS was only located on the exterior of the matrix. The group demonstrated the applicability of this method to other GAGs including dermatan sulfate, heparin and heparan sulfate [63]. The degree of crosslinking, the amount of

GAG incorporated, and biodegradability of the collagen-GAG matrix could be modified by varying the respective molar ratios of EDC, NHS, the carboxyl groups of the GAGs and the amine groups of collagen. The typical EDC/NHS synthesis method is shown in

Scheme 2.1(a).

- 19 -

Chapter 2 – Literature Review

R NCN R R R O 1 2 O 1 NCN 2 O O O O O EDC EDC SH O GAG* R + n Collagen* NH GAG* OH 3 O O GAG* GAG* + 2 OH OH O N PEG diacrylate O SH GAG* SH O OH DTT NOH O R O O or 3 O S SH O GAG* NH Collagen* OH 2NH NHS + N cysteamine O O S R GAG* O 3 O n O (a) (b)

Chitosan* O O O O O NH O O O O O O OH OH OH UV light O methacrylic anhydride n m p NH NH 2 NH GAG* O N2 GAG* OH O O + or + Synthetic polymer* O OH OH N Chitosan* O OH O OH UV light N NH2 O 3 Chitosan* NH photoinitiator glycidyl methacrylate N Chitosan* NH O O R N 2 O N Chitosan* GAG* O

Chitosan* NH NH Chitosan* N N O O Synthetic polymer* R (c) (d) 1

Scheme 2.1 Crosslinking mechanisms for covalent biomolecule incorporation, (a) EDC/NHS chemistry, (b) thiol modification, (c) lactose modification of chitosan, and (d) methacrylate group attachment. *the compound possess more than one modifiable pendant groups.

- 20 -

Chapter 2 – Literature Review

The bioavailability of GAGs attached to the collagen matrices has been assessed directly by testing their enzyme degradability and their ability to bind specific antibodies and growth factors [57, 81]. Heparinised collagen matrices formed by the

EDC/NHS method have been studied as a potential material for synthetic vascular grafts

[40, 57]. Human umbilical vein endothelial cells (HUVEC) were seeded on matrices that were pre-loaded with fibroblast growth factor-2 (FGF-2). Steffens et al. tested the angiogenic potential of a similar heparin-collagen matrix by performing a chorioallantoic assay on the membrane of chicken embryos and by implanting them subcutaneously in rats [41].

The development of GAG hydrogels, using hydrophilic synthetic polymers as crosslinkers, has also been investigated. Tae et al. investigated the formation of heparin hydrogels for the sustained release of VEGF [10]. Heparin was modified with hydrazide functional groups, using EDC as an intermediate reagent and by adding an excess of adipic dihydrazide (ADH). The heparin gel was formed by reacting the heparin-ADH compounds with the NHS group on the poly(ethylene glycol)-bis-butanoic acid (SBA-

PEG-SBA) chain. A 13 wt% gel was formed with a 2:1 Hep-ADH to SBA-PEG-SBA ratio, and VEGF was injected into the gel. After a 2-week subcutaneous implantation in mice, new blood vessels was observed in the tissue surrounding the VEGF-loaded gel while almost no vascularisation was observed around the control heparin gel. No cells or vascular structures were found to infiltrate the gel.

2.2.2.2 Thiol modification

Hydrogel formation by a disulfide crosslinking method (Scheme 2.1 (b)) was developed by the Prestwich research group [82] and has been used to form GAG hydrogel film for wound healing applications [8, 55, 83]. Low molecular weight

- 21 -

Chapter 2 – Literature Review hyaluronic acid (HA) has been modified by first attaching hydrazide molecules using the carbodiimide chemistry, then reducing the disulfide bonds on the hydrazide molecules with dithiothreitol (DTT). The resulting thiol pendant groups were oxidised to re-form the disulfide linkages and create HA hydrogel films. The thiol to disulfide conversion was reversible. Higher disulfide content was obtained when, in addition air oxidation, the film was further oxidised with 0.3% H2O2. Although mouse fibroblasts

(L929) showed increased in proliferation for up to 3 days encapsulated in the hydrogels, due to the non-sulfated nature of HA, the hydrogel was hydrophilic and did not favour the attachment cells, which possess anionic surfaces [84]. Crosslinking HA with gelatin by the same method demonstrated a degradable in vitro environment for cell growth

[85]. Using this chemistry, the covalent incorporation of drugs, such as mitomycin C

[86], has also been also demonstrated.

As with any types of surgery, minimally invasive procedures are desired under permitting circumstances. An injectable, in situ forming hydrogel system was later developed by Cai et al., using three types of GAGs [8]. HA, chondroitin sulfate (CS), and heparin were chemically modified with thiol pendant groups, and crosslinked using poly(ethylene glycol) diacrylate (PEGDA) to form hydrogels (Scheme 2.1(b)). The hydrogels were <3 wt% in composition and formed within 10 minutes. The released

FGF-2 maintained 55% of its activity over a period of 28 days [8, 87].

The antithrombogenicity of thiol-modified heparin that are immobilised on polymer surfaces has been investigated in the microfluidics studies of blood or plasma samples [88]. Heparin molecules were grafted onto a polymeric amine carrier chain

(PAV) by subsequent reactions with a heterobifunctional coupling agent, N- succinimidyl 3-(2-pyridoldithio) propionate, and DTT [89-90]. Up to 70 mol of heparin could be attached to 1 mol of 50 kDa PAV. The heparin was shown to suppress fibrin

- 22 -

Chapter 2 – Literature Review formation when coated on poly(dimethylsiloxane) (PDMS) microchannels [88] and increased the biocompatibility (in terms of antithrombogenicity) of extracorporeal devices during cardiopulmonary bypass [91-92].

2.2.2.3 Lactose-modified chitosan

Chitosan, a natural polysaccharides made of 1,4 β-linked N-acetyl-D-glucosamine units, has been investigated for use in wound healing applications. The preparation of photocrosslinkable chitosan was first investigated by Ono, et al. [93]. The amino groups on the chitosan backbone were substituted with azide (Az) and lactose (LA) groups using a two-step condensation reaction (Scheme 2.1(c)). The resulting chitosan derivative (Az-CH-LA) was photo-crosslinked. The azide groups released N2 gas upon

UV-irradiation and converted into nitrene groups, which reacted with each other or with the amino groups to form crosslinks.

Functionalising the chitosan with 2% lactose and 2.5% azide improved its solubility in physiological pH. The binding strength of the chitosan hydrogels was found to increase with macromer concentration, with a 50 mg/mL hydrogel having similar strength to commercial fibrin glue [93]. The azide groups were thought to covalently link with tissue proteins, therefore the site of rupture of the chitosan gels was not at the hydrogel-tissue interface, but within the chitosan network itself.

When FGF-1 and FGF-2 incorporated chitosan hydrogels were implanted into the back of a mouse, neovascularisation was observed around the site of implantation

[42]. Most of the growth factors (~75%) remained in the chitosan hydrogels at equilibrium, indicating that the growth factor release in vivo was done by degradation of the chitosan network. As observed in other studies, vascularisation was improved when heparin was co-incorporated with the growth factors.

- 23 -

Chapter 2 – Literature Review

- An injectable chitosan/non-anticoagulant (IO4 ) heparin hydrogel has also been developed using similar methods [48]. The chitosan was modified with lactose moieties, while the non-antiocagulant heparin was prepared by oxidation with periodate. Gelation occurred by ionic interactions of the modified materials (40:1 CH-LA:IO4-heparin, 20 mg/mL CH-LA), and >80% of heparin remained in the hydrogel at equilibrium.

Although the heparin had been modified to minimise its antithrombotic capacity, its

FGF-2 activation property remained [94-95] and neovascularisation was induced near the injected site of the hydrogel in the mouse model.

2.2.2.4 Methacrylate modification

Photo-initiated polymerisation offers a convenient way to form hydrogels of different sizes and shapes. Like chitosan, GAG molecules have been modified with photocrosslinkable pendant groups to enable formation of hydrogels, both on their own and copolymerised with other polymers. However, the insolubility of GAGs in organic solvents limits the options for their modification. The most popular pendant groups reported for attachment to GAGs have been the methacrylate groups (Scheme 2.1(d)); the methods and the methacrylate precursors used are discussed below.

The coupling of methacrylate pendant groups to dextran, a bacterial polysaccharide, had been done by reaction with glycidyl methacrylate (GMA) in dimethyl sulfoxide (DMSO) [96]. Li et al. adapted this reaction mechanism for use in a heterogeneous-phase condition to functionalise CS, a main component of native cartilage [74]. The resulting CS macromers were decorated with methacrylate (MA) groups and pure CS hydrogels could be formed from 10 mol% MA-subsituted macromers under 8 mW/cm2 UV for 10 min. CS-MA had also been prepared by reaction with methacrylic anhydride [97]. CS has been copolymerised with PEG [98]

- 24 -

Chapter 2 – Literature Review and PVA [99] to form hydrogels and were shown to support the growth of encapsulated chondrocytes.

Heparin-MA has been synthesised mostly by reaction with methacrylic anhydride, and copolymerised with PEG dimethacrylate to form a delivery vehicle for FGF-2 [9].

Heparin was shown to retain its ability to bind proteins when copolymerised with PEG, and the resulting hydrogel was found to support adhesion and osteogenic differentiation of human mesenchymal stem cells (hMSC) [9, 75]. Due to the multi-binding and multi- signalling properties of heparin, extended applications of these heparin-modified PEG hydrogels have included the myofibroblast activation of valvular interstitial cells (VICs)

[11]. The copolymer gels have also been used as a fluvastatin-releasing matrix, to stimulate the production of bone morphogenic protein 2 (BMP2) by hMSC – a process which further support the differentiation of the hMSCs [12].

In this section, the formation of heparin-carrying matrices for growth factor presentation has been discussed. The binding and signalling of growth factors by heparin in solution has been intensively characterised [100-101]. The cell proliferation and neovascularisation effects of collagen and synthetic scaffolds have been shown to increase when heparin was incorporated into the scaffolds. However the specific growth factor signalling property of the hydrogel-bound heparin has not been widely investigated. This present work will attempt to address this area, using PVA as the supporting structure of heparin.

- 25 -

Chapter 2 – Literature Review

2.3 POLY(VINYL ALCOHOL) HYDROGELS

PVA hydrogels are formed by the crosslinking of linear PVA polymer chains to create an insoluble, swellable network. The monomer vinyl alcohol does not exist in a stable form, therefore PVA is synthesised by the hydrolysis of poly(vinyl acetate) [102], resulting in multiple pendant hydroxyl groups along the polymer backbone (Scheme

2.2). These pendant hydroxyls are highly reactive and can be readily substituted using various reaction mechanisms [103]. The degree of hydrolysis, that is, the percentage of acetate groups that have been hydrolysed, affects the chemical properties of the PVA, as well as its solubility and crystallinity [31].

m n O O OH

Scheme 2.2 Poly(vinyl alcohol) (PVA). The hydroxyl group (n) results from the hydrolysis of the acetate group (m).

2.3.1 Structure and Properties

PVA has been used in its linear as well as crosslinked forms. Introducing crosslinks is a way to increase the strength of PVA polymer, as linear polymer systems are often less durable and have lower mechanical strength [104]. The method of polymer crosslinking is a major factor that influences the final properties of the resulting network. Physically crosslinked hydrogels are formed by introducing crystalline regions that behave as crosslinks (Figure 2.3, left), and a freeze/thaw method for PVA crosslinking has been extensively demonstrated by Peppas and coworkers [2,

31]. The advantage of this method is that crosslinks are formed without the addition of chemical reagents. Physical crosslinking is typically reversible, which can be desirable

- 26 -

Chapter 2 – Literature Review in applications such as drug delivery; however the resulting crosslinks are likely to be less stable than covalent crosslinks formed in chemical reactions. For this reason, chemical crosslinking is widely accepted as the most efficient way to create durable and mechanically stable hydrogels.

functional group crystalline region

Figure 2.3 Formation of PVA hydrogels by physical and covalent crosslinking. Left, physical crosslinks are created by the formation of crystalline regions. Right, covalent crosslinks are created by the polymerisation of the pendant functional groups, indicated here by the formation of kinetic chains (---).

Covalent crosslinks can be formed in chemical reactions by the use of difunctional crosslinking agents, most commonly aldehydes, in the presence catalysts such as sulfuric acid, acetic acid, or methanol [103, 105]. Crosslinking occurs by the random and homogeneous formation of stable acetal bridges linking the hydroxyl groups, however, toxic residue of the crosslinking agents are often inevitably present in the resulting hydrogels [78-79]. The removal procedure of the residue is often time- consuming, but without which the hydrogels are unacceptable for use in medical applications.

A more recent method of chemical crosslinking addresses the need for a method that allows the formation of hydrogels in conditions that are relatively compatible with cultured cells and target applications. Photo-initiated radical polymerisation has been shown to allow a rapid hydrogel formation in the presence of non-toxic and water

- 27 -

Chapter 2 – Literature Review soluble photoinitiator [106-107]. PVA chains are first modified by attaching vinyl- carrying functional groups via a substitution reaction with the hydroxyl pendant groups on PVA backbone. The abundance of these hydroxyl groups gave PVA the flexibility of being modified with multiple functional groups. Upon irradiation, photoinitiator molecules form radicals that will attack the pendant functional groups, starting the formation of a kinetic chain that link the functional groups together (Figure 2.3, right).

PVA has been modified with non-degradable functional groups such as (meth)acrylates and acrylamides [108], as well as with hydrolysable ester-carrying groups [106]. The addition of hydrolysable crosslinker groups render the hydrogel biodegradable, and has been used in the formation of scaffolds for chondrocytes delivery [30]. The functionalisation of PVA with (meth)acrylate pendant groups have been widely reported, by reactions with (meth)acrylic acid [108], glycidyl (meth)acrylate [106, 109-

110], and 2-isocyanatoethyl methacrylate [99].

The swelling and degradation profiles of the hydrogel have been shown to be tailorable by altering the amount and type of functional groups on the macromer, the macromer molecular weight, and the concentration of macromer prior to polymerisation

[111]. The change in swelling capacity by varying initial macromer concentration has been shown to affect drug permeability into PVA hydrogels [52]. The crosslinking density within a gel influences its water content and consequently its mechanical strength. The increase in crosslinking has also been shown to increase both compressive and tensile moduli but reduced the amount of water imbibed in the gel [30, 112].

2.3.2 Biomedical Applications

PVA has previously been researched for use in biomedical and pharmaceutical applications due to its many advantageous qualities. The polymer is non-toxic, non-

- 28 -

Chapter 2 – Literature Review carcinogenic, hydrophilic, and can be easily modified through its pendant hydroxyl groups [105]. PVA hydrogels have been used in contact lenses, controlled release matrices and as bioadhesives [52, 113-115]. The use of PVA hydrogels as vascular prosthesis and cathethers, covalently coated with heparin as an antithrombotic agent, has been extensively studied [116-118]. Sefton and co-workers have extensively investigated the effect of incorporating heparin on the platelet activation and thrombin adsorption of PVA hydrogels [119]. In a chronic arterio-venous canine shunt model, platelets were found to be reactive upon contact with PVA but they did not remain adherent to the hydrogels after activation [120-121]. Immobilising heparin in the PVA hydrogel had no effect on platelets, rather, the PVA dominated the interaction with the platelets. The adsorption of thrombin onto PVA was also shown to be unaffected by the presence of heparin [122]. This indicates that in a dynamic system, the PVA is still reactive towards platelets even though the presence of heparin could suppress fibrin formation.

For use as a scaffold however, PVA hydrogels, like other synthetic polymers, lack recognition sites that would enable cells to adhere to [58]. The cell adhesion peptide,

RGD, has previously been attached to PVA hydrogels and shown to improve the attachment of fibroblasts [60]. Subsequent cell functions, however, rely on more complicated growth factor signalling processes, as modulated by ECM protein in vivo.

For this reason, the copolymerisation of PVA with biological polymers has been increasingly reported [123-124]. Copolymer of PVA and chondroitin sulfate has been reported for use in the encapsulation and delivery of chondrocytes to an injured site to encourage cartilage repair [99, 123]. The formation of degradable PVA hydrogels for use in cartilage regeneration [111], as artificial heart valves [58], and as soft tissue fillers [125] has also been investigated.

- 29 -

Chapter 2 – Literature Review

2.4 HEPARAN SULFATE

Heparan sulfate (HS) is a linear polysaccharide composed of alternating hexuronic acid and D-glucosamine residues, which are substituted to a varying extent with N- and

O-linked sulfate moieties and N-linked acetyl groups [56]. With the exception of heparin and hyaluronic acid, HS and other members of the glycosaminoglycan (GAG) family are synthesised as polysaccharide chains branching off a protein core, resulting in a structure called a proteoglycan [76]. The basic structure of a HS proteoglycan consists of a protein core to which several HS chains are covalently attached.

HS proteoglycans are primarily localised in two major areas, either associated with the plasma membrane on cell surface or with the basement membranes [126-127].

They interact specifically with a vast number of growth factors and cytokines. These interactions might simply serve to immobilise and protect the proteins from proteolytic degradation, while other times the HS chains help present these proteins to their cell surface receptors and modulate their activity [128]. Because of such interactions, HS proteoglycans are critically involved in a variety of biological processes at various levels of complexity, including embryonic development, angiogenesis, wound repair, and tumour progression. Although these functions occur mainly by the interaction of the proteins with the HS chains, most of the activities of HS proteoglycans depend to some extent on the presence of the protein core [129-130].

Heparin has the same basic disaccharide units as HS for its backbone, and is often considered an oversulfated variant of HS. Unlike HS, which is produced by most mammalian cell types and widely distributed on the cell surfaces and in the basement membranes, heparin is synthesised exclusively by mast cells in connective tissues and stored in cytoplasmic granules [131]. Nevertheless, the closely related structures of

- 30 -

Chapter 2 – Literature Review heparin and HS result in the two molecules sharing similar functions. Due to its wide availability, heparin is often used as a model for the functional study of HS.

2.4.1 Biosynthesis

A variety of protein cores have been shown to carry HS chains. Those anchored to the plasma membranes, such as syndecan and glypican, form the building blocks for cell-surface HS proteoglycans. Those associated with the basement membranes form pericellular HS proteoglycans, which include agrin, perlecan, and collagen type XVIII

[56, 132]. Heparin, on the other hand, is produced as side chains that decorate serglycin, an intracellular proteoglycan [129].

The initial attachment of HS/heparin to the protein core and the subsequent chain lengthening involve complex, multistep processes that are regulated by membrane- bound enzymes in the endoplasmic reticulum and Golgi apparatus. The HS side chains are covalently added to the serine residues on the protein core through a linkage tetrasaccharide, consisting of β-xylose, β1-4-galactose, β1-3-galactose and β1-3- glucuronic acid (Figure 2.4). The alternating addition of the N-acetyl glucosamine

(GlcNAc) and uronic acid (either a D-glucuronic acid (GlcA) or L-iduronic acid (IdoA)) residues, the predominant disaccharide repeat units of HS, are mediated by the enzymes

GlcNAc transferase II and GlcA transferase II, resulting in the elongation of HS chains

[56].

- 31 -

Chapter 2 – Literature Review

GlcNSO3 O-SO3 HexA n GlcNAc GlcA n D G D A G X

Figure 2.4 The basic structure of a heparan sulfate proteoglycan. There are typically several domains (D) along the protein core and, for the case of membrane-bound HS proteoglycans, an anchoring region (A) [129].

HS chain maturation occurs during or closely after chain elongation. This process includes partial N-deacetylation/N-sulfation of GlcNAc units, epimerisation of

GlcA into IdoA, and incorporation of O-sulfate groups at the C2 position of either GlcA or IdoA, and at C3 and C6 on glucosamine [128]. This process generates regions of high sulfate and IdoA content, interspersed by regions where the modifications are less extensive and the original structure retained [126]. Heparin resembles this highly sulfated region of HS, and in general, it is synthesised with a higher degree of sulfation and epimerisation, resulting in more N- and O-sulfate groups and a higher proportion of

IdoA units than HS, which instead contains more GlcNAc and GlcA residues. This variation in structural combination leads to the intricacy of the HS/heparin molecules, but despite the heterogeneous nature of these molecules, their interactions with HS- binding proteins are mediated through structure-specific saccharide sequences. The structural similarity of heparin and HS is illustrated in Scheme 2.3 [133-134].

- 32 -

Chapter 2 – Literature Review

2CH OSO3- O

OH O

NH GlcNpAc6S HOOC 2CH OH O O O 3CH O O OH OH 2CH OH n O OH NH O O GlcAp GlcNpAc OH 3CH GlcNpS NH SO3- 2CH OSO3- Heparan sulfate O

OH O GlcNpS6S

NH SO3-

2CH OH O O COOH O O OH OH

OH 2CH OSO3- NH SO3- O IdoAp O GlcNpS COOH O O OH OH n OSO - NH SO3- 3 CH OH COOH 2 IdoAp2S GlcNpS6S O O O O OH OH Heparin NH OH GlcNpAc GlcAp O CH3

Scheme 2.3 The predominant repeating disaccharides of heparan sulfate (HS) and heparin. The additional monosaccharide substituents of the hexuronic acid and glucosamine are described.

2.4.2 Protein Interactions

Beside serving as a scaffold for the attachment of various ECM components (e.g., collagen, laminin, fibronectin), the binding of HS to certain proteins has been suggested to induce a conformational change which may lead to the exposure of novel reactive determinants or conversely stabilize an inert protein configuration [76]. In most cases, this binding facilitates but is not required for ligand-receptor interaction and signalling

- 33 -

Chapter 2 – Literature Review

[135]. HS interactions with cytokines, such as interleukin and platelet factor 4, actually serve to localise the cytokines to various extracellular compartments as well as controlling their activities. The potential interaction of the ligands with HS in the ECM serves to set up and maintain gradients of these signalling molecules, an important feature in biological processes where cells are required to migrate through various tissues and organs.

The binding of proteins to HS/heparin chains ranges from those with relatively low affinity and specificity, to highly specific interaction requiring a particular saccharide region along the HS/heparin backbone. The sulfation pattern along the chains has also been shown to play a significant impact in creating the particular saccharide sequence for proteins to adhere to. A concrete example of this highly specific interaction is the binding of antithrombin III (ATIII) to heparin, which only occurs if a certain pentasaccharide sequence – known as the ATIII binding site – is present and available on the heparin backbone [136].

2.4.2.1 Interactions with growth factors

The interaction between growth factors and HS has been studied extensively with particular focus on FGFs [101, 137]. This family of growth factors bind heparin with relatively high affinity, and therefore termed ‘heparin-binding growth factors’. This binding translates to the ability to bind HS present on proteoglycans on the cell surface and in the ECM. The minimal binding sequence of HS for FGF-2 has been variably reported to be a tetra- [138], penta- [139] or a hexasaccharide [140], containing the essential O-sulfate group at C2 of the iduronic acid. The smallest biologically active sequence required for FGF-2 signalling was determined to be a decasaccharide, where the longer chain length is necessary to bridge the physical distance between the growth

- 34 -

Chapter 2 – Literature Review factor and its cognate receptor [56, 139]. When complexed with HS, FGF-2 has been shown to be protected from proteolytic degradation, and is consequently able to maintain a capacity for long term (24–48 h) endothelial cells stimulation [141-142].

The exact nature of the interaction between HS, FGF, and the tyrosine kinase FGF receptor (FGFR) is still not fully understood. A few models have been proposed to describe the HS/FGF/FGFR interaction that facilitates mitogenic activity, some suggesting simulataneous binding of FGF and FGFR by HS in a ternary complex [101], while others allude to the idea that HS assist in FGF dimerisation which leads to the activation of FGF-2 receptors [143-144]. As for the absolute requirement of HS for

FGF-2 signalling, FGF-2 activity was shown to be abolished in the absence of HS, which was thought to occur from the inability of FGF-2 to bind to its receptor [145].

Some other studies, however, have shown that FGF-2 can signal without HS, but the presence of HS was shown to enhance cellular signalling of FGF-2 [100]. Some studies suggest that HS/heparin interacts with both FGF-2 and FGFR [139, 146], while others have suggested that HS/heparin changes the conformation of FGF-2 thus increasing the

FGF-2–FGFR affinity [145, 147-148]. In either case HS/heparin is believed to facilitate

FGFR dimerisation and subsequent activation.

The binding of another heparin-binding growth factor, VEGF, to HS has been shown to be important in vivo in the developing mouse vasculature where localised forms of the growth factor are essential for successful branching morphogenesis [132,

149-150]. N-sulfate and 2-O-sulfate groups on the HS backbone are important for

VEGF binding whereas the 6-O-sulfate groups play a key role in cell signalling events.

Understanding the nature of the interactions of FGF-2 (basic FGF) with FGFR and the role of heparin/HS in this process is a fundamental issue that is still poorly understood [151]. HSPG of the same type can have different biological effects that

- 35 -

Chapter 2 – Literature Review depend on the cell type that synthesise them and the environment. For example, perlecan has been shown to be a potent inhibitor of SMC adhesion, migration and proliferation, but its effect on EC proliferation appears to be strongly stimulatory [149].

Perlecan, a large HS proteoglycan that is expressed in the ECM and basement membranes, is the major extracellular HS proteoglycan in blood vessel wall. The different perlecan activities towards SMC and EC appear to be related to the distinct cell surface HS of the two cell types. Studies have shown that ultimately the bioactivity of

HS proteoglycan such as perlecan is dependent on its cell origin, subtle changes in structure, concentration and binding kinetics with growth factors and their specific receptors [137]. Combined with its inhibitory effect on thrombosis, perlecan has been considered for use as pharmacological treatment to promote rapid healing after stenting.

2.4.3 Role in Vascularisation

HSPG controls various aspects of vascular development and tumour angiogenesis by firstly regulating the action of angiogenic growth factors. These HS/heparin-binding

GFs are highly mitogenic for endothelial cell proliferation in vitro and can induce angiogenesis in vivo at picomolar quantities [140]. HS acts in concert with members of the FGF and VEGF families, and their receptors [132]. Despite the ubiquitous presence of these angiogenic factors in normal tissue, the endothelial cell proliferation in these tissues is usually very low. It has since been found that these factors are stored in the basement membrane, sequestered from their site of activity. In an experiment with bovine cornea, FGF-2 was found to be stored in the basement membrane, attached to

HS proteoglycans and only released when competitive binding ligands for FGF-2 were present, or when the tissue was treated with heparanase [140]. Alternatively, the release

- 36 -

Chapter 2 – Literature Review of growth factors from HSPG can also be facilitated by the cleavage of the protein core by various proteases [152].

Heparanase, a unique mammalian enzyme, appears to be the only enzyme responsible for digesting the HS chains [130]. Platelets have been shown to be a good source of heparanase. Other sources of heparanase include the endothelial cells and smooth muscle cells, although the HS-degrading activity from these cells has been reported to be less than 10% that of platelet extract [153]. Recent studies have indicated that this heparanase activity not only cause the release of HS-bound growth factors but also create highly active HS fragments [149]. Figure 2.5 illustrates the heparanase degradation of HS chains to release growth factors from the ECM, making them available to interact with their high-affinity receptor on endothelial cells.

HS/heparin HSPG protein core Growth factor

heparanase

Endothelial cells

Extracellular matrix

Figure 2.5 The release of HS-bound growth factors by enzymatic cleavage of HS chains. The released growth factors act as chemotactic signals for endothelial cell migration and stimulate the formation of endothelial capillaries [154].

- 37 -

Chapter 2 – Literature Review

2.6 SUMMARY

The presence of molecular cues in a synthetic scaffold is essential for cell recognition and interaction. The advantageous properties of hydrogels have generated increasing interest in their use as scaffolds for tissue repair, while advances in polymer technologies has enabled the tailoring of hydrogel microstructure and the covalent incorporation of molecular cues. The use of heparin, physically blended or covalently incorporated, has been prominent in both natural and synthetic scaffolds, as molecules that protects growth factors against degradation, and also as a parameter that controls the rate of growth factor delivery. The presence of heparin in the scaffolds have been shown to enhance the effect of the growth factors being delivered, in inducing cell proliferation or even local tissue vascularisation in animal models. The specific activity of heparin, as a biomaterial, in signalling growth factors in these processes, however has not been widely investigated. No study has also investigated the effect of heparanase degradation on the activity of heparinised biomaterials.

The next chapters of this thesis describe the steps and methods used in the incorporation and assessment of heparin, as a model of HS, as signalling molecules in a synthetic hydrogel. A synthesis method is selected for the modification of heparin for covalent incorporation, and investigated for its effect on heparin activity. The properties of the heparin-incorporated PVA, including structural and mechanical properties, bioactivity, and enzyme degradability, are assessed to determine their potential for use as scaffolds for tissue repair.

- 38 -

Chapter 3

From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

ovalent incorporation of biomolecules onto polymer scaffolds results in a C sustained presentation of the biomolecules. Poly(vinyl alcohol) (PVA) and heparin were both functionalised to create macromers carrying photopolymerisable methacrylate pendant groups. PVA macromers from glycidyl methacrylate (GMA) synthesis were compared to those synthesised by reaction with 2-isocyanatoethyl methacrylate (ICEMA), which has been well studied. Even though the ICEMA synthesis had higher methacrylate incorporation efficiency than GMA synthesis, GMA- and ICEMA-functionalised macromers that had similar functional group densities formed hydrogels with comparable percent macromer, swelling, and mechanical properties. The relative cell compatibilities of both (GMA and ICEMA) macromers and hydrogels were also examined. By using GMA, the same functionalisation method was able to be used to modify PVA and heparin, resulting in PVA and heparin macromers that have functional groups of the same structure and equal reactivity.

- 39 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

3.1 BACKGROUND AND AIMS

The incorporation of biological molecules into synthetic scaffolds has been variously attempted, from simple physical incorporation to the more complex covalent attachment. Covalent incorporation of biological molecules generally results in a sustained presentation of these molecules by the polymer scaffold, which is a desired characteristic when long term applications are concerned. In designing a scaffold that can support wound healing and tissue regrowth, it is important to consider the natural components of the extracellular matrix that are responsible in mediating these processes.

Heparan sulfate (HS) proteoglycans in the extracellular matrix are essential regulators of growth factors during tissue regeneration and vascularisation [56, 128, 132], therefore incorporating HS into a supporting scaffold will create a structure that may represent a basic mimic of the extracellular environment.

As outlined in Chapter 2, glycosaminoglycan (GAG) molecules like HS have been covalently immobilised onto biologically-derived and synthetic hydrogel scaffolds, in an attempt to encourage localised interactions of the scaffolds with specific proteins and cell surface ligands [41, 63, 155]. Glutaraldehyde was a common crosslinking agent used for biomolecule incorporation into hydrogels in early methods [118, 156], however residual glutaraldehyde in the resulting gels has been proven toxic to cells [77-79].

GAG incorporation into collagenous scaffolds has used amines and carbodiimides as crosslinking agents, which modify the carboxyl groups on the GAG chain [81, 157].

The attachment of functional groups such as hydrazides has also been shown to allow incorporation into vinyl-functionalised polymers [158-159]. The rate of crosslinking and the structure or mechanics of scaffolds produced by these methods could only be controlled by the ratio at which the GAG/crosslinking agent/polymer was added. The

- 40 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment reactivity among the functionalised components allow for in situ hydrogel formation and has the potential for use as injectable biomaterials [6, 8, 82]. However techniques such as photo-initiated polymerisation might be better in controlling factors such as gelation time, the homogeneity and shape of the formed hydrogel scaffolds [106], as well as still having the potential of in situ (transdermal) hydrogel formation [160].

Photo-polymerisation of synthetic polymers has been found to be a method that can allow spatial and temporal tailoring of the crosslinking mechanism [30]. This technique enables in situ hydrogel formation, with the aid of non-cytotoxic initiators

[161]. To enable this crosslinking process, the polymers need to be modified with polymerisable pendant groups, typically vinyl-carrying groups that will act as crosslinkers upon photo-initiation (Figure 3.1). The resulting polymers carrying these functional groups are called macromers. The use of divinyl macromers such as poly(ethylene glycol) (PEG) has been prominent, however multifunctional macromers would offer an additional way to tailor the network properties of the hydrogels.

Multivinyl macromers would allow copolymerisation with other multi-, di-, or even mono-vinyl macromers.

A

B Light / UV

+

photoinitiator

Figure 3.1 Photocrosslinking of polymers carrying multiple pendant functional groups. The different macromers that can be copolymerised are represented by A and B. Following light or UV initiation, the photoinitiators split into radicals ( ) that attack the functional groups on the macromers, starting the formation of kinetic chains (---). Formation of a cycle is demonstrated (
), in which the functional groups of a macromer chain react only with one another instead of with those from other macromers, therefore preventing the formation of chemical crosslinks.

- 41 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

The abundant hydroxyl groups along the backbone of poly(vinyl alcohol) (PVA) allow for multiple substitutions with various pendant groups, giving PVA the advantage for structure modification over other synthetic polymers. Several research groups have explored the synthesis of PVA macromers for photocrosslinking, notably by modifying the hydroxyl groups on PVA with methacrylate and acrylate groups. Acrylic and methacrylic acid [108], glycidyl acrylate [109], glycidyl methacrylate [106, 110], and 2- isocyanatoethyl methacrylate [99] have all been used as precursors of these functional groups.

More recently, methods used for the attachment of functional groups onto synthetic polymers have been adapted for biological polymers [54]. The synthesis of photopolymerisable macromers derived from polysaccharides has been demonstrated.

GAG macromers have been crosslinked to form pure GAG hydrogels, and have been crosslinked with synthetic macromers, most notably PEG, to form co-hydrogels.

Methacrylate substitution on chondroitin sulfate (CS) by reaction with glycidyl methacrylate [74, 98] and methacrylic anhydride [97, 99] has been reported to result in photocrosslinkable CS macromers. The methacrylic anhydride chemistry has also been extended to modify heparin [9, 75] for covalent attachment to PEG gels.

This chapter focuses on the functionalisation of PVA and heparin with pendant groups that could act as crosslinks during photopolymerisation. When copolymerisation is concerned, both PVA and heparin macromers should present functional groups of equal reactivity, to minimise selective crosslinking among individual macromers. Most of the existing work has used different synthesis methods for synthetic and biological macromers, mostly due to the insolubility of biopolymers such as GAGs in organic solvent. Moreover, no studies have investigated the cell compatibility of the pendant functional groups, an aspect that is important when in situ hydrogel formation or cell

- 42 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment encapsulation is considered, because the cells will be exposed to the functional groups prior to them forming stable crosslinks.

Therefore the aim of this Chapter was to investigate the formation of PVA and heparin macromers using the same reaction mechanism, which provides pendant groups of the same crosslinking reactivity. Glycidyl methacrylate (GMA)-functionalised PVA was compared with PVA macromer that was functionalised using 2-isocyanatoethyl methacrylate (ICEMA) in DMSO. The fabrication of PVA hydrogels from ICEMA- synthesised macromers is well known and has been used in commercial applications

[162-163] as well as in tissue engineering research [99]. The PVA hydrogels formed from GMA- and ICEMA-functionalised macromers were compared in terms of their percent macromer, which indicates the hydrogel crosslinking efficiency and their strength. The cell compatibilities of both types of the PVA macromers were also compared. The term PVA-methacrylate (PVA-MA) describes PVA macromers in general, but in the comparison study between GMA and ICEMA synthesis, the PVA macromers and hydrogels in this study were named by their methacrylate precursor and amount of functional groups. For example, GMA-synthesised PVA macromers with 3 methacrylate (MA) groups/chain, along with hydrogels formed subsequently from these macromers, will be referred to as GMA3. The heparin macromers will be referred to as heparin-methacrylate (hep-MA).

3.2 EXPERIMENTAL

3.2.1 Materials

Poly(vinyl alcohol) (PVA) (average MW 13-23 kDa, 98% hydrolysed), heparin sodium salt (grade I-A, from porcine intestinal mucosa, average MW 17-19 kDa), glycidyl methacrylate (GMA) (97% purity), 2-isocyanatoethyl methacrylate (ICEMA)

- 43 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

(98% purity), 2,6-di-tert-butyl-4-methylphenol and deuterium oxide (D2O) were obtained from Sigma and used as received. The photoinitiator, 2-hydroxy-1-[4-

(hydroxyethoxy)phenyl]-2-methyl-1-propanone (Irgacure 2959, Ciba Specialty

Chemicals), was used as supplied. Dimethyl sulfoxide (DMSO), chloroform, acetone, and toluene (Sigma) were reagent grade and used as supplied. Silicone sheets (Silastic®

Sheeting, reinforced medical grade silicone rubber, Dow Corning) were perforated for use as hydrogel moulds. Nitrocellulose ultrafiltration membranes (10 kDa molecular weight cut off, Millipore) were used as received.

3.2.2 Synthesis of PVA Macromers

2-isocyanatoethyl methacrylate (ICEMA) synthesis. Methacrylate pendant groups were introduced by reacting PVA with ICEMA [99]. PVA was dissolved in DMSO to make a 10% (w/v) solution, in an 80°C waterbath. Small amounts of 2,6-di-tert-butyl-4- methylphenol was added to the PVA solution as an inhibitor to prevent polymerisation during macromer synthesis, and the solution was purged with N2 gas for 30 minutes to remove traces of water. For 1 g PVA, 45 µL of ICEMA was added and the mixture was allowed to react with stirring at 60°C under a N2 atmosphere. After 4 h, the reaction was stopped by precipitating the PVA in toluene. The precipitate was then redissolved in water and ultrafiltered with a 10kDa MW cut-off (MWCO) membrane, before being lyophilised.

Glycidyl methacrylate (GMA) synthesis. PVA was dissolved in phosphate buffered saline (PBS) (pH 7.4) at 80ºC to prepare a 10% (w/v) solution. GMA was added drop-wise to the PVA solution with vigorous stirring, in a 1:2 molar ratio to the number of PVA repeating units. The amount of repeating unit (RU) in a given polymer chain can be calculated from by:

- 44 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

MW mol RU =polymer × mol polymer MW RU (3.1) where MW = molecular weight.

The reaction was done at room T for up to 10 d, and was stopped by precipitation in acetone. After washing with chloroform, the PVA was ultrafiltered with a 10 kDa

MWCO membrane and lyophilised.

3.2.3 Synthesis of Heparin Macromers

Heparin was reacted with GMA to provide polymerisable pendant groups using a heterogeneous-phase method that was adapted from Li, et al. [74]. In a typical experiment, 1 g heparin was dissolved in PBS, pH 7.4, to make up a 10% (w/v) solution. GMA was added in equal molar amount as the number of moles of the disaccharide RU (1 mole heparin is made of ~40 moles disaccharides). The GMA

(0.326 g) was added with vigorous stirring and the reaction solution was left to react at room temperature for 14 d. To stop the reaction the macromer was precipitated in acetone, redissolved in water and then washed twice with chloroform to remove residual

GMA. The heparin was ultrafiltered to remove compounds less than 10kDa in size and was then lyophilised to obtain a dry product.

3.2.4 Characterisation of Methacrylated Macromers

Nuclear magnetic resonance (NMR). 1H NMR (300 MHz Bruker Avance DPX-

300 spectrometer) was used to analyse the amount of methacrylate attachment. To calculate the percent methacrylation of PVA, two 1H NMR peaks representing the two methacrylate vinyl protons (see Figure 3.2, H1 and H2) (δH1 ~6.1 and δH2 ~5.7 ppm)

- 45 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment were compared to the peaks representing the protons in the PVA backbone (δ 3.8–4.1 and 1.4–1.8 ppm).

2 2 1 1 2 m n OH O OH

O O 1

H1 H2

H1 H2

6.0 5.6 5.2 4.8 4.4 4.0 3.6 3.2 2.8 2.4 2.0 1.6 1.2 (ppm) 1 Figure 3.2 H NMR of PVA-methacrylate in D2O. Inset: schematic numbering for the methacrylate pendant group and the PVA backbone.

Similarly, percent methacrylation of heparin was calculated by comparing the area of the methacrylate vinyl proton peaks (Figure 3.3, H1 and H2) to the peaks representing protons 4-11 (δ 3.4–4.6 ppm) and proton 12 (δ 3.0–3.4 ppm) on the repeating disaccharide unit of heparin [133].

The amount of methacrylate groups per polymer (PVA or heparin) chain could then be calculated by:

MW crosslinker / chain= % methacrylation × polymer (3.2) MWRU

- 46 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

OSO - 4 3 6 3 H1 H2 O 9 O OH 1 8 O 10 O 2 O O R OH 5 OH 12 3CH n 7 11 OSO - NH R' O 3 10 13 3 5

11 7 8

2 6 9 1 12 4

H1 H2

5.6 6.0 5.2 5.6 4.8 5.2 4.4 4.8 4.0 4.4 3.6 4.0 3.2 3.6 2.8 3.2 2.4 2.8 2.0 2.4 1.6 2.0 1.6 (ppm) 1 Figure 3.3 H NMR of heparin-methacrylate in D2O. Inset: schematic numbering for the - methacrylate group (left) and heparin (right). R = heparin, R’ = –SO3 (predominant) or –COCH3, which protons account for peak 13.

3.2.5 Hydrogel Formation

Hydrogels were prepared with 20% (w/w) macromer concentration in water. Once the PVA or heparin macromer was dissolved, the photoinitiator (Irgacure 2959) was added to make up a final concentration of 0.1% (w/w) initiator. The macromer solution was poured into a disc-shaped silicon mould (10 mm diameter × 1 mm thick) and cured under a UV light source (UVA 300-480nm, 365nm peak, Green Spot). The UV intensity and exposure time were varied to determine the optimum photopolymerisation condition.

Swelling property and sol fraction determination. Hydrogel sol fraction refers to the portion of macromers that are not incorporated into the network. It has been previously determined to be extracted in the first 24 h and therefore equals the hydrogel mass loss at 24 h [30, 112]. Immediately after polymerisation each hydrogel disc was

- 47 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

weighed (m0), and hydrogels discs were made in triplicates for each type of polymer. A t0 sample was taken and lyophilised without immersion in buffer, while the rest of the samples were each immersed in PBS. At 24 h the gels were removed from the buffer, patted dry and weighed to give the swollen weight of the discs (ms). The gels were then lyophilised to obtain their dry weights (md). The actual amount of macromer in the discs

(macromer fraction) was calculated using the dry weight of the t0 sample.

m macromer fraction = d,t0 (3.3) m 0,t0

The macromer fraction was used in combination with the original wet weights (m0) to obtain the hydrogels initial dry weights (mid)

mid = m 0 × macromer fraction (3.4)

The volumetic swelling ratio (Q) was calculated by: ρ Q 1+= polymer ()q −1 (3.5) ρ solvent where q is the mass swelling ratio (q = ms/md), ρpolymer is the density of the macromer

(for PVA macromers, approximated to the density of PVA = 1.2619 g/mL), and ρsolvent is the density of PBS, approximated to that of water to be ~1 g/mL.

The hydrogel sol fraction, i.e. mass loss at 24 h, was calculated as follows:

m - m mass loss =id d ×100% (3.6) mid

3.2.6 Tensile Testing of PVA Hydrogels

PVA samples were photopolymerised in a dumbbell-shaped mould, with the same macromer and initiator concentrations as described above. Immediately after polymerisation, the sol fraction of the hydrogels was extracted by incubating in PBS for

24 h to ensure all non-crosslinked macromers have leached out [30, 112]. The tensile

- 48 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment tests were done on an Instron 5543 universal testing machine, fitted with an immersion bath and an advanced video extensometer (AVE). The immersion bath was used to maintain temperature and water content in the hydrogels, as well as to eliminate the effect of sample dehydration during testing.

The testing area of the samples was 2 mm wide and 1 mm thick, and two dots were drawn on the ends of the hydrogel to mark the tested gauge length (Figure 3.4).

Samples were mounted onto corrugated grips on the instrument to prevent slippage, and then immersed in the bath. The AVE system used a camera to detect the two marks on the hydrogel and monitor the distance between the two points during testing. Tensile tests were performed at a 3-mm/min strain rate until failure.

Gauge length

Figure 3.4 Hydrogel specimen for immersed tensile testing. A dumbbell-shaped, 1-mm thick hydrogel is shown with the marked gauge length and dimensions.

3.2.6 Cell Growth Inhibition Assay

Confluent mouse fibroblasts (L929) were trypsinised and seeded at 1×105 cells per 35-mm tissue culture dish (~1×104 cells/cm2). The cells were incubated in 2 mL media (Earle’s minimum essential medium (EMEM) + 10% FBS) per dish for 24 h

(37ºC, 5% CO2) to produce a sub-confluent layer of adherent cells. The culture dishes were then washed with sterile DPBS before test or control solutions were introduced to

- 49 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment the cells. The controls used were a null (media only) and ethanol (EtOH) solutions at 4,

5, 6 and 7.5% (v/v) in media as positive controls.

Preparation of test solutions. Macromer solutions. Test solutions of PVA and

PVA-MA (10 wt% in saline), were filter-sterilised and diluted with media at a volumetric ratio of 1:3 to give a final polymer concentration of 2.5% (w/v). A sterile saline-only solution was also mixed 1:3 with media as a control. Hydrogel extracts.

PVA hydrogel discs (10 mm diameter × 1 mm thick) were formed and each placed in an extraction vial. The sol fraction of each disc was extracted in 1.5 mL saline, at 37ºC for

24 h. A saline-only solution was incubated in an extraction vial as a control. The extract solutions were then filter sterilised and mixed in a 1:3 ratio with media.

The cells were incubated in the test and control media for 48 h, and each sample was done in triplicate. At the end of incubation period the cells were harvested from the dishes. The cell number was counted using a Vi-Cell cell counter apparatus (Coulter), which also had a built-in Trypan blue staining system to measure viability. Cell growth inhibition was calculated as the decrease (%) in cell number for each sample type relative to the null. Positive controls in 7.5% EtOH was expected to show greater than

70% inhibition to indicate the assay was valid. A sample was considered cell inhibitory if showing higher than 30% cell growth inhibition [164].

Statistical analysis of cell growth inhibition. To compare the effects of methacrylate groups on the L929 cell growth, the calculated percent cell growth inhibition was analysed with an analysis of variance model [165] with two fixed factors

(macromer type and amount of methacrylate group) and one random factor (experiment) using Minitab 15 [166].

- 50 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

3.3 RESULTS AND DISCUSSION

The objective of this chapter was to modify PVA and heparin with functional groups with equal reactivity for photocrosslinking. Functionalisation using the same precursor provides one way to achieve the same structure and reactivity of the resulting pendant groups. Methacrylate functionalisation of PVA has been done by reaction with various methacrylate precursors in aqueous and organic solvents. The ICEMA synthesis, in particular, has been well studied and has been shown to be easily controlled with an incorporation efficiency of 80-100% [99, 162-163]. Chemical modification of biopolymers such as heparin, on the other hand, has been limited due to their insolubility in organic solvents.

In this work, ICEMA-synthesised PVA macromers were made and photopolymerised, so that the crosslinking efficiency and the mechanical properties of the ICEMA hydrogels could be used as a reference for PVA synthesised using GMA.

The cell compatibility of the macromers and hydrogels were also examined. The GMA synthesis can be performed in aqueous media and has been shown to functionalise

GAGs similar to heparin [74], therefore the viability of using GMA synthesis on both

PVA and heparin was assessed, and the optimum copolymerisation conditions were determined.

3.3.1 Synthesis of PVA Macromers

The PVA-ICEMA reaction in this study was carried out in DMSO and the amount of methacrylation was dependent on the ratio of ICEMA addition to the amount of hydroxyl groups on PVA. The average substitution efficiency for all ICEMA synthesis was 80%. The synthesis of PVA macromer by reaction with GMA demonstrated a time dependent attachment of MA groups onto PVA backbone (Table 3.1).

- 51 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

Table 3.1 Efficiency of PVA-methacrylate synthesis by reaction with glycidyl methacrylate. Reaction efficiency as a function of reaction time. Reaction time MA-attachment MA groups/ Substitution (d) (mol%) PVA chain efficiency (%)

1 0.4 1.5 (~1) 0.8

4 0.975 3.5 (~3) 2.0

7 1.09 4.0 (~4) 2.2

10 1.41 5.1 (~5) 2.8

14 1.87 6.8 (~7) 3.7

Scheme 3.1 (A) and (B) illustrate the synthesis of PVA macromers by reactions with GMA and ICEMA, respectively. Despite carrying the same methacrylate end groups, the overall structure of the crosslinkers molecules from GMA and ICEMA syntheses differed slightly. This might have an effect on cell compatibility of the macromers, which is of significant importance when hydrogels are to be used for tissue engineering scaffold.

A

O m + O n OH n OHO OH OHOH O OH

O O

B O + NCO m OH O n n O O OH OH OH OH NH

O

O

Scheme 3.1 Synthesis of poly(vinyl alcohol) (PVA) macromer. Functionalisation of PVA by reaction with glycidyl methacrylate (A) and 2-isocyanatoethyl methacrylate (B).

- 52 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

3.3.2 Effect of Macromer Attachment on the Cell Compatibility of PVA

Therefore the PVA macromers were assessed in a cell growth inhibition (CGI) study, in order to examine the effect of adding methacrylate pendant groups on the cell compatibility of PVA and to compare the GMA- and ICEMA-sourced functional groups. The non-modified PVA has been found to be low in toxicity, with an acute oral toxicity (LD50) of 15-20 g/kg [167]. All polymer solutions were prepared at the same concentration in saline, and at 2.5% (w/v) (25 mg/mL) the polymer concentration was higher than would be expected from the release of hydrogel sol fraction.

GMA and ICEMA macromers with 3 (GMA3 and ICEMA3) and 5 (GMA5 and

ICEMA5) MA groups per chain were dissolved in saline and applied to the culture media of L929 mouse fibroblasts. The effect of these macromers on cell growth after 48 h was examined in terms of cell number and viability, and was compared to results from

PVA. Cell growth inhibition was determined as the decrease in cell number in test solutions compared to an unperturbed control, which were cells incubated in culture media only. Figure 3.5 shows the amount of cell growth inhibition as a response to the varying macromer type and crosslinker density.

45

30

15 Cell growth Cell inhibition (%)

0 media only saline PVA PVA-MA PVA-MA PVA-MA PVA-MA EtOH 4% EtOH 5% (null) GMA3 GMA5 ICEMA3 ICEMA5 (+ve) (+ve)

Figure 3.5 L929 cell compatibility of PVA macromers. PVA and PVA-MA were at 2.5% (w/v) in media. Percent cell growth inhibition was calculated as the increase in cell number compared to cell cultured in media only.

- 53 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

Figure 3.5 above shows that incubation with 2.5% (w/v) (25 mg/mL) PVA caused

15 ± 6 % inhibition in cell growth, however the presence of methacrylate functional groups in PVA-MA did not appear to elevate the cell growth inhibitory property of

PVA. Analysis of variance confirmed the effect of methacrylate functional groups on cells to be insignificant (p>0.05). All the positive controls showed elevated cell inhibition, with EtOH concentrations of 5% and higher causing significantly higher cell inhibition compared to the null and the test samples (p<0.01). Cells that were incubated with 7.5% EtOH experienced a cell growth inhibition above the 70% level, confirming the validity of the assay [164]. From these results, the presence and amount of crosslinkers on the PVA macromer have been statistically shown to have no effect on cell growth.

3.3.3 PVA Hydrogels from GMA- and ICEMA-Synthesised Macromers

3.3.3.1 Hydrogel swelling and sol fraction

To characterise the network structure and strength of hydrogels made from GMA- synthesised PVA macromers, they were compared to gels made of PVA-MA that was synthesised by reaction with ICEMA. The ICEMA synthesis was previously described to prepare PVA macromers for hydrogel contact lenses [162], and used in a study investigating hydrogels for cell encapsulation [99]. The concentration of macromers has been shown to have the strongest effect in influencing the final structure and strength of the hydrogels [112]. Therefore, by keeping the percent macromer constant at 20 wt% and varying the amount of functional groups per polymer chain (i.e. crosslinker density), the GMA and ICEMA macromers were investigated for their crosslinking efficiency, network structure and strength, and also cell compatibility.

- 54 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

The swelling capacity of a hydrogel depends on the type of network formed by its crosslinked macromers. Macromers with higher crosslinker density have been shown to form a tighter network than those with lower number of crosslinkers, consequently decreasing its water-swelling capacity. Macromers with higher crosslinker density also typically result in hydrogels with lower sol fraction. After the 24-h period needed for sol fraction extraction, hydrogels with non-degradable crosslinks are considered at equilibrium while hydrogels with hydrolysable crosslinker groups such as ester continue to degrade and lose more mass [111]. Both GMA- and ICEMA- sourced functional groups do not have sites that permit chain scission by hydrolysis.

The photocrosslinking efficiency of multifunctional macromers is not simply dependent on the conversion of the functional groups. Martens and Anseth showed that even though a 100% conversion of acrylate pendant groups was achieved, PVA gels were still formed with ~35% sol fraction [30]. Cycles might be formed, where functional groups within one polymer chain react only with one another but not with those on adjacent chains (see previous Figure 3.1, (
)). The cycles are not covalently crosslinked and may be released; even if they are entangled with other chains, they do not contribute to the structure and strength of the hydrogels. The formation of cycles and having functional groups of different reactivity might be reasons behind this deviation from the ideal chain crosslinking process. Increasing the functional group density seemed to compensate for this irregularity in crosslinking. Earlier work done in our group showed a decrease in sol fraction by nearly 4-fold when PVA macromers with 7, instead of 4, acrylamides/chain were used [112], thus improving the overall crosslinking efficiency.

The trend of increased crosslinking efficiency with higher functional group density was seen with both the GMA and ICEMA hydrogels. Increasing the functional

- 55 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment group density of the GMA and ICEMA macromers from 3 to 5 decreased the sol fraction of the resulting hydrogels. GMA5 and ICEMA5 hydrogels had sol fractions of around 7%, compared to the GMA3 and ICEMA3 hydrogels that had 22 and 24% sol fractions, respectively (Table 3.2). Smaller sol fraction also means higher percent macromer in the resulting gels, consequently decreasing the swelling ratio of the gels.

GMA3 hydrogels swelled to ~7.1 times in volume, while GMA5 hydrogels only had a volumetric swelling ratio (Q) of ~5.5.

Table 3.2 Properties of PVA hydrogels from GMA- and ICEMA- synthesised macromers. All hydrogels were formed with an initial 20 wt% macromer.

Type of PVA Crosslinker density Volumetric Sol Tensile hydrogel of macromer (MA swelling fraction moduli groups/chain) ratio, Q (%) (kPa)

GMA3 2.8 (~3) 7.1 ± 0.05 24.7 ± 0.7 69 ± 22

GMA5 4.8 (~5) 5.5 ± 0.1 7.2 ± 0.3 179 ± 12

ICEMA3 3.0 (~3) 7.6 ± 0.05 24.3 ± 0.8 64 ± 6

ICEMA5 5.0 (~5) 5.1 ± 0.06 6.6 ± 0.6 181 ± 26

There were no significant differences in sol fraction and swelling properties of

GMA and ICEMA hydrogels with similar crosslinker densities, despite the difference in structure of the crosslinkers carrying the methacrylate end groups. In fact, the sol fraction results are directly comparable to gels made from acrylamide-functionalised

PVA. Hydrogels formed from macromers with 4 acrylamides had a Q of 7.4 and a sol fraction of 17%, while those formed from 7 functional groups per chain macromer had

Q of 5.3 with a 4.3% sol fraction [112]. This indicates that the crosslinking efficiency is not as much influenced by the type of functional groups present; rather the crosslinker density is a more determining factor. The swelling ratio, however, depends more on the

- 56 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment type and length of the crosslinker molecules, as the swelling ratios of GMA3 and

ICEMA 3 hydrogels are comparable to that of a PVA acrylamides with 4 functional groups per chain.

3.3.3.2 Mechanical properties

The mechanical properties, both tensile and compressive, of hydrogels have been found to be mainly influenced by, and increase with, the amount of polymers in the hydrogel network [30, 99, 112]. Factors such as polymer molecular weight and functional group density affect the crosslinking efficiency of photopolymerised hydrogels and thus influence the amount of macromers incorporated in the hydrogels.

The increase in tensile modulus and strength with increasing macromer content has been shown in co-hydrogels of 2-hydroxyethyl methacrylate (HEMA) and PEG, regardless of the copolymer composition [168]. A study on oligo(poly(ethylene glycol)fumarate)

(OPF) hydrogels, a PEG derivative, showed that hydrogels made from 860 Da macromers had higher tensile modulus (~89.5 kPa) compared to that of OPF gels made from 9600 Da macromers (~16.5 kPa), due to the higher degree of crosslinking in the former [169].

The tensile moduli of the PVA hydrogels tested in this study are summarised in

Table 3.2 above. The decrease in water swelling capacity when the amount of functional group density was increased from 3 to 5 MA groups/chain, was accompanied by an increase in the mechanical strength of the hydrogels. The average tensile modulus for

PVA hydrogels formed from the GMA3 macromers was 69 ± 22 kPa, while that from the GMA5 macromers was 179 ± 12 kPa. Although the tensile properties of PVA increased with the increase in MA pendant groups per macromer chain, the resulting gels had smaller strain at failure.

- 57 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

GMA and ICEMA hydrogels made from macromers with similar amount of methacrylate groups per chain (i.e. GMA3 vs. ICEMA3 and GMA5 vs. ICEMA5) were found to have comparable tensile moduli. The difference in modulus between the GMA and ICEMA hydrogels was found to be statistically insignificant.

3.3.3.3 Cell growth inhibition

When choosing the synthesis methods for biomaterials, it was important to compare the biological performance of the resulting materials. The physical characteristics of the GMA- and ICEMA- synthesised PVA hydrogels had been found to be comparable. The four types of PVA hydrogels were now tested for cellular toxicity

(cytotoxicity) in vitro. Performing cytotoxicity assay on hydrogel extracts allowed all cells to be exposed to the test materials, making it a sensitive study compared to the direct contact method [125, 170].

As stated previously, the presence of MA groups was shown not to increase the level of cell growth inhibition of PVA (see Figure 3.5). The sol fraction extracts from the four hydrogel types were tested in a CGI study, to examine their effects on mouse fibroblasts. This test would give a preliminary insight on cell reactions upon the implantation of hydrogels immediately after polymerisation and to the release of any sol fraction. All extractions were performed in saline, and therefore a saline control was used.

Comparing the mean cell responses to the hydrogel extracts, the amount of cell inhibition was mainly influenced by percentage of sol fraction released by the hydrogels

(Figure 3.6). As discussed previously, hydrogels made from GMA3 and ICEMA3 had higher sol fractions than those made from GMA5 and ICEMA5. The sol fraction of

- 58 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

PVA hydrogels of a 3-crosslinkers/chain macromer was around 20%, while hydrogels made from 5-MA groups/chain macromers had sol fractions of less than 10%.

45

30

15 Cell growth Cell inhibition (%)

0 media only extract I2959 extract extract extract extract EtOH 4% EtOH 5% (null) saline GMA3 GMA5 ICEMA3 ICEMA5 (+ve) (+ve)

Figure 3.6 L929 cell compatibility to the PVA hydrogel extracts. Extracts were sol fractions of the hydrogels. Percent cell growth inhibition was calculated as the increase in cell number compared to cell cultured in media only.

At the completion of photopolymerisation, residual initiator might be present in the hydrogel network. The initiator I2959, also known as D2959 in some literature, has been shown to cause minimal cytotoxicity across six mammalian cell types, even when activated into radicals by UV exposure [161]. Compared to several other water-soluble photoinitiators, I2959 was found to cause minimal cytotoxicity. When exposed to NIH

3T3 fibroblasts, I2959 was also shown to be cytocompatible at concentrations ≤ 0.5%

(w/w) [171]. Photoinitiators have been shown to induce less cell growth inhibition compared to chemical reagents such as reductants and oxidants used in the redox crosslinking process [125]. In this study, a 20 ± 2 % cell growth inhibition was observed when cells were incubated with 0.1% (w/w) I2959 in culture media (Figure 3.6 above).

This 0.1 wt% concentration, at which the I2959 was tested, was the initial amount of

I2959 present in PVA-MA solution prior to polymerisation. The amount of residual initiator released from the gels is expected to be less and thus would have lower cell growth inhibition.

- 59 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

The viability of the cells remained above 90% for all cases except for the 7.5%

EtOH control. The effect of the test materials on cell growth was seen in the difference in cell number compared to the null control. The high cell viability, coupled with the decreased cell number, indicated that the presence of the test materials might have decreased the rate of cell proliferation but did not directly caused cell death. The inhibitory effects of the 4, 5, and 7.5% EtOH controls from three independent experiments were consistent; the 7.5% EtOH resulting in less than 50% cell viability and ~85% relative cell growth inhibition. Saline contributed to the cell growth inhibition as shown by the saline controls in both the cell growth assays on the hydrogel extracts (Figure 3.6) and on the polymer solution (previous Figure 3.5). However, this increase in inhibition by saline compared to the null, and by the PVA hydrogel extracts compared to saline, were not statistically significant.

Results from the swelling, mechanical strength, and cell compatibility studies on the PVA hydrogels showed comparable properties between GMA and ICEMA hydrogels. The biggest drawback of GMA synthesis is the low methacrylate incorporation efficiency compared to ICEMA synthesis, however the same amount of methacrylation could be achieved. Percent methacrylation has been shown to be the property that most influences the resulting hydrogel’s structure and strength. Even though the GMA synthesis has low incorporation efficiency, this work aimed at low percent methacrylation. High swelling capability is one of the attractive properties of hydrogels for use as cell scaffold, and this could only be achieved when macromers have sufficiently low amount of methacrylation. Since all the properties of GMA and

ICEMA hydrogels were the same, the GMA synthesis could be chosen to functionalise

PVA, especially if it can be achieved with heparin.

- 60 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

3.3.4 Synthesis of Heparin Macromers When forming hydrogels by the photo-crosslinking of polymer chains, it is important for the crosslinkers that are attached on the chain to have similar reactivity to prevent selective crosslinking among groups of higher reactivity. Acrylate groups have been shown to be more reactive than methacrylate groups, while acrylamides are more reactive compared to methacrylamides [108]. PVA was modified with methacrylate groups by reaction with GMA in an aqueous media, and this method was now tested on heparin.

Bacterial polysaccharides such as dextran are soluble in DMSO and have been functionalised with methacrylate pendant groups for photocrosslinking [96], however

GAGs such as heparin have limited solubility in organic solvents. In the present study, heparin was functionalised in a heterogeneous-phase reaction using a method adapted from Li, et al., who investigated the modification of chondroitin sulfate (CS) with methacrylate pendant groups by a reaction with GMA [74]. Methacrylate substitution initially occurred by a transesterification reaction with the hydroxyl groups on CS, which was consistent with results found in the GMA-dextran modification in DMSO

[172]. This transesterification was followed by a GMA-ring opening mechanism that allowed MA substitutions on the sulfate and carboxyl groups on CS.

Heparin and CS are both members of the GAG family, and although differing in functions, their basic polysaccharide structure provides sites (e.g. hydroxyl groups) for modification by GMA. It was expected that similar reaction mechanisms would occur with heparin, in that heparin would be modified via both the transesterification and ring opening reactions (Scheme 3.2).

- 61 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

CH2 O O - CH Hep OH O O 3 + S A O O 2CH O O O CH2 O OH OH CH O O 2 O OH O OH + 3CH O n - - B 3CH O O NH O O O S O SO O S O O O O O + OH Hep CH2 O 3CH O O O Hep Scheme 3.2 Synthesis of heparin macromer. Heparin functionalisation by reaction with glycidyl methacrylate. A = transesterification, B = ring opening.

In theory, the proportion of MA-substitution by ring opening polymerisation could be calculated by comparing the NMR proton peaks that indicate ring-opening substitution on a sulfate (δ ~5.5 ppm) or on a carboxyl group (δ ~5.2 ppm) [74].

However in the case of heparin, these ring-opening indicator peaks overlap with protons

1 and 2 on the heparin backbone (see previous Figure 3.3). The overlapping peaks meant that it was not possible to quantify the proportion of MA-substitution from either the transesterification or ring opening reactions. Nonetheless, the total amount of MA- substitution could still be calculated from comparing the peaks of MA protons H1 and

H2 (δ ~6.1 and ~5.7 ppm, respectively), with the heparin proton peaks.

The GMA functionalisation of heparin was found to produce heparin macromers with 8 mol% substitution after a 14-d reaction, a result similar to the functionalisation of

CS [74]. To enable chemical crosslinking of polymer chains, the minimum average number of functional groups that needs to be present is two functional groups per polymer chain. The 8 mol% substitution translated to approximately 3 MA crosslinkers per heparin chain, which fulfilled the requirement of two crosslinkers per polymer chain to enable crosslinking to form heparin hydrogel. The formation and characterisation of heparin hydrogels will be further discussed in Chapter 5.

- 62 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

The efficiency of GMA functionalisation on PVA and on heparin was compared.

In terms of molar percentage, the functionalisation of heparin was more efficient than that of PVA. For a 14-d reaction with GMA, heparin had 8 mol% MA substitution while

PVA was only substituted with 4 mol% MA. However, these numbers become less relevant when calculating the actual number of MA groups that are present, on average, on a single PVA or heparin chain. An 8 mol% substitution on heparin equals approximately 3 MA groups per heparin chain, and to get the same number of functional groups on PVA, only a 2 mol% substitution is needed. Figure 3.7 shows the increasing amount of methacrylate substitution on PVA and heparin as the reaction time was extended.

8

7

6

5

4

3

2 MA groups/polymer chainMA

1

0 0 2 4 6 8 10 12 14 16 Reaction time (d) Figure 3.7 Methacrylate group attachment as a function of reaction time with glycidyl methacrylate. The amount of methacrylate groups attached to PVA (●) and heparin (○) are shown against the reaction time.

The attachment of MA groups on PVA was found to increase almost linearly with time and was still increasing at 14 d, at which the amount of functionalisation reached an average of 7 MA groups per PVA chain. Meanwhile, the attachment of MA to

- 63 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment heparin seemed to have reached a plateau by 7 d. The number of MA groups per heparin chain was ~3 after a 7-d reaction, and did not increase after extending the reaction time to 14 d. For the copolymerisation of PVA and heparin, PVA-MA and hep-MA macromers with an equal average amount of ~3 MA/chain were used.

3.3.5 UV-Polymerisation Conditions for PVA and Heparin Macromers

PVA and heparin macromers were synthesised using the same method to provide crosslinker groups of equal reactivity. Even though both macromers were of similar average molecular weights, and were functionalised with similar amount of crosslinkers

(~3 per polymer chain), the structure of the polysaccharide heparin chain is more complex than the linear PVA chain. PVA is a neutral polymer while heparin chains are negatively charged, and this might have an effect on their photopolymerisation behaviour.

The optimum polymerisation conditions for the formation of individual PVA and heparin hydrogels were determined by varying the intensity of the UV light and the length of exposure. The sol fraction of the hydrogel indicates the proportion of macromers that was not crosslinked into the network, and has been shown to be released in the first 24 h of incubation. Therefore measuring the sol fraction is a way to analyse the crosslinking efficiency of functional macromers, and complete crosslinking of the macromers was considered achieved when subsequent changes in UV intensity and exposure time did not alter the amount of sol fraction of the hydrogels.

Homopolymer hydrogels of PVA and heparin were formed and their sol fraction analysed. Figure 3.8(A) showed the sol fraction of the hydrogels as a function of the UV intensity that they were crosslinked at. Although there was a trend of decreased sol fractions with lower UV intensities, the amount of PVA sol fraction stayed similar

- 64 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment across the range of UV intensities used, suggesting that the light intensity had minimal influence on the efficiency of crosslinking of PVA macromers. This unexpected trend was more evident in heparin hydrogels. The amount of heparin macromers incorporated was lower at higher UV intensities, as indicated by the higher sol fraction. When polymerised at intensities lower than 50 mW/cm2, heparin hydrogels had less than

~35% sol fraction, but this increased when heparin was polymerised at intensities higher than 90 mW/cm2, up to ~60% sol fraction at 150 mW/cm2.

70 A 60

50

40

Sol fraction (%) 30

20

10 0 20 40 60 80 100 120 140 160 2 UV intensity (mW/cm )

60 B 50

40

30

20 Sol fraction (%) fraction Sol

10

0 0 5 10 15 2 UV, 30 mW/cm , exposure time (min) Figure 3.8 Photopolymerisation efficiency as a function of UV intensity (A) and UV exposure time (B). The sol fraction was used as a measure of efficiency in the crosslinking of PVA (●) and heparin (○) macromers.

- 65 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment

Although unexpected, polymerisation at high UV intensities (120−850 mW/cm2) has been reported to result in lower conversion in the crosslinking of mutifunctional methacrylates and acrylates macromers [173-174]. Initiator molecules dissociate into radicals when subjected to irradiation. Higher light intensity creates a greater number of radicals at a time, which leads to the formation of more, but shorter, kinetic chains

[175]. The decrease in kinetic chain length lowers the rate of polymerisation in crosslinked system thus increases the required polymerisation time [174, 176], which means that the crosslinking of PVA and heparin at UV intensities higher than 90 mW/cm2 (Figure 3.8(A)) would take longer than those polymerised at lower intensities.

The sol fractions of both PVA and heparin hydrogels had no significant changes following polymerisation at intensities lower than 30 mW/cm2. Therefore taking this intensity to use, PVA and heparin hydrogels were formed with different exposure times.

No change in sol fraction was observed for PVA after a 3-min polymerisation, however it took 6 min for heparin hydrogels to fully crosslink (Figure 3.8(B)). The anionic nature of heparin, which affects its conformation and the accessibility of pendant MA groups to react with each other, might also prevent physical entanglements of the chains. As physical entanglements increases crosslinking density, this would explain the higher sol fraction that a heparin gel had compared to a PVA gel, even when both were polymerised with the same original macromer concentration. Lower number of crosslinks per chain and smaller molecular weight are also factors that have been shown to result in higher hydrogel sol fraction [30]. If the heparin chains were fragmented by UV at higher intensities, the smaller macromers would have less average number of functional groups per chain and when polymerised will have less amount of crosslinking compared to longer-chained macromers. However when analysed by size

- 66 -

Chapter 3 – From Macromer Synthesis to Hydrogel Formation: Compatibility Assessment exclusion chromatography, heparin was found to remain intact after UV exposure, even after exposure to 0.3 W/cm2 UV for 30 min (result shown in Chapter 4).

The optimum UV-crosslinking condition for PVA and heparin macromers was determined to be 30 mW/cm2 intensity for 6 min. Decreased UV intensity and increased polymerisation time from this condition did not improve the amount of sol fraction of both PVA and heparin macromers. While PVA required shorter polymerisation time (3 min), the addition of the time needed to crosslink heparin (6 min) had no negative impact on PVA sol fraction and thus the same intensity and curing time could be used later on in PVA/heparin copolymerisation, which will be covered in Chapter 5.

3.4 CONCLUSION

The copolymerisation of two macromers by photocrosslinking requires the macromers to carry functional groups of similar reactivity. The choice of synthesis methods that can be used to functionalise both PVA and heparin is limited, due to the non-solubility of heparin in organic solvent. PVA was functionalised using the well- known ICEMA synthesis and by GMA synthesis, which could be performed in aqueous media. Properties of ICEMA-synthesised PVA macromers and hydrogels were used as a reference for the tested GMA method. Although the GMA method had low methacrylate incorporation efficiency, PVA hydrogels from GMA and ICEMA macromers were found to be comparable in structure and strength, with minimal cell growth inhibition. In addition to producing hydrogels that had the same properties as

ICEMA hydrogels, GMA synthesis was also successfully adapted to modify heparin.

Therefore, GMA-synthesised PVA macromers were chosen for the impending copolymerisation with heparin macromers, with both macromers carrying methacrylate groups of the equal reactivity.

- 67 -

Chapter 4

Methacrylate–Modified Heparin for Hydrogel Incorporation

eparan sulfate (HS), a glycosaminoglycan polysaccharide present in the Hbasement membrane and on the cell surface, binds growth factors and cytokines and enhances the signalling of these ligands by forming complexes with their receptors.

In this study, heparin was used as a model of HS and was modified with methacrylate functional groups to facilitate co-polymerisation with poly(vinyl alcohol) (PVA). Co- polymerising heparin and PVA was aimed at imparting the growth factor activation property of heparin to the synthetic PVA scaffold, therefore it was important for heparin to retain its bioactivity following chemical functionalisation. It was shown that the methacrylate group attachment on heparin did not result in the fragmentation of heparin molecules, and that the biological activity of the methacrylated heparin was preserved as determined by tests on its anticoagulation properties and ability to signal fibroblast growth factor-2 (FGF-2).

- 68 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation

4.1 BACKGROUND AND AIMS

Heparan sulfate (HS) interacts with proteins in a structurally specific manner, meaning only a certain sequence of the saccharide units can act as a specific binding site for a certain type of protein [56, 177]. Most of the work investigating the relationships of HS with proteins has used heparin as a model of HS. HS and heparin have the same basic disaccharide units of glucosamine and hexuronic acid, with heparin resembling a more sulfated version of HS [133-134]. Consequently, heparin and HS share similar functions, from protecting proteins from proteolytic degradation to regulating growth factor action in angiogenesis [128, 132].

In its own right, heparin has been used for decades as an anticoagulant in treating thrombotic diseases [178]. Heparin binds a plasma cofactor, antithrombin III (ATIII), converting the protein from a slow to a rapid inhibitor of thrombin, a process that delays or stops the activation of the clotting ‘factors’ in the coagulation cascade. ATIII inhibits the serine proteases that include coagulation factors IIa (thrombin) and Xa, thus preventing fibrin formation. The binding of ATIII to heparin is facilitated by a specific pentasaccharide sequence in the heparin chain (Scheme 4.1(A)) [136].

A O CH OSO - 2CH OSO3- C OH 2CH OSO3- 2 3 O O O O O OH O O O OH O OSO - O O OH OH 3 OH - OSO NH SO - HN OH NH SO3- 3 3 O B O CH OSO - 2CH OSO3- C OH 2CH OSO3- 2 3 O O O O O OH O O O O OSO - O O OH OH 3 OH - OH OH OSO NH SO - NH NH SO3- 3 3 O

Scheme 4.1 The specific heparin pentasaccharide sequence for ATIII binding. Structure of the active sequence (A) and after periodate-oxidation (B).

- 69 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation

Disruption of the pentasaccharide sequence, for example by enzymatic degradation of the heparin chain [179], has been shown to result in the loss of the anticoagulant activity of the heparin. Moreover, when used for purposes other than as an anticoagulant, heparin has been chemically modified to reduce its strong anticoagulant activity [180-181], in order to prevent bleeding complications when administered in high doses. For example, the structure of a glucosamine ring in the ATIII binding region

- was found to be altered following periodate (IO4 ) oxidation (Scheme 4.1(B)) [181]. The resulting IO4-heparin was unable to bind to ATIII, amounting to a lower anticoagulation activity.

A commonly used method to detect blood coagulation abnormalities is the activated partial thromboplastin time (aPTT) assay, which measure clotting time of plasma, in the presence of phospholipids as platelet substitutes, calcium chloride, and more recently kaolin [182]. This assay has been used extensively as a method to discover the coagulation factors in plasma that are involved in the coagulation cascade.

Aside from ATIII binding capacity, heparin also binds a protein called the plasma co- factor II, which would also deactivate thrombin and prevent fibrin formation. Plasma co-factor II has lower binding strength to heparin compared to ATIII [183]. This indicates that when both ATIII and plasma co-factor II are present, most heparin would bind ATIII and therefore the anticoagulant activity mainly depends on the activation of

ATIII.

Looking back at the purpose of including heparin in the PVA scaffold, the property of heparin that was of our interest was its growth factor signalling activity. One of the major functions of HS/heparin in vivo is in regulating the activity of various growth factors [150]. Members of the fibroblast growth factor (FGF) family, in particular FGF-2, have been shown to bind specifically to heparin, and are referred to as

- 70 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation heparin-binding growth factors. HS and heparin interact with FGFs and their receptors

(FGFRs) to form FGF-HS-FGFR signalling complexes [101]. The minimal binding sequence for FGF-2 on HS has been shown to be a hexasaccharide [184], however FGF-

2 activation requires a decasaccharide, the longer chain length being necessary to bridge the physical distance between the growth factor and its associated cell receptor [56,

185].

In this work, the purpose of modifying heparin with functional groups was to covalently attach and introduce heparin as biomolecular cues within a hydrogel scaffold, by copolymerisation with PVA using a UV-initiated polymerisation technique. The cytocompatibility of the covalently attached methacrylate groups has been assessed in

Chapter 3. Unlike the modification of synthetic polymers, chemical modifications can denature biological molecules. Therefore it was important to ensure that the methacrylate modification and UV exposure were not detrimental to the integrity of heparin. The aim of this Chapter was to address the effect that the functionalisation process have on the structure and activity of heparin. The molecular weight profiles of heparin prior to and after methacrylation were compared, as well as their anticoagulant and growth factor-binding properties.

4.2 EXPERIMENTAL

4.2.1 Materials

Methacrylated heparin (hep-MA) with an average of ~3 methacrylate groups per chain was synthesised as outlined in Chapter 3. Toyopearl® HW-50S media (Tosoh

Bioscience) for size exclusion chromatography was washed prior to packing into a

Pharmacia HR10/30 column. Heparin samples (heparin sodium salt from porcine intestinal mucosa) of different molecular weights were used as standards: heparin (grade

- 71 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation

I-A, average MW 17-19 kDa, Sigma), low molecular weight (LMW) heparin (average

MW 4-6 kDa, Sigma), and Deligoparin™ (average MW 2-3 kDa, Opocrin, Italy). Fresh frozen human plasma (Australian Red Cross Blood Service), phospholin ES activated partial thromboplastin time (APTT) reagent (ellagic acid based reagent with soybean phospholipids) (R2 Diagnostics, Inc.), and recombinant human fibroblast growth factor-

2 (FGF-2) (Invitrogen) were used without further purifications. Dyes 1,9- dimethylmethylene blue (DMMB) (Sigma), AlamarBlue (Invitrogen), and CellTiter 96®

AQueous One Solution Reagent ([3-(4,5-dimethylthiazol-2-yl)-5-(3- carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium, inner salt (MTS))

(Promega) were also used as received.

4.2.2 Size Exclusion Chromatography

Size exclusion chromatography (SEC) was performed in a HR10/30 column packed with Toyopearl® HW-50S to obtain molecular weight distribution profile of methacrylated heparin (hep-MA) and UV-exposed heparin (UV-hep), using a method described by Melrose and Ghosh [186]. Hep-MA and heparin standards were dissolved in buffer at 2 mg/mL, of which ~0.4 mL was injected into the column. To prepare UV- hep, 2 mg/mL heparin was prepared in buffer and irradiated with 0.3 W/cm2 UV for 30 minutes prior to injection. 0.25 mL eluted fractions were collected from the column and the heparin content was analysed by DMMB assay. 50 µL aliquots of each fraction were transferred to a 96-well plate, after which 200 µL DMMB solution was added to each well, followed by absorbance reading at 535 nm. The absorbance properties of the

DMMB dye will be altered when it binds heparin [187], and the amount of heparin was quantified by measuring the change in absorbance against a standard curve of absorbance vs. heparin concentration.

- 72 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation

The profile was then compared with those of the following heparin standards: non-modified heparin (17-19 kDa) and two low molecular weight heparin samples (4-6 kDa and 2-3 kDa). The retention time of the molecules in the column indicates their molecular weight, and was represented by the partition coefficient, Kav. The Kav was calculated by:

Ve −V0 )( K av = (4.1) t −VV 0 )( where Ve = elution volume of the heparin molecule, V0 = void volume of the column, and Vt = total volume of the column.

4.2.3 Clotting Time Assay

Heparin and hep-MA was dissolved in 0.9% NaCl solutions at concentrations ranging from 0.5 to 5 µg/mL. 180 µL human plasma was mixed with 20 µL heparin or hep-MA solution and warmed at 37ºC for 60 seconds. 200 µL pre-warmed phospholin

ES APTT reagent was added, mixed and further incubated at 37ºC for 3-5 minutes.

Coagulation was initiated by adding 200 µL pre-warmed CaCl2, at which point the timing began. Clotting time was recorded when the first sign of fibrin formation was observed. As a control, the clotting time of pure human plasma was measured.

4.2.4 BaF3 Cell Proliferation Assay

The maintenance of BaF3 FR1c cells, a heparan sulfate proteoglycan (HSPG)- deficient myeloid cell line expressing the FGF receptor 1c (FGFR1c) isoform, and the mitogenic assay were essentially performed as described by Knox, et al. [137]. Prior to starting the assay, the BaF3 cells were washed with and transferred to RPMI 1640 medium containing 10% FBS, and incubated at 37°C for approximately 15 h. Cells were

- 73 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation then washed and resuspended at 1×105 cells/mL with 3 nM FGF-2 and the appropriate heparin or hep-MA concentrations. The cell suspension (200 µL) was seeded into 96- well plates and incubated at 37°C for 72 h. Afterward 20 µL AlamarBlue was added into each well, incubated for 6 h, before reading the plates for absorbance at 570 and

600 nm (VersaMax Tunable Microplate Reader, Molecular Devices).

Cell proliferation was assayed by measuring the degree of AlamarBlue reduction, which corresponds to the cell number and is calculated by:

117,216× A − 80,586 × A %ABreduction = 570 600 ×100 (4.2) 155,677× AB600 − 14,652 × AB570 where A570 = absorbance of test wells at 570nm, A600 = absorbance of test wells at

600nm, AB570 = absorbance of blank wells (medium without cells) at 570nm, and AB600

= absorbance of blank wells at 600nm.

The data was normalised following the method by Padera, et al. [100], in which cell proliferation index was calculated by dividing the increase in cell number for each test condition by the increase in cell number for the positive control. The positive control was BaF3 cells incubated with 3 nM FGF-2 and 30 nM heparin.

Statistical analysis of cell proliferation results. To compare the effects of heparin and hep-MA on the BaF3 cell proliferation, the logarithm of the proliferation index was analysed with an analysis of variance model [165] with two fixed factors (heparin type and heparin concentration) and one random factor (experiment) using Minitab 15 [166].

- 74 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation

4.3 RESULTS AND DISCUSSION

4.3.1 Structural Analysis of Methacrylated Heparin

Heparin was modified with methacrylate pendant groups, and in Chapter 3 the methacrylated heparin (hep-MA) with approximately 3 MA groups per chain was able to form pure heparin hydrogels by photopolymerisation. Functionalisation of biomolecules such as glycosaminoglycans to form hydrogels [54, 86] or for covalent incorporation into synthetic polymers [9, 74] have been reported for various purposes, however bioactivities of these molecules following chemical modification have not been closely examined. As the purpose of incorporating heparin into the scaffold was to impart bioactivity, it was sensible to examine whether the modification altered its bioactivity.

The structural integrity of heparin was confirmed by SEC to determine if the hep-

MA macromer chain was cleaved following the functionalisation process. Heparin and hep-MA were eluted through a SEC column to obtain their molecular weight profiles.

Non-modified heparin and two low molecular weight heparin samples (4-6 kDa and 2-3 kDa) were used as standards. The SEC result showed no shift in the molecular weight distribution profile of heparin after methacrylate addition compared to that of non- modified heparin (Figure 4.1), indicating that the modification did not result in the fragmentation of heparin molecules.

- 75 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation

0.4

0.3 535

0.2

Absorbance,A 0.1

0 0 0.1 0.2 0.3 0.4 0.5 0.6

Partition coefficient, Kav

Figure 4.1 Molecular weight distribution profiles of hep-MA and heparin standards. Molecular weight distribution profile of hep-MA (○) and UV-hep (×) compared to profiles of non- modified heparin (17-19 kDa,
) and two low molecular weight heparins of 4-6 kDa (■) and 2-3 kDa (▲). Amount of eluted heparin is represented by the absorbance value of the colorimetric DMMB dye at 535 nm.

Extended exposure (30 min) of heparin to UV at 0.3 W/cm2 also did not fragment the heparin chain, even though the applied UV intensity was ten times the intensity ultimately used to crosslink the heparin macromers. In Figure 4.1 above, the amount, i.e. the absorbance, of UV-hep detected appeared higher compared to that of heparin.

This might have been caused by a small difference in the volume of UV-hep solution that was injected into the column, causing an increase in the amount detected by

DMMB without necessarily shifting the peak.

For macromer chains to form hydrogels by covalent crosslinking, they need to be decorated with at least two functional groups per chain. Fragmentation of the heparin chain would result in smaller chains that might contain little or no MA- functionalisation, which consequently may not be incorporated into the hydrogel.

Fragmentation would also change the way the smaller heparin chains interact with proteins. Depending on the size and the sequence that is available in the fragments, less protein interactions might occur due to the disruption of binding sites. An enhancement

- 76 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation in activity, as has been seen in some growth factor signalling studies with low molecular weight heparin, was also a possibility. However, the main concern was for heparin to be functionalised with sufficient MA functional groups for covalent crosslinking. This result confirmed that the reaction with glycidyl methacrylate (GMA) was safe for the integrity of the heparin chains.

4.3.2 Antithrombin III Activation

As outlined in the previous Chapter, methacrylate groups attachment on heparin can be done by substituting the hydroxyl, carboxyl or O-sulfate groups on the heparin chain. Even when the length of the heparin chain remained unchanged, the biological functionality of the resulting hep-MA might be compromised depending on where and how many of those substitutions occurred. Oxidation/ reduction reactions as well as variations in the amount of sulfation have been shown to alter the activity of HS/heparin towards various proteins [56, 94]. Figure 4.2 illustrates the importance of specific sequences on the heparin chain for the binding and activation of proteins.

Heparin backbone Protein Protein binding sequence Activated protein MA pendant group

Figure 4.2 Illustration of the binding of proteins to specific sequences on heparin chain. Attachment of methacrylate (MA) pendant groups would alter the structure of these binding sequences and inhibit the binding and activation of proteins.

As a first step in confirming the activity of heparin following MA group attachments, the anticoagulation property of hep-MA was assessed using an activated

- 77 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation partial thromboplastin time (aPTT) assay and the result compared to that of heparin. The aPTT assay had been used to measure the anticoagulant properties of heparin-coated surfaces, and also to test the thrombogenicity of various blood-contacting medical materials [136, 188-189]. In this work, the clotting times of plasma incubated with heparin and hep-MA were measured and represented as a function of heparin or hep-

MA concentrations (Figure 4.3).

600

500

400

300

200 Clotting time (s)

100

0 0 1 2 3 4 5 6 Heparin concentration in plasma (µµµg/mL)

Figure 4.3 Plasma clotting time as a function of heparin and hep-MA concentrations. Anticoagulation properties of heparin (●) and hep-MA (○) measured in terms of clotting time as a function of heparin concentration in plasma. The clotting time of plasma with no sample solutions introduced was 72 s.

It was found that without heparin, plasma clotted in approximately 72 seconds. It has been well documented that the addition of heparin increases the clotting time [178].

As expected, increasing heparin concentration delayed the clot formation, and the same was observed with hep-MA. For both heparin and hep-MA, the clotting time was sharply increased at concentrations higher than 5 µg/mL, deviating from the trend at the lower concentrations. This result suggests that the MA-modification had minimal effect on the ability of heparin to delay clot formation. The heparin used in this study (17-19

- 78 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation kDa) consisted of approximately 75-85 saccharide units, and after modification had on average 3 MA groups randomly attached along the chain. It was concluded that this amount of MA substitution was not enough to substantially alter the anticoagulation, and that the pentasaccharide sequences for ATIII binding were mostly unmodified and still active.

4.3.3 Cellular Signalling of FGF-2

In a study of the periodate-oxidated heparin [181], earlier mentioned in the introduction of this Chapter (Scheme 4.1), the anticoagulation activity was decreased when the structure of the pentasaccharide sequence was altered. At the same time, the binding affinity to FGF-2 and the heparanase inhibitory activity of the IO4-heparin were found to be largely unchanged [95]. These results mean that structure modification of heparin may have minimal effect on some of its activity, but can result in the loss of other functions.

The growth factor activation property of the modified heparin was of special interest in this work, for which different binding sequences than that for ATII binding would be required. FGF-2 interacts with a specific hexasaccharide sequence that contains a 2-O-sulfated iduronic acid (IdoUA(2S)) and an N-sulfated glucosamine

(GlcNS) [184, 190]. For receptor signaling, a dodecasaccharide containing this binding region as well as 6-O-sulfated GlcNS residues is also required [139]. This means although not of the same structure, the signalling of FGF-2 to cells requires twice the saccharide length of that required for ATIII activation.

Most mammalian cells synthesise HS in vivo, which was then distributed on cell surface and in the extracellular matrix (ECM). Cell-surface HSPG helps anchor growth factors for binding with its cell surface receptor [135, 144]. Although the exact role of

- 79 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation

HS/heparin in enhancing the binding and signalling of FGF-2 remains to be confirmed, with different conformations of the heparin/FGF/FGFR proposed [101], HS/heparin has been shown to be essential for the cell signalling of FGF-2. Addition of heparin was shown to improve growth in FGF-responsive cells that were depleted of cell surface

HSPGs. Studies have shown the use of chlorate on Swiss 3T3 fibroblasts [139] and transfection of B-lymphocytes to impair their ability to synthesise HSPG [137, 145].

To further assess the bioactivity of the methacrylated heparin, a cell proliferation study was performed to investigate its FGF-2 signalling activity. Mouse B lymphocytes

(BaF3), which were transfected to express the cell surface receptor of FGF-2 (FGFR 1c isoform), were used [191-192]. The BaF3 cells were deficient of HSPG on their cell surfaces, therefore eliminating the chance of competitive binding between the cells’ own HS and the heparin being tested. The BaF3 cells were dependent on interleukin-3

(IL-3) for proliferation, but in the absence of the cytokine cell growth can be regulated by FGF-2 (Figure 4.4). In the interest of studying cellular signalling of FGF-2, the cells were deprived of IL-3 prior to the start of all mitogenic assays.

Figure 4.4 Illustration of a BaF3 cell and its cell surface receptors. Cell proliferation is dependent on IL-3, in the absence of which can be substituted by FGF-2. Figure courtesy of S. Yogisaputra.

A series of 3-d proliferation studies on the BaF3 cells was done using FGF-2 and heparin of varying concentrations. The experimental results were found to agree with a

- 80 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation theoretical model (Equation 4.3) that was developed by Padera, et al. [100] based on the relationships between FGF-2, FGFR1 and heparin:

0.60+ 0.25H y = 0 (4.3) 1 0.93+ 0.25H0 + F0 where y is the proliferation index, H0 is the initial heparin concentration, and F0 is the initial FGF-2 concentration.

The BaF3 cells did not proliferate in the absence of FGF-2 (Figure 4.5(A)), confirming their dependency on the factor for growth. When incubated in 3 nM FGF-2, the degree of cell proliferation continued to increase with heparin concentration, up to

30 nM heparin. After this point, any further increase in heparin concentration did not result in improved proliferation. The FGF-2 mediated cellular response on the BaF3 cells has been shown to be a function of both FGF-2 and HS/heparin concentrations

[145], and a reduction of one of the components can be compensated for by an increase in the other.

Figure 4.5(B) shows that while FGF-2 can signal in the absence of heparin, the presence of heparin enhances the signal, effectively reducing the amount of FGF-2 required for growth. For example, to get a proliferation index of ~0.5, less than 0.1 nM

FGF-2 was needed when 30 nM heparin was present while 3 nM FGF-2 was needed when no heparin was added to the culture medium. No substantial increase in proliferation was observed after the FGF-2 reached 3 nM (Figure 4.5(B)). Therefore 3 nM FGF-2 and 30 nM heparin was used as the positive control for the culture conditions.

- 81 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation

1.2

1.0 A

0.8

0.6

0.4 Proliferationindex 0.2

0.0 0.01 0.1 1 10 100 Heparin concentration (nM)

1.2 B 1.0

0.8

0.6

0.4 Proliferationindex 0.2

0.0 0.001 0.01 0.1 1 10 100 FGF2 concentration (nM)

Figure 4.5 BaF3 cell proliferation as a function of heparin (A) and FGF-2 (B) concentrations. (A) response shown with no FGF-2 () and with 3 nM FGF-2 (●) added, after 72 h incubation; (B) response shown with no heparin (
) and with 30 nM heparin (■) added, after 72 h incubation. The lines represent the theoretical proliferation index calculated from the model described in Equation 4.3.

To compare the mitogenic activities of heparin prior to and after methacrylation, the BaF3 cells were incubated with 3 nM FGF-2 and a range of heparin and hep-MA concentrations. The cells incubated with hep-MA showed similar trends in proliferation compared to those cultured with the non-modified heparin (Figure 4.6). There was substantial variation between repeat experiments (n=3), as indicated by the error bars.

The proliferation index was generally higher for native heparin, however the difference between cell responses to heparin and hep-MA was not statistically significant (p=0.24).

- 82 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation

2

1

0.5 Proliferation index 0.2

0.1 0.01 0.1 1 10 100 Heparin concentration (nM)

Figure 4.6 Comparison of FGF-2 signalling activity of heparin and hep-MA. The FGF-2 signalling activity was measured in terms of BaF3 cell proliferation as a function of heparin (●) and hep-MA (○) concentrations, in the presence of 3 nM FGF-2, after 72 h incubation.

It can be contemplated that the MA substitution had more effect on the cellular signalling of FGF-2 than on anticoagulation, due to the longer saccharide sequence on heparin chain that is needed to activate FGF-2 than ATIII. Nevertheless, the results from the two assays also suggest that the addition of ~3 methacrylate groups per heparin chain allowed heparin to remain active, while enabling it to be covalently incorporated into PVA.

4.4 CONCLUSION

In this work, heparin was modified with methacrylate pendant groups, which enabled it to be covalently attached to a synthetic polymer. Heparin-binding proteins are only able to attach to their own unique binding sites on the heparin chain. Disruptions of these binding sites, by fragmentation or structure alteration, would result in a loss of protein activation. Heparin was successfully modified with photopolymerisable groups which would allow it to be co-polymerised with PVA to make a synthetic/biological co- polymer hydrogel construct. The modification of the heparin did not result in the

- 83 -

Chapter 4 – Methacrylate–Modified Heparin for Hydrogel Incorporation fragmentation of molecules, or the alteration of its anticoagulation activity. FGF-2 signaling also confirmed that the hep-MA molecules performed similarily to the unmodified heparin. Therefore, it can be concluded that this method of functionalising the heparin is a viable route, and will allow for the successful incorporation of biological cues into the PVA scaffolds.

ACKNOWLEDGEMENTS

The author would like to acknowledge and thank Stella Yogisaputra for the illustration in Figure 4.4.

- 84 -

Chapter 5

Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

ynthetic scaffolds show great promise for use in tissue engineering due to their Sability to mimic some aspects of the extracellular matrix, however, their use has been hindered by the lack of inherent recognition sites that are required for protein and cell interactions. The study outlined in this chapter focuses on the formation of photopolymerised hydrogels derived from methacrylated macromers of poly(vinyl alcohol) (PVA) and heparin, with the aim of imparting the growth factor activation property of heparin to the synthetic scaffolds. The addition of heparin into the PVA hydrogels resulted in an increase in mass swelling ratio from 5.8 for pure PVA to 6.5 and 6.6 for PVA/heparin co-hydrogels of 19/1 and 17.5/2.5 (w/w) compositions, respectively. The aim of this work was to add heparin molecules into a synthetic PVA scaffold without adversely affecting the structural and mechanical stability of the PVA scaffold. The tensile moduli of the co-hydrogels remained close to that of PVA hydrogels (61 kPa), even up to 2.5% heparin composition (PVA/hep 17.5/2.5). Finally, the co-hydrogels were found to retain the growth factor signalling activity of heparin at equilibrium.

- 85 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

5.1 BACKGROUND AND AIMS

The extracellular matrix (ECM) provides structural support for cells and acts as a reservoir for molecular components responsible for protein upregulation and the ensuing cellular activities [5, 7]. The way the molecular signals are presented to the cells is of utmost importance, especially in tissue engineering approaches and in the design of a scaffold capable of providing the necessary molecular signals to facilitate tissue regeneration. Hydrogels have been used as vehicles for the controlled release of these signalling molecules in vivo [6, 52, 113], and also as a supporting matrix for the sustained presentation of the molecules to surrounding tissues [9, 41, 49, 159, 193].

As discussed in the literature review in Chapter 2, vascularisation of tissue engineered constructs has been of great interest for some time. The key to cellular infiltration into the engineered constructs will rely on the interaction of cells with the biomaterial making up the constructs. Cell adhesion to biomaterials is mediated by interactions of the cell-surface receptor with proteins that are adsorbed onto the biomaterials. Poly(vinyl alcohol) (PVA), the base polymer used for hydrogel formation in this study, has demonstrated biocompatibility in several biomedical applications. Like other synthetic materials, however, the resistance of PVA hydrogels to protein adsorption and cell adhesion is a drawback for applications such as tissue engineering

[58]. Cell adhesion peptides have been introduced to improve the attachment of cells to these hydrogels [60], however subsequent processes such as cell growth and tissue formation are reliant on an array of growth factors.

Delivery and presentation of heparan sulfate (HS)/heparin from natural and synthetic hydrogels to enhance tissue and hydrogel vascularisation have been investigated. In its growth-factor binding capacity, heparin has been introduced to natural and synthetic polymers as physically bound or covalently crosslinked

- 86 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels biomolecules, particularly for growth factor delivery and activation [10, 41, 63, 120].

However earlier studies of heparin coupled to various polymers such as PVA aimed at using the anticoagulant property of heparin, to improve the antithrombogenicity of the polymer surfaces [118, 156]. Fabrication of hydrogels based on heparin [194] and other glycosaminoglycans, including hyaluronic acid (HA) [82, 87] and chondroitin sulfate

(CS) [8, 99], has more recently been explored.

Three main aspects of the formation and characterisation of PVA and heparin hydrogels are addressed in this Chapter. The first focused on the suitable polymerisation conditions for PVA and heparin. As outlined in Chapter 3, various pendant groups were introduced to the PVA and heparin chain to create crosslinkable PVA and heparin macromers. Photo-initiated radical polymerisation of these functional groups allows rapid hydrogel formation in the presence of non-toxic and water soluble photoinitiator

[106], which fulfil the need for mild polymerisation conditions that are relatively compatible with the cultured cells and target applications.

The next focus concerned the structural and mechanical stability of the

PVA/heparin co-hydrogels. As with any copolymerisation, the combination of two compounds of different properties inevitably changes the original properties of the individual compounds. The copolymerisation of PVA with both synthetic and biological polymers has increasingly been reported in the literature [106, 118, 123-124].

In separate studies, poly(ethylene glycol) (PEG) [111] and chondroitin sulfate (CS) [99,

123] have been copolymerised with PVA to create scaffolds for cartilage regeneration.

The studies showed the use of composite PVA hydrogels for the encapsulation of chondrocytes, with the goal of delivering the cells to an injury site to initiate cartilage formation. Results from these studies demonstrate that the properties of the resulting hydrogels could be tailored by adjusting the ratio of copolymers. The degradation time

- 87 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels for PVA/PEG co-hydrogels increased with higher PEG to PVA ratio, ranging from less than 1 d for pure PVA hydrogels to 34 d for PEG hydrogels [111] The mass swelling ratio (q) and compressive modulus (K) of a 20 wt% PVA hydrogels (q ~3.2, K ~680 kPa) was observed to increase up to a q of ~9.0 and a K of ~900 kPa, the properties of a

20 wt% CS hydrogels [99]. Finding the optimum copolymerisation conditions is necessary to ensure not only the combination but also preservation of the advantageous properties of the individual polymer component.

The last aspect to be investigated, which ultimately is also the most important, is the availability and bioactivity of the heparin in the co-hydrogels. Heparinised hydrogel scaffolds have been analysed for the binding and controlled release of growth factors [8-

10], and used to support cell adhesion and differentiation [11-12]. Heparin has been shown to retain its ability to bind proteins when copolymerised with PEG hydrogels, and was found to support osteogenic differentiation of human mesenchymal stem cells

(hMSC) [75]. However, the specific function of covalently bound heparin as growth factor-activating molecules for cell growth has not been extensively studied, and there has also been a lack of reports investigating the effect of adding biological molecules on the mechanical stability of synthetic hydrogels.

Therefore, the aims of this Chapter were to address the effect of heparin incorporation into PVA as co-hydrogels, and to test the bioactivity of heparin in the resulting matrix. Photopolymerised hydrogels were formed from the methacrylated macromers of PVA and heparin. The optimum photopolymerisation conditions for the two macromers were analysed, and the efficiency of hydrogel crosslinking was characterised by measuring the sol fraction of the hydrogels. The effect of heparin addition to the structural and mechanical stability of PVA was investigated by measuring changes in mass loss, swelling capacity and tensile properties. The

- 88 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels bioavailability of the incorporated heparin was examined by performing a plasma clotting assay and a BaF3 cell proliferation study.

5.2 EXPERIMENTAL

5.2.1 Materials

PVA (~3 crosslinkers per chain) and heparin (~3 crosslinkers per chain) macromer were synthesised by reaction with glycidyl methacrylate and purified as described in Chapter 3. The starting materials were obtained from Sigma: PVA (average

MW 13-23 kDa, 98% hydrolysed), heparin sodium salt (grade I-A, from porcine intestinal mucosa, average MW 17-19 kDa) and glycidyl methacrylate (97% purity).

The photoinitiator, 2-hydroxy-1-[4-(hydroxyethoxy)phenyl]-2-methyl-1-propanone

(Irgacure 2959, Ciba Specialty Chemicals), was used as supplied. Silicone sheets

(Silastic® Sheeting, reinforced medical grade silicone rubber, Dow Corning) were perforated for use as hydrogel moulds. Dyes 1,9-dimethylmethylene blue (DMMB)

® (Sigma), AlamarBlue (Invitrogen), and CellTiter 96 AQueous One Solution Reagent ([3-

(4,5-dimethylthiazol-2-yl)- 5- (3-carboxymethoxyphenyl)- 2- (4-sulfophenyl)- 2H- tetrazolium, inner salt (MTS)) (Promega) were also used as received. The cell culture media RPMI-1640 and M199 were obtained from Sigma, prepared following the manufacturer’s instructions, and sterile filtered (0.2 µm). Recombinant human fibroblast growth factor-2 (FGF-2) (Invitrogen) was used without further purifications.

5.2.2 Hydrogel Formation

All hydrogels were formulated with 20% (w/w) total macromer concentration.

Methacrylated PVA (PVA-MA), methacrylated heparin (hep-MA), or a combination of the two macromers was first dissolved in water. Heating at 80°C for at least 2 h was

- 89 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels required to completely dissolve PVA-MA. To prepare the co-hydrogels, PVA-MA and hep-MA were dissolved separately before mixing at room temperature. The photoinitiator, Irgacure 2959, was added to make up a final concentration of 0.1% (w/w) initiator. The macromer solution was placed in a mould and photopolymerised with a

UV light source (UVA 300-480nm, 365nm peak, Green Spot) at an intensity of 30 mW/cm2 for 6 minutes.

5.2.3 Hydrogel Characterisation

5.2.3.1 Swelling study and mass loss analysis

Swelling and mass loss studies were carried out as described in Chapter 3. In brief, hydrogel discs (10mm diameter × 1mm thick) of PVA, heparin and their copolymers were made. A t0 sample was taken and lyophilised without immersion in buffer, while the rest of the samples were incubated in PBS (pH 7.4) at 37ºC and taken out at various time points up to 7 d. Samples were made in triplicates for each type of hydrogel and for each time point that they were collected. By obtaining the initial (m0), swollen (ms), and final dry (md) weights of the samples, the initial dry weight (mid), mass swelling ratio

(q) and mass loss of the hydrogels were calculated following equations derived in

Chapter 3. Below is a summary of the equations used to characterise the swelling and mass loss of the hydrogels.

m macromer fraction = d,t0 (5.1) m 0,t0

The hydrogels’ initial dry weights (mid).

mid = m 0 × macromer fraction (5.2)

m q = s (5.3) md

- 90 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

m - m mass loss =id d ×100% (5.4) mid

Again, the hydrogel sol fraction refers to the portion of macromers that are not incorporated into the hydrogels. It has previously been reported to be released within 24 h of swelling, and therefore equals the mass loss of hydrogels at 24 h [30, 112].

5.2.3.2 Composition of co-hydrogel mass loss

PVA/heparin 17.5/2.5 co-hydrogel discs were formed and their mass loss determined as above. At the end of each incubation time, the sink volume was sampled and diluted. 50 µL aliquots of the diluted samples were transferred to a 96-well plate, after which 200 µL DMMB solution was added to each well, followed by absorbance reading at 535 nm (VersaMax Tunable Microplate Reader, Molecular Devices). The absorbance properties of the DMMB dye will be altered when it binds heparin [187], therefore the amount of heparin lost from the co-hydrogel into the sink solution was quantified by measuring the change in absorbance against a standard curve of absorbance vs. heparin concentration. Water was removed from the remaining sink volume by freeze-drying to leave behind dry powder, which was sampled and analysed

1 by H NMR in D2O to confirm the presence of PVA and heparin in the co-hydrogel mass loss. Knowing the amount of heparin from DMMB assay (mhep,loss) and coupled with the mass loss data, the amount of PVA (mPVA,loss) in the co-hydrogel mass loss was then calculated:

lossPVA lossPVA did −−= m)mm(m hep loss (5.5) where mid and md were the initial and final dry weights of the co-hydrogel, respectively.

- 91 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

For comparison, the theoretical amounts of heparin and PVA in the co-hydrogel mass loss were determined. The initial dry weight of heparin in the co-hydrogel (mid,hep) was first obtained by

mid,hep = % heparin in copolymer× mid (5.6)

Using the mass loss of pure heparin hydrogels (mass losshep), the theoretical amount of heparin in the co-hydrogel mass loss (mhep,theory) was estimated by

m hep theoryloss, m hepid, ×= mass loss hep (5.7)

Similarly, the theoretical amount of PVA in the co-hydrogel mass loss (mPVA,theory) was calculated:

m theoryloss,PVA m PVAid, ×= mass loss PVA (5.8) where mass lossPVA refers to the mass loss of pure PVA hydrogels.

5.2.3.3 Final composition of co-hydrogels at equilibrium

The actual amounts of PVA and heparin that remained crosslinked in the co- hydrogel, mPVA and mhep, respectively, were determined by substituting the initial dry weight of the PVA or heparin, by the respective amount lost.

PVA PVAid, −= mmm lossPVA (5.5)

hep hepid, −= mmm hep loss (5.6)

By determining the final macromer fraction of the co-hydrogel at equilibrium using the calculated mass loss and initial macromer fraction,

macromer fraction final = − mass1( loss) × macromer fraction (5.7) the final composition of the PVA and heparin in the co-hydrogel was then calculated by:

m PVA PVA eq = × m acromerfraction final (5.8) PVA + mm hep

- 92 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

m hep hepeq = × m acromerfraction final (5.9) PVA + mm hep

5.2.3.3 Mechanical testing

PVA and co-hydrogel samples were photopolymerised in a dumbbell-shaped mould, with the same macromer and initiator concentrations as described in the previous section. The testing area of the samples was 10 mm long, 2 mm wide and 1 mm thick. Immediately after polymerisation, the sol fraction was extracted by incubating in PBS for 24 h to ensure all non-crosslinked macromers were removed [30,

112]. Samples were mounted onto an Instron 5543 universal testing machine, with fine sandpaper covering the grips to reduce slippage. Tensile tests were performed at a 3- mm/min strain rate until failure.

5.2.4 Bioactivity of PVA/Heparin Co-Hydrogels

5.2.4.1 Antithrombin III activation

PVA and co-hydrogel discs were formed and sol fraction-extracted for 24 h in

PBS. Each hydrogel was placed in 200 µL human plasma and warmed at 37ºC for ~60 seconds. 200 µL pre-warmed phospholin ES aPTT reagent was added, mixed and further incubated at 37ºC for 3-5 minutes. Coagulation was initiated by adding 200 µL pre-warmed CaCl2, at which point the timing began. Clotting time was recorded when the first sign of fibrin formation was observed. As controls, heparin solutions prepared in 0.9% NaCl were introduced to plasma, with concentrations ranging from 0 to 5

µg/mL plasma. The clotting time of pure plasma was also measured.

- 93 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

5.2.4.2 Cellular signalling of FGF-2

PVA and co-hydrogel discs were formed in sterile conditions, and their sol fraction was extracted by incubation in sterile DPBS for 24 h. Each hydrogel was then transferred to a well in a 24-well plate and BaF3 cells were seeded on top of the gels at

6×104 cells/well. Positive controls were BaF3 cells incubated in RPMI-1640 media containing 0 to 30 nM heparin. Culture medium for all controls and test hydrogels contain 0.3 nM FGF-2, and the cells were incubated at 37°C for 72 h. MTS reagent was then added at 10% (v/v) and the cells were incubated for a further 6 h before the culture medium was measured for absorbance at 495 nm (Tecan Infinite F200 Microplate

Reader) to assess cell proliferation.

5.3 RESULTS AND DISCUSSION

5.3.1 Hydrogel Characterisation

5.3.1.1 Swelling study and mass loss analysis

PVA and heparin macromers, each with an average of 3 MA groups/chain, were first photopolymerised to form individual PVA and heparin hydrogels. The optimum photopolymerisation condition for both the PVA-MA and heparin-MA was previously determined in Chapter 3. All hydrogels were formed by UV-induced crosslinking at an intensity of 30 mW/cm2 for 6 min.

To investigate the covalent incorporation of hep-MA into the hydrogels, non- modified heparin was mixed and polymerised with PVA-MA using the same ratio as hydrogels formed with hep-MA and PVA-MA (PVA/heparin 17.5/2.5). The hydrogels were immersed in PBS, collected at various time points, and their supernatants analysed for their heparin content by a DMMB assay. Figure 5.1 shows the amount of heparin

- 94 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels released from the two types of PVA/heparin hydrogels, one crosslinked with hep-MA, the other simply containing non-modified heparin.

100

80

60

40 Heparin loss from

PVA/heparin hydrogels (%)PVA/heparin hydrogels 20

0 0 1 2 3 4 5 6 7 Incubation time (d) Figure 5.1 Comparison of heparin loss from PVA/heparin 17.5/2.5 hydrogels when heparin was covalently linked and when physically blended. Heparin was functionalised (●) (heparin-MA, 3 crosslinker/chain), or non-functionalised (○).

The non-modified heparin was released at a higher amount than hep-MA, and unlike the hep-MA, it continued to be released from the PVA hydrogels after the initial sol fraction extraction, reaching ~90% loss at 7 d. There was no further release of hep-

MA after a 1-d incubation, indicating covalent crosslinking of the heparin into the hydrogel network. Around 50% of the originally incorporated hep-MA remained within the hydrogels at equilibrium. Heparin stability in the co-hydrogel network is particularly attractive for long-term applications, where continuous presence of biological activity within the scaffold is needed.

Once the copolymerisation of PVA and heparin was confirmed, PVA/heparin co- hydrogels were formed at various compositions. The swelling behaviour and mass loss of the co-hydrogels were measured and compared with the pure PVA and heparin hydrogel. Figure 5.2 shows the swelling and mass loss profiles of the PVA, heparin and

- 95 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels the co-hydrogels over a period of 7 d. Sol fractions of the PVA and heparin hydrogels were rapidly released within the first few hours of incubation, and equilibrium was considered to be reached by 24 h [30, 112]. The swelling profiles of the hydrogels were also found to stabilise after 24 h. There was no increase in mass loss following the initial 24-h incubation period, and after 7 d a typical 20 wt% PVA hydrogel had 25% mass loss with a mass swelling ratio of 5.8, while heparin hydrogels had 32% mass loss with a swelling ratio of 17.

22 50

20 40

30 18

20 16

8 10

6 massHydrogel loss (%) 0 Hydrogel massHydrogel swelling ratio, q 4 0 1 2 3 4 5 6 7 0 1 2 3 4 5 6 7 Incubation time (d) Incubation time (d)

Figure 5.2 Swelling and mass loss profiles of PVA, heparin, and their copolymer hydrogels. The profiles of PVA (■), heparin (●), and PVA/heparin 17.5/2.5 (
) hydrogels were plotted against incubation time and were found to stabilise after 1 d of incubation at 37ºC.

The relative sizes of the hydrogels following a 24-h incubation is shown in Figure

5.3. The original diameters of the hydrogels were 10 mm. At equilibrium swelling, the diameters of PVA and PVA/hep 19/1 hydrogels were higher, at ~10.5 and ~11 mm, respectively. The diameter of the heparin hydrogel, on the other hand, increased approximately by 25% to ~13 mm. Both PVA and heparin hydrogels were formulated with the same percent macromer (20 wt%), and had approximately the same crosslinker density, therefore the high swelling capacity of heparin gels was attributed to the ionic

- 96 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels nature of heparin due to the presence of sulfate groups. The equilibrium properties of the hydrogels are summarised in Table 5.1.

PVA PVA/hep 19/1 Heparin

Figure 5.3 PVA, heparin and PVA/heparin hydrogels at their equilibrium swelling. The hydrogels were equilibrated in PBS pH 7.4 for 24 h.

Table 5.1 Equilibrium swelling, mass loss and tensile properties of PVA, heparin and PVA/heparin hydrogels. The hydrogels were equilibrated in PBS pH 7.4 at 37ºC.

Mass swelling Mass loss* (%) ratio*, q

PVA 5.8 ± 0.04 24.7 ± 0.7

PVA/hep 19/1 6.5 ± 0.1 26.7 ± 0.4

PVA/hep 17.5/2.5 6.6 ± 0.1 27.1 ± 2.3

Heparin 17.3 ± 0.5 31.9 ± 1.6

* after 7-d incubation

Copolymerising PVA with ionic molecules such as chondroitin sulfate (CS) has been shown to increase the overall mass swelling ratio. In one study, the mass swelling ratio increased from 7.2 for a 10 wt% PVA hydrogel to around 8 for a PVA/CS copolymer with 4/1 (w/w) ratio [99]. Heparin molecules are highly sulfated and more negatively charged than CS, therefore copolymerising PVA with heparin was expected to cause a similar, if not higher, increase in the co-hydrogel swelling ratio, for a lower amount of heparin incorporated. This trend was observed in PVA/heparin co-hydrogels

- 97 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

of 19/1 and 17.5/2.5 (w/w) compositions, which had swelling ratios of 6.5 and 6.6,

respectively.

While the mass losses of the hydrogels may seem high, they are within the normal

range seen for (meth)acrylate modified hydrogels [30, 99, 110]. High sol fraction in

hydrogels with low crosslinker density is not unexpected. The PVA and heparin

macromers each had an average of 3 methacrylate groups per polymer chain, which was

considered a low crosslinker density considering the minimum number of crosslinkers

per polymer chain that is needed to facilitate crosslinking is 2. Since some polymer

chains might have less than the average crosslinker density, or even none, some of these

chains are bound to be released from the hydrogels during the swelling process.

Therefore it is not unexpected for hydrogels with low crosslinker density to have high

sol fraction. Varying the crosslinker density provides a way of controlling a hydrogel’s

sol fraction, swelling, and mechanical properties; this aspect has been described in more

detail in Chapter 3 of this thesis.

5.3.1.2 Equilibrium mass loss and co-hydrogel composition

Pure heparin hydrogels had a higher mass loss than pure PVA hydrogels, and the Heparin copolymers had mass loss values in between those of the individual hydrogels (Table

5.1). Therefore it was important to assess the effectiveness of PVA-heparin

copolymerisation, by evaluating the actual amount of heparin retained in the hydrogels

at equilibrium. To do this, the PVA/hep 17.5/2.5 hydrogels’ mass loss was analysed for

its heparin and PVA content. A DMMB assay was used to quantify the amount of

heparin in the co-hydrogel mass loss and it was found that by 24 h, as previously shown

by Figure 5.2, ~50% of the heparin originally incorporated had been released from the

co-hydrogel network, with no additional heparin loss up to 7-d incubation.

- 98 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

The 7-d mass loss contents of the PVA, heparin, and PVA/hep 17.5/2.5 hydrogels are illustrated in Figure 5.4. The theoretical mass loss of the co-hydrogels was estimated on the basis that the PVA and heparin components were released at the same proportion as from the pure PVA and pure heparin hydrogels, respectively. When comparing the theoretical mass loss profile of the PVA/hep 17.5/2.5 hydrogels to their actual mass loss, it was observed that the PVA loss from the co-hydrogel was approximately the same as the theoretical prediction, while heparin loss from the co-hydrogel was noticeably higher than the theoretical amount.

40 35 heparin PVA 30 25

20 15 10

Hydrogel (%)lossmass 5

0 PVA PVA/hep PVA/hep heparin 17.5/2.5 17.5/2.5 theoretical* experiment^

Figure 5.4 Comparison of hydrogel mass loss and their compositions. *estimated using the mass loss of pure PVA and heparin hydrogels, ^determined by DMMB assay.

The higher loss of heparin than PVA from the co-hydrogel suggested that the crosslinking of heparin with PVA was not as effective as the crosslinking among PVA macromers, possibly due to the steric hindrance caused by the complex structure of the heparin and this effect was observed as phase separation of the two polymers when the heparin/PVA ratio exceeds 1/8 (2.5% heparin in a 20% macromer mixture). The water- solubility of PVA and heparin is also different at all pH levels. It is likely that the charges on heparin are arranged unevenly along the chain, producing patches of positive

- 99 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels and negative charges. When non-ionic polymers such as PVA are present, they are not capable of absorbing the charges from heparin and thus enhancing the phase separation.

This kind of phase separation has been observed in mixtures of proteins and polysaccharides, when two non-interacting random coil polymers are present together at a sufficiently high concentration, typically above 10 wt% [195].

Nonetheless, heparin was not lost continually from the PVA/hep 17.5/2.5 gels and

~50% of the heparin initially loaded was still retained at equilibrium. This result appeared to be consistent in the other co-hydrogels of different compositions. While covalent incorporation of heparin was confirmed, it is noted that the incorporation efficiency was still relatively low, and will need to be improved in order to make this a commercially feasible device. Various techniques could be done to increase the incorporation efficiency, for example by increasing the functional group density, which has been shown to increase the polymer content of PVA hydrogels in Chapter 3 and in previous works [108, 112].

By knowing the amount of heparin (and thus PVA) that remained in the co- hydrogels and the final macromer concentration, the actual composition of the co- hydrogels at equilibrium could be calculated. The composition of the PVA/hep 17.5/2.5 hydrogel after sol fraction extraction was determined to be 13.4/1.2 PVA/heparin, with a final macromer concentration of 14.6 wt%. The final compositions of the hydrogels are summarised in Table 5.2 below.

- 100 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

Table 5.2 Nominal and equilibrium compositions of PVA, heparin and PVA/heparin hydrogels. The hydrogels were initially formulated with 20 wt% macromer. Hydrogel Equilibrium Equilibrium composition (%) macromer content (%) PVA Heparina

PVA 15.1 15.1 -

PVA/hep 19/1 14.7 14.2 0.5

PVA/hep 17.5/2.5 14.6 13.4 1.2

Heparin 13.6 - 13.6

a determined by DMMB

5.3.1.3 Mechanical properties

Despite showing structural stability as indicated by the equilibrium mass loss, pure heparin hydrogels were found to be too weak to be mechanically tested and copolymerisation with PVA gave heparin structural support. While the addition of heparin proved to be useful for altering the water uptake properties of the resulting network, it was also important to minimise the amount of heparin added so that the beneficial mechanical properties of the PVA hydrogels would not be adversely affected.

Copolymerising 2-hydroxyethyl methacrylate (HEMA) with PEG diacrylate was previously shown to increase the swelling property of pure poly(HEMA) hydrogels.

[168]. The tensile modulus of the poly(HEMA) was increased with the addition of PEG, from 4.5 MPa up to 7.9 MPa in hydrogels with 30% PEG, despite studies reporting PEG gels to have much lower tensile moduli compared to poly(HEMA) [169]. The resulting poly(HEMA)/PEG co-hydrogels, however, had lower elongation and strength. Another study showed that the addition of CS, a glycosaminoglycan like heparin, into PVA hydrogels resulted in an increase in both swelling and compressive moduli in the co- hydrogels – properties that are usually a trade-off in neutral hydrogel systems [99]. The charged molecules of CS were thought to alter the properties of PVA hydrogels so that

- 101 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels the increase in mechanical properties was achieved without sacrificing the hydrogel water content.

In this study, PVA/heparin hydrogels were formed at the same total macromer concentration (20 wt%) as PVA to reduce variability arising from the percent macromers, which has been shown to influence strongly the tensile modulus [112]. The average tensile properties of pure PVA hydrogels were UTS of 122 ± 22 kPa, 168 ± 26

% strain, and an elastic modulus of 61 ± 14 kPa. A decrease in tensile strength and strain with increasing heparin composition was observed. Representative stress vs. strain curves of the hydrogels are depicted in Figure 5.5.

0.15

0.12

0.09

0.06 stress (MPa) stress

0.03

0 0 50 100 150 200 strain (%)

Figure 5.5 Representative stress vs. strain curves for PVA and PVA/heparin hydrogels. PVA (―), PVA/hep 19/1 (---), and PVA/hep 17.5/2.5 (···) hydrogels.

A decrease in tensile strength and strain was observed with increasing heparin concentration. However, it was difficult to determine definite stress and strain values of hydrogels at failure, due to micro-defects in their structure such as air bubbles. For this reason, the tensile modulus was a more reliable feature that is less prone to errors in the testing and is therefore a more accurate way to compare the hydrogels [112]. The overall tensile moduli of all co-hydrogels were found to closely match that of pure PVA

- 102 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels hydrogels (Table 5.3), which suggests that addition of up to 2.5 wt% heparin did not affect the mechanical stability of PVA gels.

Table 5.3 Tensile properties of PVA and PVA/heparin hydrogels. The hydrogel sol fraction was extracted prior to testing and the hydrogels tested in their hydrated state. Tensile modulus (kPa)

PVA 61 ± 14

PVA/hep 19/1 67 ± 13

PVA/hep 17.5/2.5 68 ± 8

5.3.2 Bioactivity of PVA/Heparin Co-Hydrogels

Results in the previous sections of this Chapter demonstrated that the inclusion of up to 2.5 wt% heparin in the PVA hydrogels was stable and did not disturb the network integrity of the hydrogels. In Chapter 4, heparin has been shown to retain its anticoagulation and growth factor signalling properties following methacrylate modification and UV exposure. Once the heparin molecules were incorporated into the hydrogel network, it was important to test for the ability of the co-hydrogels to initiate or enhance the binding and signalling of proteins to their complementary receptors.

To bind a protein, the specific binding sequences on the heparin chains, which are now an integral part of the co-hydrogels, must be accessible to the protein for attachment. The amount of heparin that needed to be incorporated in the hydrogels depended on the availability of the binding sequences of the covalently crosslinked heparin, and on the relative compatibility of the PVA component. Therefore the hydrogels needed to be tested for their ability to present bioactive heparin. For this purpose, PVA/heparin co-hydrogels (20 wt%) of different compositions were made and their sol fractions extracted before the commencement of any bioactivity tests.

- 103 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

5.3.3.1 Anticoagulation property

When a foreign material is exposed to blood, plasma proteins are adsorbed onto the surface of the material. This leads to the activation of clotting factors or followed by the adhesion and activation of platelets, resulting in the formation of a fibrin network [189].

Studies measuring the anticoagulant activity of heparin-modified polymer surfaces have been done, with the purpose of improving the blood compatibility of the polymers used in blood contacting devices such as catheters [92, 196]. While earlier studies on the development of non-thrombogenic biomaterials rely on the release of heparin molecules, Merrill et al. described the formation of such materials by the covalent crosslinking of heparin with PVA by glutaraldehyde, producing hydrogels with evenly distributed heparin molecules throughout its structure [156]. The potency of these glutaradehyde-crosslinked PVA/heparin hydrogels in facilitating thrombin inactivation was demonstrated, and was confirmed in more detail in similar studies [116-117].

Grafting heparin onto polyurethanes [188-189] and collagen tubes [196] has also been shown to cause suppressed thrombus formation and increased plasma recalcification time. Thrombin adsorption on PVA hydrogels was shown to increase when heparin was present in the PVA as a crosslinked component [119]. It was indicative that the antithrombogenicity of the hydrogel would arise from the interaction of adsorbed thrombin with PVA-bound heparin and the antithrombin that is available in plasma.

In Chapter 4, heparin’s anticoagulant property was compared prior to and after methacrylate modification, with the result showing no alteration in the heparin activity following methacrylate group attachment. Similar testing was thus done on the co- hydrogels using the same plasma clotting assay. In this work, heparin did not act as a surface coating of the co-hydrogels but rather was distributed throughout the hydrogels.

The activation of proteins such as ATIII by the covalently incorporated heparin would

- 104 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels depend on the accessibility of the specific protein binding sites by the protein (Figure

5.6).

Figure 5.6 Illustration of the accessibility of ATIII binding sites in the PVA/heparin co- hydrogels. Figure courtesy of S. Yogisaputra.

Normal human plasma was incubated with either PVA or PVA/heparin hydrogels, and its clotting time was recorded and compared to a series of heparin controls. In order to facilitate anticoagulation, hydrogel bound heparin chain must be directly accessible for ATIII to bind to and be activated, via high-affinity binding to the pentasaccharide sequence on heparin. The inhibition of coagulation factors IIa (thrombin) or Xa will occur when these factors gain access to the ATIII-heparin unit and bind covalently to

ATIII [178]. The presentation of anticoagulation activity by the hydrogels would then have to rely on the accessibility of the ATIII binding sequence on the heparin chains, by the ATIII and thrombin.

The clotting time of plasma with no heparin added was 110 ± 3 s. Plasma incubated with pure PVA hydrogels clotted at similar time (102 ± 4 s), suggesting that

PVA hydrogels did not have a significant influence in ATIII activation or inhibition, for that matter. The delay in fibrin formation started to be seen when plasma was incubated

- 105 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels with PVA/heparin 19.5/0.05 co-hydrogels. A gradual increase in clotting time was observed with increasing amount of heparin (Figure 5.7), reaching up to 166 ± 13 s when plasma was incubated with PVA/hep 19.8/0.2 co-hydrogels.

500 92 180 400 160 300 43 140 200 120 21 0 5 100

Plasma clotting100 time (s) Plasma clotting time (s) 0 80 -1 0 1 2 3 4 5 6 PVA PVA/hep PVA/hep PVA/hep PVA/hep Heparin concentration in 19.99/0.01 19.95/0.05 19.9/0.1 19.8/0.2 p las m a (µµµg /m L )

Figure 5.7 Anticoagulant activity of PVA and PVA/heparin hydrogels. The anticoagulant activity was determined by measuring plasma clotting time as a response to the hydrogels (left). Each hydrogel was sol fraction-extracted prior to incubation with 200 µL plasma. The number above each bar indicates the equilibrium heparin content (µg) for the co-hydrogel composition that the bar represents. As a control, the anticoagulant activity of heparin in solution was measured (right).

As the sol fractions of all hydrogels were removed prior to the clotting assay, any anticoagulant activity detected from the co-hydrogels could only result from hydrogel- bound heparin. The actual heparin content of the co-hydrogels at equilibrium, as determined by DMMB assay, was indicated by the numbers above the bar graph in

Figure 5.7. The equilibrium heparin content of PVA/heparin co-hydrogels was determined to be approximately 50% of the original heparin composition prior to sol fraction extraction, and the values are summarised in Table 5.4 below.

- 106 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

Table 5.4 Average equilibrium heparin content in the PVA/heparin co-hydrogels. The amount of heparin released during sol fraction extraction was determined by DMMB and thus the amount remaining in the co-hydrogels could be calculated.

Hydrogel composition (%) Equilibrium heparin Equilibrium hydrogel a PVA heparin composition (%) heparin content (µµµg)

20 0 0 0 19.99 0.01 0.006 5 19:95 0.05 0.02 21 19.9 0.1 0.05 43 19.8 0.2 0.10 92 a as determined by DMMB

The PVA/hep 19.8/0.2 hydrogels were determined to have an equilibrium heparin composition of ~0.1% following sol fraction extraction (Table 5.4). This composition correlates to ~92 µg heparin per co-hydrogel, and when the PVA/hep 19.8/0.2 co- hydrogel was incubated in 0.2 mL plasma, the concentration of hydrogel-bound heparin in the plasma could be calculated to be approximately 460 µg/mL. Looking at the controls, which tested various heparin concentrations on plasma (Figure 5.7, right), a heparin concentration of 5 µg/mL in plasma was sufficient to extend the clotting time of normal plasm by more than 5 min. If all of the hydrogel-bound heparin in the co- hydrogels was accessible for ATIII binding and activation, which amounts to about 100 times more than required to delay plasma clotting in the controls, the resulting clotting time from incubation with PVA/hep 19.8/0.2 would be much higher than with the 5

µg/mL heparin control. However the opposite happened. Incubation with PVA/hep

19.8/0.2 resulted in clotting time of just under 3 minutes, while heparin controls with lower total heparin concentration could delay the clot formation for longer. From these results, it was evident that not all heparin in the co-hydrogels was accessible to the

ATIII, possibly due to the mesh size of the hydrogels (~1.35 kDa, calculated from the

- 107 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

Peppas-Merrill equation [5, 30]) that was too small for ATIII molecules (~58 kDa)

[197] to diffuse through.

When plasma was incubated with co-hydrogels that had heparin content higher than that of PVA/hep 19.8/0.2, fibrin clot did not form even at 3 h, after which the timing was discontinued. Thrombin has been shown to adsorb strongly to PVA and

PVA/heparin hydrogels, and the presence of heparin has been shown to facilitate the thrombin-ATIII complex formation [122]. It was likely that the critical amount of heparin that needed to be available for thrombin and ATIII binding on the surface of the gels had been reached by having 0.2% heparin in the co-hydrogels.

5.3.3.2 Growth factor activation

The ultimate goal for choosing heparin/HS as the biological component in the scaffold was to have the heparin molecules attract growth factors and signal these factors to surrounding cells. Heparin/HS molecules are present in the ECM and on cell surfaces, providing a stable reservoir of FGF-2 that can be released in response to appropriate stimuli for cell signalling [198]. Heparin form ternary complexes with FGF and FGF receptors. Once the heparin molecules were incorporated into the hydrogel network, it was important to test for the ability of the co-hydrogels to signal FGF-2 to live cells bearing the FGF receptor. The availability of the FGF-2 binding sites on the co-hydrogels is illustrated in Figure 5.8.

- 108 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

Figure 5.8 Illustration of the accessibility heparin on PVA/heparin co-hydrogels for the signalling of FGF-2 to its cell receptor. Figure courtesy of S. Yogisaputra.

The FGF-2 dependent BaF3 cells were seeded onto co-hydrogels containing 0.5, 1 and 2% heparin. The BaF3 cells are B lymphocytes, non-adherent cells that require direct contact with or close proximity to other cells in order to facilitate interaction via ligands and their cell surface receptors [199]. Cell proliferation assays using BaF3 cells would therefore rely on the amount of cell contact with the hydrogels, and whether or not they are able to access ligands that are crosslinked into the hydrogels.

The BaF3 cells in this study were thus seeded directly on top of the hydrogels and allowed to settle. Similar to the plasma clotting assay, as heparin is distributed throughout the PVA/heparin co-hydrogels, unless the FGF-2 could diffuse into the hydrogels through the gap between hydrogel crosslinks (1.35 kDa), FGF-2 binding to heparin could only occur with heparin present on the hydrogel surface . Direct seeding of the cells on the hydrogels would maximise chance of contact with the randomly distributed heparin. After 72 h incubation, the BaF3 cells seeded on PVA/heparin co- hydrogels showed increased proliferation compared to those on PVA only hydrogels

(Figure 5.9).

- 109 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

1.00 hydrogels 0.80 controls

0.60

0.40 Proliferation index 0.20

0.00 PVA PVA/hep PVA/hep PVA/hep 0 nM 3 nM 30 nM 19.5/0.5 19/1 18/2 heparin heparin heparin (-ve) (+ve) (+ve)

Figure 5.9 BaF3 cell proliferation on PVA and PVA/heparin hydrogels. Cell proliferation on the PVA hydrogels and the co-hydrogels was measured after 72 h incubation. Cells incubated with no heparin (negative) and with 3 and 30 nM heparin in solution (positive) served as controls. All test and control samples were incubated with 0.3 nM FGF-2. Experimental results were n=3.

The increase in proliferation when cells were incubated with PVA/heparin co- hydrogels, compared to with PVA-only hydrogels, demonstrated that the incorporated heparin was able to bind FGF-2, and that it could signal the cells to grow. The positive controls, BaF3 cells incubated with heparin in solution, showed increased cell proliferation with increasing heparin concentration. The cells incubated with the co- hydrogels, however, produced similar cell responses despite the different heparin concentrations in the co-hydrogels. These results suggested that the maximum amount of hydrogel-bound heparin that was accessible to the cells was already reached with

0.5% heparin in the hydrogels. FGF-2-dependent cell proliferation was still observed at lower heparin compositions in the co-hydrogels. Figure 5.10 depicts further the

PVA/heparin co-hydrogels activity in the cellular signalling of FGF-2, with the purpose of showing the required amount of heparin in the hydrogels to elicit cell response.

- 110 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels

0.48 hydrogels controls 0.32

0.16

Proliferation index Proliferation 0.00

-0.16 PVA PVA/hep PVA/hep PVA/hep 0 nM 0.3 nM 0.6 nM 19.95/0.05 19.5/0.5 19/1 heparin heparin heparin (-ve) (+ve) (+ve)

Figure 5.10 The increase in BaF3 cell proliferation with increasing heparin content in the PVA/heparin co-hydrogels. Cell proliferation on the co-hydrogels was compared to cell responses to negative (no heparin) and positive (3 and 30 nM heparin) control conditions. All test and control samples were incubated with 0.3 nM FGF-2. Experimental results were n=3.

The results showed an increasing trend in cell proliferation with heparin content, even though the difference was not statistically significant in compositions between

0.05% heparin and 0.5% heparin. Even at 0.05%, the presence of heparin in the co- hydrogels still greatly improved the proliferation of the BaF3 cells compared to when they were seeded on pure PVA hydrogels.

Having a minimum, growth factor-regulating, amount of heparin in the co- hydrogels might be advantageous in applications such as tissue engineered constructs.

This way, the anticoagulant effect of the co-hydrogels could be minimised, reducing the risk of excessive bleeding [56]. The results also mean that the bioactive PVA/heparin co-hydrogels could be formulated with a minimum amount of heparin that would not adversely alter the physical properties of PVA hydrogels, the supporting construct. The plasma clotting assay and cellular FGF-2 signalling study supported the hypothesis that the amount of heparin for protein binding has reached saturation at initial heparin compositions of lower than 1 wt% in the PVA/heparin co-hydrogels. These tests essentially measured co-hydrogel activity based on the surface availability of heparin,

- 111 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels since the hydrogel mesh size was too small for proteins to diffuse through. With this low, but effective, amount of heparin, the co-hydrogels inherited the antithrombogenicity and growth factor signalling capability of heparin, while only experiencing slight alteration in the structure and strength of the PVA hydrogels (Tables

5.1 and 5.3).

5.4 CONCLUSION

To fulfil the role as scaffolds for tissue repair, the advantageous resemblance of hydrogels to the ECM in vivo needs to be coupled with their ability to present specific signalling molecules to surrounding cells. The localised presence of molecules that would encourage cellular infiltration and vascularisation into the scaffold is of utmost importance to ensure the viability of the engineered tissue. As a model of HS, the glycosaminoglycan that is responsible for regulating the formation of vascular networks, functionalised heparin molecules were successfully copolymerised with PVA to form co-hydrogels. The main aims of the Chapters were to address the effect of heparin incorporation into PVA and to investigate the bioactivity of the resulting co- hydrogels. Therefore the stability of heparin in the co-hydrogels and of the resulting hydrogel structure was examined prior to investigating aspects of the co-hydrogels’ bioactivity. Covalent coupling with PVA, as opposed to simple mixing, enabled heparin to be presented in a sustained manner. The PVA/heparin co-hydrogels were found to be capable of presenting and activating FGF-2 to promote cell proliferation even at low concentrations, suggesting that the co-hydrogels could be formulated with minimum amount of heparin as to least compromise the strength of the surrounding synthetic scaffold. This study demonstrated that the composite PVA/heparin hydrogels created act

- 112 -

Chapter 5 – Structural and Functional Characterisation of PVA/Heparin Co-Hydrogels as scaffolds with controllable network properties, capable of presenting signalling molecules to cells.

ACKNOWLEDGEMENTS

The author would like to acknowledge Stella Yogisaputra for help with the illustrations in Figures 5.6 and 5.8.

- 113 -

Chapter 6

Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

he extracellular matrix (ECM) in vivo is continually remodelled by the action of Tvarious enzymes that specifically target different components of the matrix. The mammalian enzyme responsible for the degradation of heparan sulphate (HS) chains from the ECM and cell surfaces is known as heparanase. In humans, platelets have been known to contain high levels of heparanase and were used as the enzyme source for this study. The aim of this Chapter was to investigate the activity of heparinised hydrogels following incubation with platelet extract (PE), in order to simulate the responses that might occur when the hydrogels, as tissue engineered scaffolds, come in contact with blood products at the sites of injury. The heparanase activity of the PE on heparin that was used as a model of HS was confirmed by the decrease in molecular weight. PE treatment of heparin diminished its anticoagulation property but increased its FGF-2 signalling activity, suggesting that the PE’s heparanase activity cleaves at the 3-O- sulfated glucosamine to produce large fragments that can signal cell receptors. When poly(vinyl alcohol) (PVA)/heparin co-hydrogels were incubated with PE, the release of heparin fragments was confirmed by the alteration of heparin activity in the supernatant of PE-degraded co-hydrogels compared to that of the non-degraded co-hydrogels.

- 114 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

6.1 BACKGROUND AND AIMS

Tissue engineering scaffolds are designed to mimic the extracellular matrix

(ECM) by their structure and interactions with the surrounding host environment within.

Heparan sulfate (HS) molecules, which are a major part of the ECM, are continually synthesised and replaced in the body, a process that helps maintain homeostasis and regulate angiogenesis. HS molecules are synthesised as a HS proteoglycan (HSPG), which consists of a protein cores that carry HS chains as branched structures [56]. The

HSPGs sequester numerous enzymes, growth factors, and cytokines on the cell surface and in the extracellular matrix (ECM), often as an inactive reservoir [200]. Cleavage of the HSPGs or HS chains releases fragments of these proteins or polysaccharides from the ECM, and can effect their bioactivity [154]. In humans, the protein core of HSPG is susceptible to degradation by several proteases, while the HS chains are cleaved by a single heparanase enzyme, which is an endo-β-glucuronidase and is often referred to as heparanase-1.

Heparanase cleaves HS at specific sites to release HS fragments from the ECM

[201-202]. Proteins such as growth factors that are trapped within the ECM or on the cell surface are released with these HS fragments, making them available to their receptors on the cell surface, causing a change in the environmental conditions that may modulate local tissue responses. One of the primary functions of heparanase is to degrade basement membrane HSPGs at sites of injury or inflammation [192, 203-204].

This action will disrupt the basement membrane, allowing the migration of inflammatory cells and releasing HS-bound growth factors that induce the migration and enhance the survival of endothelial cells [152]. Heparanase was found to specifically release active basic fibroblast growth factor (FGF-2) by cleaving HS from

- 115 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity the ECM and cell surfaces and has been correlated traditionally with cell invasion associated with angiogenesis and inflammation [198].

Platelets have been shown to be a good source of heparanase, and research has demonstrated that heparanase activity was dependent on pH, with its optimum pH being reported in the range of 5.1 to 6.8 [153, 203, 205-206]. Other sources of heparanase include endothelial cells (EC) and smooth muscle cells (SMC), although the HS- degrading activity from these cells has been reported to be less than 10% that of platelet extract [153]. Nevertheless cell lysates from both ECs and SMCs have been shown to contain factors that can further activate platelet heparanase.

Enzymatic degradation studies of biologically derived hydrogels have been performed previously to examine the substrate availability on the crosslinked biopolymers for enzymatic cleavage of the gels. Collagen, for example, is degraded by metalloproteases, especially collagenase, and serine proteases, allowing degradation to be locally controlled by cells present in the engineered tissue [16]. Most of the studies have often used bacterial enzymes instead of those from mammalian sources, such as in the degradation studies of hydrogels formulated from chondroitin sulfate (CS) [99] and fibrin [207]. Mammalian heparanase differs from other HS-degrading enzymes such as heparinase and heparitinase from Flavobacterium heparinum, which are eliminases type enzymes that typically cleaves HS into di- or tri-saccharide units [208]. Therefore, to simulate the degradation of heparin in vivo and examine the activity of the resulting heparin fragments, human platelet extract (PE) was used in this study as a source of heparanase.

The aim of this Chapter was to expose the heparin-functionalised poly(vinyl alcohol) (PVA) hydrogels to enzymatic degradation, in order to simulate a condition that the hydrogels were likely to encounter in vivo. By investigating the outcome of this

- 116 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity degradation on the activity of heparin presented by the hydrogels, it would be possible to predict some of the effects that the hydrogels will have on the local environment surrounding the implant site. The heparin-degrading activity of PE was examined by analysing the molecular weight (MW) of heparin following incubation with PE, as well as the anticoagulation and FGF-2 signalling properties of the heparin fragments. The activity of heparin may be influenced by its chain length, therefore low molecular weight (LMW) heparin was compared to heparin in terms of its activity. The degradability of the covalently-crosslinked heparin from the PVA/heparin hydrogels was then examined. Following incubation with PE, extracts from heparin and

PVA/heparin hydrogels were collected and analysed to assess their molecular weight and tested for their corresponding bioactivities.

6.2 EXPERIMENTAL

6.2.1 Materials

Human platelet-rich serum (Australian Red Cross Blood Service) was used as received. Toyopearl® HW-50S media (Tosoh Bioscience) for size exclusion chromatography was washed prior to packing into a Pharmacia HR10/30 column. Three heparin samples, sodium salt from porcine intestinal mucosa, were used: heparin (grade

I-A, average MW 17-19 kDa, Sigma), low molecular weight (LMW) heparin (average

MW 4-6 kDa, Sigma), and Deligoparin™ (average MW 2-3 kDa, Opocrin, Italy).

Coomassie Plus Reagent, bovine serum albumin (BSA), casein, antibodies (7B5, a-ser,

HPA-1), biotinylated anti-mouse antibodies (Amersham), Streptavidin-conjugated horseradish peroxide (Amersham), Tween-20 (0.01% in PBS), ABTS reagent was prepared as instructed by the manufacturer.

- 117 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

PVA and PVA/heparin hydrogels, formed as described in Chapter 5, were used in this Chapter to assess turnover by heparanases and platelet extract. Briefly, 20 wt%

PVA and PVA/heparin hydrogels were formed by photo-polymerisation of methacrylated PVA and heparin macromers. The starting materials were obtained from

Sigma: PVA (average MW 13-23 kDa, 98% hydrolysed), heparin sodium salt (grade I-

A, from porcine intestinal mucosa, average MW 17-19 kDa) and glycidyl methacrylate

(97% purity). Dyes 1,9-dimethylmethylene blue (DMMB) (Sigma), AlamarBlue

® (Invitrogen), and CellTiter 96 AQueous One Solution Reagent ([3-(4,5-dimethylthiazol-

2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium, inner salt (MTS))

(Promega) were also used as received. The cell culture media RPMI-1640 was obtained from Sigma, prepared following the manufacturer’s instructions, and sterile filtered (0.2

µm). Recombinant human fibroblast growth factor-2 (FGF-2) (Invitrogen) was used without further purification.

6.2.2 Extraction of Heparanase from Platelets

The extraction of heparanase from platelets was done using a freeze-thaw method adapted from Graham et al. [153]. The platelet-rich serum (total volume 329 mL, 7.3 ×

108 platelets/mL) was centrifuged at 1940 g for 20 mins at 15ºC. After discarding the supernatant, the platelet pellets were resuspended gently in DPBS at room temperature and re-centrifuged. The platelets were resuspended in minimum amount (17 mL) of

DPBS, and subjected to 5 cycles of freeze-thaw using a -70ºC freezer and thawing in an ethanol bath. The platelet extract (PE) was clarified by centrifugation at 15,600 g for 15 min at 4ºC, and filter-sterilised using a 0.22 µm filter.

The protein content of the PE was determined by a Coomassie Plus assay, using bovine serum albumin (BSA) (0–1 mg/mL in DPBS) as protein standards. Briefly,

- 118 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity samples of the PE and BSA standards (10 µL) were aliquoted in quadruplicates into a

96-well plate. A volume of 300 µL Coomassie Plus Reagent was added to each well.

The plate was placed on a shaker for 30 s to mix the reagents. After 10 min incubation at room temperature, the absorbance was read at 595 nm. The absorbance values of the samples were correlated to the BSA standards to quantify the protein content in the PE.

In order to confirm the presence of heparanase in the PE, an enzyme-linked immunosorbent assay (ELISA) was used as described previously [209]. In ELISA, typically a specific antibody that binds the compound of interest (the antigen) is used to identify the presence and amount of the antigen in a sample. The antibody is linked to an enzyme, and in the final step a reagent would be added so that the enzyme will give a signal when binding is detected. PE aliquots were transferred to a 96-well plate to coat.

After washing and incubation with blocking solution for 1 h, separate sample wells

(duplicates) were probed with a monoclonal anti-heparanase antibody (HPA-1) or with either one of the negative controls: a monoclonal antibody against perlecan (7B5) or against the polyclonal serglycin (a-ser). As briefly mentioned in Chapter 2, perlecan and serglycin are forms of proteoglycans, consisting of proteins cores, to which HS chains are attached when they were newly synthesised. Since PE was prepared in DPBS, DPBS solution was used as a control in the assay. The absorbance of the samples was read at

405 nm on a Tecan Infinite F200 Microplate Reader.

The presence of heparanase was confirmed by an increase in absorbance detected in the ELISA when the samples were probed with antibodies against heparanase antigen in the PE, compared to results for the proteoglycan antigens (perlecan and serglycin) and the absorbance achieved in the absence of coated antigen (DPBS).

- 119 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

6.2.3 Heparanase Activity of PE on Heparin

The specific activity of the isolated heparanase was determined by examining the amount of heparin digested as a function of time and pH. Heparin was prepared in phosphate buffered saline (PBS) at pH 5.1 and 7.4, at a concentration of 2 mg/mL, to which 50 µL of PE (40 µg protein) was added per mg of heparin. The solutions were incubated at 37ºC. At 15 h, 1 d and 3 d, samples were collected and heated up at 80ºC for 15 mins to denature the enzyme. As a control and to ensure the preservation of heparin activity after heating, a sample at t = 0 with the same heparin and PE concentrations was heated up at 80ºC.

PE activity on heparin. The PE-treated heparin samples were run through a size exclusion chromatography (SEC) column packed with Toyopearl® HW-50S media.

Fractions (0.25 mL) were collected from the column and analysed by a DMMB assay as described in Chapter 4. Diluted samples (50 µL) of each fraction, along with a series of heparin standards (0-60 µg/mL), were each mixed with 200 µL DMMB reagent in a 96- well plate. The absorbance of the solutions was read at 535 nm, and the amount of heparin in the fractions was obtained by correlating the absorbance of the samples to that of the heparin standards. The profiles of the PE-treated heparin were then compared to those of non-treated heparin.

Heparanase cleaves heparin at the glycosyl linkages and affects the ability of the

DMMB dye to bind to heparin [210]. Previous assays used to determine heparanase activity have used radio- or biotin-labelled HS as a substrate for the enzyme [180, 211], where the heparanase activity was estimated by measuring the loss in detection of the label, attached to the cleaved HS. By adapting this concept to the DMMB assay, the amount of heparin degradation could be estimated by:

- 120 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

 A   PE−heparin  percent degradatio 1n −=  × %100 (6.1)  Aheparin 

where Ax was the area under the curve for sample x, while PE-heparin refers to

PE-treated heparin.

Molecular weight of heparin fragments. An aqueous gel permeation chromatography (GPC) system was used to estimate the molecular weight of the PE- treated heparin samples. The GPC system consisted of a pump (LC-10ATVP,

Shimadzu), an auto injector (SIL-10ADVP, Shimadzu), two PL aquagel-OH mixed 8

µm columns (Polymer Laboratories) and a refractive index detector (RID-10A,

Shimadzu). The GPC used polystyrene sulfonate as internal standards, therefore the results needed to be calibrated. To construct a heparin standard curve, three heparin samples of known molecular weights, namely heparin, LMW heparin and deligoparin, were run through the GPC. The retention times (RT) of the heparin standards were plotted against their weight-average molecular weights (Mw) given by the supplier. The

RTs of the PE-treated heparin samples were applied against the heparin standard curve to estimate their Mw.

6.2.4 Effect of Molecular Weight on Heparin Activity

The anticoagulation activity of heparin, prior to and after PE treatment, was compared using a plasma clotting assay (aPTT) as described in Chapter 4. Briefly, the 2 mg/mL heparin (+/-) PE samples were heated at 80ºC for 15 min to stop PE activity.

The samples were then diluted to 30 µg/mL in saline. A volume of 20 µL from each sample was added to 180 µL plasma and warmed at 37ºC for ~60 seconds. 200 µL pre- warmed phospholin ES aPTT reagent was added, mixed and further incubated at 37ºC for 3-5 minutes. Coagulation was initiated by adding 200 µL pre-warmed CaCl2, at

- 121 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity which point the timing began. Clotting time was recorded when the first sign of fibrin formation was observed.

The FGF-2 signalling activity of heparin and LMW heparin was compared in a

BaF3 FR1c cell proliferation assay. This assay was also used to compare the activity of heparin prior to and after degradation with PE. The maintenance of the BaF3 cells and the mitogenic assay were performed as described in Chapter 4. Prior to starting the assay, BaF3 cells were washed with and transferred to RPMI 1640 medium containing

10% FBS, and incubated at 37°C for approximately 15 h. Cells were then washed and resuspended at 1×105 cells/mL with 0.3 nM FGF-2 and the appropriate heparin or LMW heparin concentrations.

The cell suspension (100 µL) was seeded into 96-well plates and incubated at

37°C for 72 h. Afterward 10 µL MTS dye was added into each well, incubated for 6 h, before reading the plates for absorbance at 495 nm (Tecan Infinite F200 Microplate

Reader). The data were normalised following the method by Padera, et al. [100], in which cell proliferation index was calculated by dividing the increase in cell number for each test condition by the increase in cell number for the positive control. The positive control was BaF3 cells incubated with 0.3 nM FGF-2 and 30 nM heparin.

6.2.5 Enzymatic Degradation Profile of Heparin from Hydrogels

Heparin and PVA/heparin hydrogels (20 wt%) were prepared as described in

Chapter 5. In brief, heparin or PVA/heparin hydrogels (10 mm diameter, 1 mm thick) were formed by photopolymerisation at 30 mW/cm2 UV for 6 min. The sol fractions of the hydrogels were extracted for 24 h prior to the addition of the PE. The amount of heparin in the sol fraction was analysed by a DMMB assay as described in Chapter 5.

From this result, the amount of heparin left in the hydrogels after sol fraction extraction

- 122 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity could be determined. PE was added at 50 µL per mg heparin left in the hydrogels, and the hydrogels were incubated with PE at pH 5.1 and 7.4, for 1 d and 3 d. Control hydrogels were also incubated in PBS without the addition of PE. At the end of incubation time, supernatants of the PE-treated hydrogels were collected, heated to stop

PE activity, and tested in a plasma clotting assay and a cell proliferation assay described above.

6.3 RESULTS AND DISCUSSION

6.3.1 Heparanase Activity of the Platelet Extract

Platelets have been shown to be the most reliable source from which heparanase could be obtained [153, 204, 212]. To examine the contents of the platelet extract (PE) used in this study, tests were done to determine the amount of protein and the presence of heparanase in the PE. The overall protein concentration of the PE was determined in the Coomasie Plus assay to be 810 ± 28 µg/mL. An ELISA was done on the PE, using an anti-heparanase antibody (HPA-1) as a probe for heparanase. Antibodies against perlecan (7B5) and serglycin (a-ser) served as negative controls. The PE was tested with the antibodies and compared to DPBS, which was the solution used during platelet extraction. In the ELISA, a higher signal (absorbance) from the PE relative to the DPBS thus indicates antibody binding to the antigen. The presence of heparanase in the PE was confirmed by the increased signal when PE was probed with HPA-1 (Figure 6.1).

- 123 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

0.3 HPA-1 0.25 7B5 a-ser

405 0.2

0.15

0.1 Absorbance,A

0.05

0 PE DPBS

Figure 6.1 Content of platelet extract (PE) as determined by ELISA. Antibodies against heparanase (HPA-1), perlecan (7B5) and serglycin (a-ser) were used to probe for heparanase and HSPG content of the PE. An increase in absorbance in PE compared to that in DPBS confirms the presence of the probed antigens.

The heparanase activity of the PE was then tested on heparin, at physiological pH

(7.4) and at pH 5.1, a condition at which heparanase has been found to be most active towards HS [212]. This correlates to the heparanase activity in vivo; the sites of inflammation and tumours in vivo have been recorded to have relatively acidic pH, enabling heparanase-mediated cleavage of HS from the ECM [206]. The heparanase activities of the PE on heparin at both pH 5.1 and 7.4 were thus characterised by SEC, to analyse the amount of heparin that was degraded following incubation with PE, and by GPC , to determine the molecular weight of the heparin fragments.

6.3.1.1 Amount of heparin degradation following PE treatment

DMMB dye has been used for the quantification of sulfated glycosaminoglycans

(GAGs) content in tissues [187] and the reactivity of the dye was found to decrease following enzymatic-degradation of GAGs, presumably into smaller oligosaccharides

[210]. This property has in the past been used to analyse the content of a mixture of sulphated GAGs. Specific GAG-degrading enzymes could be applied to eliminate the

- 124 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

DMMB-binding capacity of other GAGs in the mixture, leaving behind the GAG of interest for quantification by DMMB.

It was expected that heparin would experience a similar loss in DMMB reactivity if cleaved by the heparanase contained in the PE. This would present a complication in that it may hinder the detection of the total amount of heparin (by DMMB) in the fractions eluted from the SEC column. However, since only heparin that has been degraded will lose its reactivity with DMMB, it was proposed in this study that this loss in detection, as previously used for radio-labelled HS [180], could also be used to estimate the amount of heparin that was digested by the heparanase (Equation 6.1).

The degree of heparin degradation over time was analysed by observing the elution profiles of PE-treated heparin after chromatography through a SEC column.

Fractions of 0.25 mL were collected from the column and analysed for their heparin content by the DMMB assay. In Figure 6.2, the molecular weight profiles of PE-treated heparin at various time points were compared to the profiles of non-degraded heparin.

When treated with PE at pH 5.1, the amount of heparin detected by DMMB continued to decrease when heparin was incubated with PE for longer time periods, as shown by the MW profiles of heparin in Figure 6.2(A). The amount of heparin that was cleaved by the PE was estimated at ~90% after 1d.

- 125 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

A 0.2 535

0.1 Absorbance,A

0.0 0 0.1 0.2 0.3 0.4 0.5 0.6

Partition coefficient, Kav

B 0.2 535

0.1 Absorbance,A

0.0 0 0.1 0.2 0.3 0.4 0.5 0.6

Partition coefficient, Kav

Figure 6.2 Heparin degradation profiles at pH 5.1 (A) and 7.4 (B). The amount of degradation is represented by the loss of binding to DMMB dye, resulting in a decrease in absorbance at 535 nm. Profiles of the PE-degraded samples at 6h (●), 15h (), and 1d (▲) were compared to that of non-treated heparin (–) at both pH values.

Heparanase was previously shown to only be partially active at pH 7.2, where it acted as a cell adhesion molecule instead of an enzyme [206]. Here, heparin samples treated with PE at pH 7.4 did show significant degradation as detected by a decrease in

DMMB reactivity (Figure 6.2(B)). However, the samples treated at pH 7.4 were less degraded than those at pH 5.1 and only reached ~68% degradation by 1 d. At this pH, both the 15h and the 1d treatments resulted in heparin with similar profiles, indicating no increase in degradation was after 15h of incubation.

- 126 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

6.3.1.2 Molecular weight of PE-treated heparin

Heparanases are classified as endo-type glycosidases, which are enzymes that cleave polysaccharides at the non-reducing end of glycosidic linkages [208]. The heparin and HS degrading activities of platelet heparanase have specifically been associated with an endo-β-D-glucuronidase. Unlike heparin/HS degrading enzymes from bacterial sources (endo-type lyases), which act on the α14 linkage regions on the sugar chains to yield di- and oligo-saccharides, mammalian heparanases cleave

HS/heparin into fragments that are still long enough to contain specific binding sequences and be active [153]. Thus, the maintenance of growth factor signalling activity has suggested that the heparanase-cleaved fragments might be longer than a decasaccharide (~4.5 kDa) in length [154].

In the previous section, PE was shown to degrade heparin at both pH 5.1 and 7.4.

To confirm that the degradation was a result of heparanase activity, the PE-treated heparin samples were analysed by GPC to estimate their molecular weight post degradation. Using heparins of known molecular weights as standards, the molecular weights of the heparin fragments were estimated. Heparin samples after different incubation times with PE at pH 5.1 and 7.4 were eluted through a GPC column to obtain their molecular weight profiles. Figure 6.3 confirms the degradation of the heparin chains, at pH 5.1 and 7.4, respectively.

- 127 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

1.2

1.0

0.8

0.6

0.4 Normalised response Normalised 0.2

0.0 18 19 20 21 22 Retention time (min)

Figure 6.3 Heparin degradation by platelet extract at pH 5.1 and 7.4. Molecular weight profiles of heparin after 1 d incubation at pH 5.1 (+), and at pH 7.4 (○), were compared to heparin standards: heparin (17-19 kDa, –), low molecular weight heparin (4-6 kDa, --) and deligoparin (2-3 kDa, -·-)

The shift in retention time for heparin following PE-treatment at pH 5.1 was greater than when it was treated at pH 7.4. This result supported the previous observation with the SEC column that more heparin was being degraded at pH 5.1, yielding heparin with lower average molecular weight. Using heparin samples of known molecular weight as standards, the weight average molecular weight (Mw) of the PE- treated heparin was determined. The change in the Mw of heparin with prolonged incubation time are summarised in Table 6.1. The decrease in Mw was most noticeable during the first 15-24 h of incubation, after which the drop in Mw became less evident.

Heparin, with an original Mw of 18 kDa, was reduced to ~9.5 and ~14 kDa after a 1d- incubation with PE at pH 5.1 and 7.4, respectively.

- 128 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

Table 6.1 Molecular weight estimate of platelet extract (PE)-treated heparin. Heparin was incubated with PE at 37ºC, sampled at various time points and analysed by GPC.

Incubation Weight average molecular weight (Mw) (kDa) time pH 5.1 pH 7.4

0 18 18 15 h 10.3 16.7 1 d 9.5 14.1 3 d 8.8 11.7

From this molecular weight analysis of PE-treated heparin, it can be concluded that heparanase in the PE cleaved the heparin at one or two sites along the chain, resulting in molecules about half its original length. The results also confirmed the glucuronidase activity typical of platelet heparanase. The Mw of heparin post PE- treatment reflects fragments that were longer than 19 saccharides (~8.6 kDa), which is important for the preservation of protein signalling capacity of heparin which requires no less than 10 saccharides.

6.3.2 Effect of Molecular Weight on Heparin Activity

Apart from being structure-specific, the bioactivities of heparin are also dependent on the length of the heparin chain. The activity of PE-degraded heparin fragments were thus investigated. PE-treated heparin were tested in a plasma clotting assay and on BaF3 cells. Prior to the clotting assays and following incubation with PE, heparin degradation was stopped by heating the solution (80ºC for 15 min) to denature the heparanase activity of the PE. Although heating did not cause any further chain scission, it might affect the conformation of the heparin chains and thus the activity of the molecule.

To investigate this effect of heating on the activity of heparin, non-heated and heated heparin samples were tested on plasma. If the anticoagulation activity of heparin decreased after heating, the clotting time of plasma treated with heated heparin samples

- 129 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity would be shorter. Figure 6.4, however, shows that there was no difference in plasma clotting time between heated and non-heated samples. The result confirmed the preservation of heparin activity as an anticoagulant, after heating to stop PE degradation.

210 non-heated 200 heated

190

180

170 Plasma Plasma clotting time(s) 160

150 pH 5.1 pH 7.4

Figure 6.4 The effect of heating on the anticoagulation property of heparin. The anticoagulation property (left) and growth factor activation (right) of non-heated and heated heparin were compared. Heated heparin was incubated at 80ºC for 15 min, the same temperature and time that were used to denature heparanase activity of PE.

The effect of heating on the ability of heparin in modulating cellular signalling of

FGF-2 was also examined. Compared to heparin controls that were not heated, the heating step affected heparin activity by decreasing cellular signalling of FGF-2, but not affecting the plasma clotting time (Figure 6.5). The amount of proliferation, as indicated by the proliferation index, decreased by approximately 50% in BaF3 cells that were incubated with heated heparin relative to those in the presence of non-heated heparin.

- 130 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

0.8 non-heated heated

0.6

0.4 Proliferation index 0.2

0.0 pH 5.1 pH 7.4

Figure 6.5 The effect of heating on the FGF-2 signalling activity of heparin. The anticoagulation property (left) and growth factor activation (right) of non-heated and heated heparin were compared. Heated heparin was incubated at 80ºC for 15 min, the same temperature and time that were used to denature heparanase activity of PE.

Heparin has been reported to maintain its anticoagulant activity following autoclave sterilisation at 121ºC for up to 10 min [213], but the effect of heating in growth factor signalling has not been reported. As the anticoagulation property was preserved, it meant that the pentasaccharide sequence required for antithrombin III

(ATIII) binding was unaltered, but it was possible that the side groups of the saccharides that are responsible for growth factor binding were disturbed. The binding region for FGF-2 has been identified as a pentasaccharide sequence containing a single essential O-sulfate group, at C2 of iduronic acid [190]. Selective O-desulfation at this site has been shown to reduce the ability of the heparin to induce high affinity binding of the FGF-2 to its receptor [139]. Therefore, one possibility is that the heat was affecting these side groups at the reducing end of the chain. For consistency, all heparin samples in this study were heated after incubation with PE, prior to any bioactivity assays.

- 131 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

The anticoagulant property of PE-treated heparin was examined and compared to that of non-treated heparin. As discussed in Chapter 4, heparin binds to ATIII, a naturally occurring plasma protease inhibitor, significantly accelerating the rate at which ATIII inhibits the activity of the coagulation proteases (factor Xa and thrombin).

In a 20 kDa heparin chain there may be a maximum of two ATIII binding sites.

Therefore in LMW heparin preparations below 8 kDa, it is unlikely that they would contain more than one ATIII binding sequence [214]. Figure 6.6 shows the decrease in plasma clotting time with shorter heparin chains.

200

180

160

140

120

100

Plasma Plasma clotting time(s) 80

60

40 6 8 10 12 14 16 18 20 Mw of PE-treated heparin (kDa)

Figure 6.6 Changes in the anticoagulant activity of heparin, after an incubation period with PE. The original weight average molecular weight (Mw) of heparin was 18 kDa, and decreased with prolonged PE-treatment at pH 5.1 (○) and 7.4 (●). All heparin samples had the same initial concentration of 3 µg/mL plasma. Plasma without heparin clotted at 55 ± 2 s.

The decrease in plasma clotting time with the decrease in heparin molecular weight indicates a loss of anticoagulant activity of the PE-degraded heparin, as a result of reduced number or elimination of the ATIII binding sites. It has been observed that heparanase can cleave the HS/heparin chain so that the binding sites for ATIII are

- 132 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity destroyed [179, 215]. Scheme 6.1 shows a HS chain containing the pentasaccharide sequence recognised by ATIII. When this pentasaccharide was part of a decasaccharide molecule or longer, heparanase cleaved the HS at the β-glycosidic linkage (marked by arrow in Scheme 6.1) next to the central glucosamine residue that contains the 3-O- sulfonate group essential for the interaction with AT-III [136, 215].

O CH OSO - 2CH OSO3- C OH 2CH OSO3- 2 3 O O O O O O O OH OH OH O O O O O O OH O OH O OSO - O OH OH 3 OH OH - OSO NH SO - OH OH HN OH NH SO3- 3 3 O

Scheme 6.1 The β-glycosidic linkage on heparan sulfate that is cleaved by heparanase. The ATIII binding region is indicated by the bracket .

The clotting assay on PE-treated heparin showed the decrease in anticoagulant activity, caused by the disruption of the ATIII specific pentasaccharide sequence. The minimum HS/heparin binding sequence required to facilitate the cellular signalling of

FGF-2, on the other hand, has been shown to be a decasaccharide [139, 184, 216].

While this required binding site for FGF-2 is longer than that for ATIII, it has been reported that heparanase-cleaved heparin fragments were more active in signalling growth factors [154]. To test whether low molecular weight (LMW) heparin has an increased capacity compared to heparin in presenting FGF-2 to its cell surface receptor,

BaF3 cells were incubated with both LMW heparin (4-6 kDa) and heparin (17-19 kDa) at various concentrations.

Cells incubated with LMW heparin showed a higher proliferation than those incubated with heparin across the concentration range (Figure 6.7). Heparin had 38-42 saccharide units, while the LMW was made up of approximately 9-13 saccharides

- 133 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

(approximately 25% of the length of heparin). Both molecular weight preparations had the required minimum of 10 saccharide units for signalling of FGF-2 [56, 185].

1

0.8

0.6

0.4 Proliferation index Proliferation

0.2

0 0.0001 0.001 0.01 0.1 1 Heparin concentration (µµµg/mL)

Figure 6.7 The effect of heparin molecular weight on the cellular signalling of FGF-2. BaF3 cells were incubated with heparin (●) or LMW heparin (○) in the presence of 0.3 nM FGF- 2 for 3d. The proliferation index is the normalised cell response with reference to cells incubated with 30 nM heparin and 3 nM FGF-2.

The anticoagulation activity of the heparin preparations, as given by the manufacturer, was 180 USP (United States Pharmacopeia) unit/mg for heparin [217] and 134 IU (International Unit)/mg (~143 USP/mg) for LMW heparin [218]. This means the LMW heparin is more active on a w/w basis, which was also confirmed by results in Figure 6.7 above. This indicates that heparin, which had the higher molecular weight, contains inhibitory or non-signalling regions [139]. The release of HS from the

ECM by heparanase activates HS-bound growth factors that are previously dormant

[209], therefore heparanase (PE)-degraded heparin, either in solution or from a biomaterial, is hypothesised to be more active.

Heparin samples were prepared at the same concentration before PE was added.

The PE treatment was stopped at different time points, and the samples were diluted and

- 134 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity introduced to BaF3 cells. From the molecular weight analysis in the previous section of this chapter, heparin that was incubated with PE at pH 5.1 had a lower average molecular weight at each time point compared to those incubated at pH 7.4. The cell responses were then plotted against the Mw of the heparin samples (Figure 6.8).

0.5

0.4

0.3 Proliferation index 0.2

0.1 6 8 10 12 14 16 18 20 Mw of PE-treated heparin (kDa)

Figure 6.8 Changes in the FGF-2 cellular signalling activity of heparin, after an incubation period with PE. The original weight average molecular weight (Mw) of heparin was 18 kDa, and decreased with prolonged PE-treatment at pH 5.1 (○) and 7.4 (●). All samples had the same initial heparin concentration.

A trend of increased cell proliferation was observed with shorter heparin chains.

Even when produced in different pH conditions, heparin fragments of similar Mw induced comparable cell proliferation. This finding supports the previous results examining the increased FGF-2 signalling of LMW heparin (see previous Figure 6.7), that lower molecular weight heparin is more active due to elimination of inhibitory sites for growth factor signalling, causing a change in chain conformation and enhanced growth factor signalling [219].

The results from the BaF3 cell proliferation and plasma clotting assays suggest that the heparanase cleaves heparin chain at regions that still allow growth factor

- 135 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity signalling yet inhibit the anticoagulant activity. These findings agree with existing literature, and as the role of HS in the body is to regulate growth factor signalling and cell migration during tissue regeneration, the inhibition of anticoagulant activities at the repair site might also be significant [203]. Platelets have been reported to exhibit both anticoagulant and procoagulant role, depending on its mode of activation [220]. The release of heparanase activity from platelets is thought to play a role in modifying

HS/heparin molecules in the ECM [221], which might encourage thrombus formation at sites of injury. Intensive research has been done to better understand the HS-degrading action of heparanase on a molecular level, providing a platform for heparanase study on biomaterials. However no reports have explored the effect of platelet heparanase on the activity of heparin-based hydrogels. In light of this, the present work investigated the degradation capacity of heparanase (PE) on hydrogel-bound heparin, and its resulting impact on the hydrogel’s activity.

6.3.3 Enzymatic Degradation Profile of Heparin from the Hydrogels

It is likely that only a small portion of the basement membrane proteoglycan HS chains are accessible by the heparanase in vivo. This is because a variety of matrix proteins, cell adhesion proteins, and growth factors will be bound at various sites along the glycosaminoglycan chain [222]. By recognizing a specific sequence or modification, heparanase does not need to rely on other structural features of the glycosaminoglycan that may be obscured when the proteoglycan is incorporated in the matrix. Similar to the

HS in the ECM, access to heparin in the PVA/heparin co-hydrogels might also be restricted due to the crosslinking of the heparin with PVA chains.

In this study, heparin and PVA/heparin hydrogels were incubated with PE to investigate the degradability of heparin from the hydrogels. Sol fraction-extracted

- 136 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity hydrogels were incubated with PE at pH 5.1 and 7.4. The sol fractions of the hydrogels were extracted in order to remove any non-incorporated heparin from the hydrogels.

Once the non-incorporated heparin was removed, additional release of heparin could only be facilitated by chain degradation. PVA/hep 19/1 co-hydrogels were incubated with PE and sampled after incubation at 1 d and 3 d. As mentioned previously, the ability of DMMB dye to bind heparin and to be used as a detection method is impaired following enzymatic degradation. Therefore the amount of heparin digested and released could be indirectly measured by testing the supernatant from the PE-treated hydrogels.

Supernatants from the 1-d and 3-d PE-treated co-hydrogels were tested for their anticoagulant activities. Compared to the supernatant from non-treated co-hydrogels

(PVA/hep only), an increase in plasma clotting time was observed when plasma was incubated with the supernatants from the 1 d PE treatment, at both pH 5.1 and 7.4

(Figure 6.9). This increase in clotting time indicates the presence of more heparin fragments, containing ATIII binding sites, in the supernatants. Although the heparanase has been shown to cleave at ATIII binding region [179, 215] and reduce the anticoagulation properties of heparin in solution (previous Figure 6.6), the released heparin chains might still contain ATIII binding sites and therefore provide more sites for ATIII activation. Moreover, once the ATIII binds to a coagulation factor, heparin dissociates from the ATIII-factor complex and can activate more ATIII [178].

- 137 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

100

pH 5.1 80 pH 7.4

60

40

Plasma clotting time(s) clotting Plasma 20

0 Supernatant, Supernatant, Supernatant, PVA/hep only PVA/hep + PE, 1d PVA/hep + PE, 3d

Figure 6.9 Anticoagulant activity of PVA/heparin 19/1 hydrogel supernatants. Hydrogel supernatants were collected after 1- and 3-d incubation with PE, and compared with the supernatant from non PE-treated hydrogels. Plasma without hydrogels clotted at 55 ± 2 s.

Incubation with the 3-d supernatants, however, caused reduced plasma clotting time for the pH 5.1 sample, but a slightly increased clotting time for the pH 7.4 sample.

PE degradation of heparin in solution at pH 5.1 for 3 d was previously shown to create fragments with reduced anticoagulation properties (previous Figure 6.6).

The 3-d treatment of the co-hydrogels with PE at pH 5.1, however, produced a supernatant that had lower anticoagulation activity than the supernatant from 1 d PE- treatment (Figure 6.9). On the other hand, PE treatment for 3 d at pH 7.4 still produced a supernatant that had a slight increase in clotting activity (80 ± 8 s) compared to the 1 d samples (73 ± 3 s). PE degradation of heparin in solution at pH 5.1 for 3 d was previously shown to result in heparin fragments smaller than those from PE-treatment at pH 7.4 (Table 6.1), and consequently with lower anticoagulant activity (Figure 6.7).

Therefore, although more heparin chains might be released from the co-hydrogels after

3-d incubation, it was possible that they have been degraded further at pH 5.1 compared to at pH 7.4. The results indicate that PE-treatment at both pH 5.1 and 7.4 released

- 138 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity heparin chains from the PVA/hep co-hydrogels. The loss in activity in the pH 5.1 supernatant, but not in the pH 7.4 sample, confirmed the higher heparanase activity at that pH compared to at pH 7.4.

The supernatants of the PE-treated hydrogels were then tested for their growth factor signalling property. During incubation with PE, as suggested in the plasma clotting assay, some heparin chains might be released from the PVA/hep 19/1 co- hydrogels by the heparanase activity of the PE. Heparanase-degraded HS/heparin has been reported to actively signal various growth factors including FGF-2 to cells [149,

154, 209]. The BaF3 cells used in this growth factor signalling assay were dependent on

FGF-2 for growth. Heparin has been shown to increase the effect of FGF-2 in increasing

BaF3 cell proliferation (Chapter 4). Therefore, any increase in proliferation when the cells were incubated with the supernatants can be attributed to the presence of heparin fragments, as a result of degradation from the co-hydrogels.

As all the supernatants contained PE, a control was done in which cells were incubated with PE but without heparin. Cells incubated with PE only in the presence of

FGF-2 did not show an increase in proliferation compared to cells incubated with FGF-

2 only and without heparin. This ruled out PE as a factor that increases FGF-2 signalling in cells. Figure 6.10 shows the proliferation results of BaF3 cells incubated with the supernatants of PE-treated PVA/hep 19/1 co-hydrogels, and also with those of non- treated co- hydrogels (PVA/hep only) as controls.

- 139 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity

0.50 pH 5.1 pH 7.4 0.40

0.30

0.20 Proliferation index 0.10

0.00 Supernatant, Supernatant, Supernatant, PVA/hep only PVA/hep + PE, 1d PVA/hep + PE, 3d

Figure 6.10 FGF-2 signalling activity of PVA/heparin 19/1 hydrogel supernatants. Hydrogel supernatants were collected after 1- and 3-d incubation with PE. Proliferation index indicates the increase in cell number in samples relative to that in the positive control (cells incubated in 3 nM FGF-2, 30 nM heparin)

In general, longer PE treatment of the co-hydrogels produced supernatants that induced higher cell proliferation (Figure 6.10). PE-treatment of heparin at pH 5.1 has been shown to create heparin fragments that, although shorter, were more active in

FGF-2 signalling to cells than those produced at pH 7.4 (previous Figure 6.8). A similar outcome was reflected here, in that the hydrogel supernatants from the pH 5.1 PE treatment had higher proliferation effects that those from the pH 7.4 treatment. It can be concluded that the PE-treatment of the PVA/heparin co-hydrogels produced supernatants that had higher FGF-2 signalling activity, resulting in the increased cell proliferation.

These results support the findings from the plasma clotting assay, and show that sites for enzymatic heparin cleavage are still available when heparin was covalently incorporated in the hydrogels. The dual activity observed in the hydrogel supernatants confirmed the activity of PE (heparanase) that was previously observed with heparin in solution. PE cleaved heparin into fragments about half its original length (~18 kDa, 40

- 140 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity saccharides). Although the activation of ATIII only requires a pentasaccharide sequence on heparin, heparanase cleaves heparin specifically at this sequence, resulting in the loss of ATIII binding and anticoagulation activity. The heparin fragments were still sufficiently long to present the decasaccharide sequence required for FGF-2 signalling, as proven by the increase in cell proliferation. The enhanced effect of this dual activity on the co-hydrogels at pH 5.1, a pH value typical in an inflamed site in the body, compared to that at neutral pH suggests the importance of increased FGF-2 signalling and suppression of anticoagulant activity during the process of wound healing.

6.6 CONCLUSION

The aim of this Chapter was to investigate the activity of heparin that has been functionalised and polymerised as part of a biosynthetic hybrid biomaterial, and incubated with platelet extract in an attempt to model what might happen when the material is exposed to blood or activated plasma, both of which may be present at sites of injury. Platelet extract (PE) has a high concentration of heparanase, the naturally occurring enzyme in mammals that is responsible for the turnover of HS/heparin. The

PE activity was pH dependent, and cleaved heparin at an average of one site, decreasing the average molecular weight by a factor of two. This result was consistent with the action of glucuronidase-type enzymes, confirming the heparanase activity of the PE.

The PE was also found to be capable of releasing covalently-crosslinked heparin from the hydrogels. PE-degraded heparin was found to have higher FGF-2 signalling capacity, however its anticoagulant activity was diminished. This suggests that the heparanase cleaves the chain at regions that still allow growth factor signalling yet inhibit the anticoagulant activity. This interesting difference in the bioactivities between the treatments supports the hypothesis that platelets have the capacity to limit the anti-

- 141 -

Chapter 6 – Enzymatic Degradation of Heparinised Hydrogels and Its Effect on Bioactivity coagulant activity and as such will promote a pro-thrombogenic activity that is critical to successful thrombus formation. It also suggests that this same action is capable of liberating growth factor activities that enable the signalling of FGF2 to cause cell proliferation, which is critical to the downstream process of wound healing.

This study on the enzymatic degradability of heparin from PVA/heparin co- hydrogels and its potential effect on a molecular level is an important and novel finding of the work presented in this thesis. No report has previously shown the dual activity of heparanase on heparin incorporated into a biomaterial. Understanding these changes in heparin activity when the co-hydrogels are subjected to physiological factors is essential, and this work has contributed into providing better insight on the potential effects of heparinised scaffolds when applied to a site of injury for tissue repair.

ACKNOWLEDGEMENTS

The author would like to acknowledge and thank Bill Cheng for his help in the analysis of the platelet extract content, and Litania Lie for additional data on the aPTT and BaF3 cell assays.

- 142 -

Chapter 7

Conclusions and Future Work

he incorporation of heparan sulfate (HS) as signalling molecules in a hydrogel Tscaffold was investigated in this thesis. The extracellular matrix (ECM) acts as the body’s natural scaffold during tissue regeneration, therefore the properties of the

ECM need to be considered when designing scaffolds for tissue engineering. Hydrogels resemble the ECM in structure due to their high swelling capacity and elasticity, and the use of synthetic materials offers greater control over the resulting hydrogel properties.

For use in assisting tissue regeneration, however, synthetic scaffolds lack recognition sites that are essential for encouraging cellular interactions at the site of injury.

Scaffolds need to present molecular cues that not only encourage cell adhesion, but also regulate growth factors for cell migration and assembly. HS facilitates such cues by binding and signalling proteins via specific binding sequences. HS can be incorporated into hydrogel scaffolds by simple physical blending but they may diffuse out of the scaffold over time. Therefore covalently linking HS to the hydrogel network would be highly beneficial. Heparin, the highly sulfated variant of HS, was used in this study as a model of HS.

The challenges associated with the covalent incorporation of glycosaminoglycans like HS/heparin into synthetic hydrogels include the requirement for the biopolymers to undergo chemical modification, which has the potential to disrupt the binding sites for

- 143 -

Chapter 7 – Conclusions and Future Work biological function. To facilitate copolymerisation, the modification of the biopolymers should be such that the functional groups on the biopolymers have similar reactivity as those on the synthetic polymers. Most importantly, the incorporation of biopolymers should be a means to impart bioactivity to, and without compromising the structural integrity of, the scaffold.

The present work strived to address these challenges, with an overall objective of creating a scaffold material that has structural stability and strength, capable of signalling growth factors to cells. Co-hydrogels of PVA and heparin were formed by photo-initiated radical polymerisation. PVA and heparin were functionalised with pendant methacrylate groups that would act as crosslinkers upon irradiation.

Copolymerisation requires the pendant groups on both polymers to have similar reactivity. While PVA has been functionalised with (meth)acrylate pendant groups by reactions in both organic and aqueous media [108, 162, 223], the functionalisation of heparin is limited by its insolubility in organic solvents.

Two synthesis methods for the functionalisation of PVA were considered: the well studied 2-isocyanatoethyl methacrylate (ICEMA) synthesis [163] and the glycidyl methacrylate (GMA) synthesis. Although ICEMA synthesis was more efficient in methacrylate incorporation, the GMA method produced PVA macromers and hydrogels that had comparable cell compatibility and strength with those from ICEMA. The photopolymerisation of these methacrylated PVA still result in some sol fraction, but a more efficient crosslinking system such as glutaraldehyde [30] may present complications for biomedical applications due to the presence of toxic residues [77, 79], and are thus not employed in this research. The GMA synthesis was successfully adapted to functionalise heparin and hereby chosen for the functionalisation of PVA.

- 144 -

Chapter 7 – Conclusions and Future Work

Heparin contains specific saccharide sequences, through which it binds and activate various proteins. Disruptions of these binding sites, by fragmentation or modifications such as methacrylate attachment, might result in the loss of protein activation. The structural integrity of the methacrylated heparin was confirmed by molecular weight analysis. The activities of heparin and hep-MA were compared in a plasma clotting assay, and in a proliferation assay using BaF3 cells, cells that express receptors specific for fibroblast growth factor (FGF-2). Heparin macromers induced similar anticoagulation effects and cell proliferation as the unmodified heparin, suggesting that the binding sites for antithrombin III (ATIII) and FGF-2 remained available and active following the addition of ~3 methacrylate groups per heparin chain.

The functionalised heparin was successfully copolymerised with PVA to form co- hydrogels. The covalent incorporation of heparin was confirmed, with ~50% of heparin remaining in the hydrogels at equilibrium (post 24 h). The swelling capacity of the co- hydrogels increased with heparin content, and the addition of up to 2.5 wt% heparin did not compromise the strength of the surrounding synthetic scaffold. The co-hydrogels were able of presenting and activating FGF-2 to promote cell proliferation, with as low as 0.05 wt% heparin incorporated in the hydrogels.

The effect of enzymatic degradation on the activity of heparin, and the degradability of the covalently-crosslinked heparin from the poly(vinyl alcohol)

(PVA)/heparin hydrogels were studied. Platelet extract (PE) was used as the source of heparanase, which activity was dependent on pH. The PE was found to cleave heparin

(~18 kDa), reducing them to ~8 kDa (pH 5.1) and ~14 kDa (pH 7.4) after a 1d incubation period. These PE-degraded heparin fragments were found to have higher

FGF-2 signalling capacity, however their anticoagulant activity was diminished. This suggests that the heparanase activity in PE cleaved heparin on average in one or two

- 145 -

Chapter 7 – Conclusions and Future Work points along the chain, at the ATIII binding sites, leaving fragments longer than the minimum decasaccharide required for cellular signalling of FGF-2.

SUMMARY OF CONTRIBUTIONS

The localised presentation of heparin as protein-signalling ligands in a synthetic hydrogel was demonstrated by the formation of PVA and heparin as co-hydrogels.

Biomaterials from collagen and synthetic polymers have invariably been modified with heparin as a means to control the delivery or presentation of growth factors [8, 40, 156].

These heparin-modified matrices have been shown to promote growth in various cell lines and induce vascular formation in animal models [49, 57, 75, 159]; however the specific function of the matrix-bound heparin as growth factor-activating molecules has not been extensively studied. In addition, no studies have examined the activity of heparinised matrices following degradation by their naturally-occurring enzymes. The present work attempts to address these unexplored aspects of the research area, with the contributions summarised as follows.

(1) The same synthesis method for methacrylate group attachment (GMA) could be adapted for both PVA and heparin, resulting in them carrying methacrylate groups of equal structure and reactivity;

(2) Heparin maintained structural integrity and activity following modification with

GMA and UV irradiation, meaning that heparin functionalisation by reaction with GMA is a viable route that will allow the incorporation of heparin into PVA, by photopolymerisation;

(3) The structural and mechanical studies of the PVA/heparin co-hydrogels confirmed the integrity of PVA hydrogel when up to 2.5 wt% heparin was incorporated

- 146 -

Chapter 7 – Conclusions and Future Work in the network, provided an insight to the effect of biomolecule incorporation on the mechanical properties of synthetic scaffolds;

(4) When applied with FGF-2, the PVA/heparin co-hydrogels induced higher cell proliferation than PVA hydrogels. The cells used were FGF-2-dependent for growth, therefore confirming that any increase in proliferation to be a direct result of FGF-2 presentation by heparin in the co-hydrogels.

(5) The low amount (<<2.5 wt%) of heparin required for activity in the co- hydrogels suggests that heparin-functionalised hydrogels could be formulated with minimal amounts of heparin so as not to disturb structural integrity of the supporting scaffold.

(6) The dual effect observed when the co-hydrogels were incubated with platelet extract supports the hypothesis of platelets having the capacity to limit anticoagulation and thus promoting blood clot formation, which may be critical in the wound healing process of tissue repair; and

(7) The heparanase activity in platelet extract was able to cleave the covalently- incorporated heparin, resulting in the enhanced FGF-2 activation by heparin to promote cell proliferation, which is also critical to the process of tissue regeneration.

RECOMMENDATIONS FOR FUTURE WORK

The advantages of having high growth factor signalling but low anticoagulation effects may be advantageous in tissue repair. Non-anticoagulant heparin has previously been developed to prevent bleeding complications when an injectable hydrogel system was administered [48]. Although the heparin had been oxidised to reduce its anticoagulant property, its FGF-2 activation property remained [94-95] and neovascularisation was induced near the injected site of the hydrogel in a mouse model.

- 147 -

Chapter 7 – Conclusions and Future Work

In the present study, heparin of different molecular weights was shown to present different strength of FGF-2 activation. LMW heparin had higher FGF-2 signalling on a weight-to-weight basis and less ATIII binding sites, and therefore might be used in place of heparin to modify PVA hydrogels.

To investigate the feasibility of LMW heparin to be covalently crosslinked to

PVA, a preliminary experiment was done to attempt methacrylate-functionalisation of

LMW heparin and examine its incorporation into PVA hydrogels. The functionalisation of LMW heparin by reaction with GMA was successful with 0.8 mol% methacrylate substitution, which corresponded to 1 methacrylate group per chain. The degree (mol%) of methacrylate attachment was the same as in heparin, suggesting that the efficiency of methacrylate functionalisation was independent of the molecular weight of the heparin chains. The methacrylated LMW heparin was copolymerised with PVA, and was found to be covalently incorporated. By optimising the degree of functionalisation on LMW heparin, co-hydrogels could be formed derived from the high growth factor-signalling

LMW heparin.

The present work has successfully demonstrated growth-promoting capacity of heparinised hydrogels. Further investigation on cell survival inside a 3D form of the hydrogels would confirm the feasibility of using the PVA/heparin co-hydrogels to support cell growth. Even though the macromers used to form the current co-hydrogels only had low functionalisation (average of 3 crosslinkers per chain), the mesh size of the hydrogels limited protein diffusion, let alone cellular infiltration. This is a challenge that has been associated with biosynthetic hydrogels, in that unless they are degradable, cell migration and vessel formation within the hydrogels are unlikely. Degradable PVA- poly(ethylene glycol) (PEG) hydrogels, for example, have been shown to support growth of encapsulated cells while allowing degraded regions to be replaced by new

- 148 -

Chapter 7 – Conclusions and Future Work tissue [111]. Scaffold porosity is another important factor that can encourage cellular infiltration. The formation of pores in hydrogels has been done using the more conventional lyophilisation technique [224-225], and more recently with photolithography [226]. The creation of porosity within the PVA/heparin co-hydrogels may expose additional activity of heparin in the construct, while the ubiquitous presence of heparin lining the pores of the hydrogels may serve as a chemotactic signal for cell migration.

In summary, HS-functionalised PVA hydrogels show promise for use as scaffold material in tissue repair. Favourable structure and mechanical properties, as well as the valuable sustained growth factor presentation were achieved by the covalent crosslinking of heparin with PVA. The degradability of hydrogel-bound heparin by platelet extracts, showed responses that simulate the liberation of HS activity from the

ECM by the platelets at sites of injury.

- 149 -

Bibliography

[1] Langer R, Vacanti JP. Tissue engineering. Science. 1993;260:920-6.

[2] Peppas NA, Langer R. New challenges in biomaterials. Science. 1994;263:1715-20.

[3] Hubbell JA. Biomaterials in tissue engineering. Nat Biotechnol. 1995;13:565-76.

[4] Griffith LG, Naughton G. Tissue engineering-current challenges and expanding opportunities. Science. 2002;295:1009-16.

[5] Peppas NA, Hilt JZ, Khademhosseini A, Langer R. Hydrogels in biology and medicine: From molecular principles to bionanotechnology. Adv Mater. 2006;18:1345- 60.

[6] Lee KY, Mooney DJ. Hydrogels for tissue engineering. Chem Rev. 2001;101:1869- 80.

[7] Lutolf MP, Hubbell JA. Synthetic biomaterials as instructive extracellular microenvironments for morphogenesis in tissue engineering. Nat Biotech. 2005;23:47- 55.

[8] Cai S, Liu Y, Zheng Shu X, Prestwich GD. Injectable glycosaminoglycan hydrogels for controlled release of human basic fibroblast growth factor. Biomaterials. 2005;26:6054-67.

[9] Benoit DSW, Anseth KS. Heparin functionalized PEG gels that modulate protein adsorption for hMSC adhesion and differentiation. Acta Biomater. 2005;1:461-70.

[10] Tae G, Scatena M, Stayton PS, Hoffman AS. PEG-cross-linked heparin is an affinity hydrogel for sustained release of vascular endothelial growth factor. J Biomater Sci Polym Ed. 2006;17:187-97.

[11] Cushing MC, Liao J-T, Jaeggli MP, Anseth KS. Material-based regulation of the myofibroblast phenotype. Biomaterials. 2007;28:3378-87.

[12] Benoit DSW, Collins SD, Anseth KS. Multifunctional hydrogels that promote osteogenic human mesenchymal stem cell differentiation through stimulation and sequestering of bone morphogenic protein 2. Advanced Functional Materials. 2007;17:2085-93.

[13] Mooney DJ, Mikos AG. Growing new organs. Sci Am. 1999;280:60.

[14] Patel ZS, Mikos AG. Angiogenesis with biomaterial-based drug- and cell-delivery systems. J Biomater Sci Polym Ed. 2004;15:701-26.

[15] Moroni L, Elisseeff JH. Biomaterials engineered for integration. Materials Today. 2008;11:44-51.

- 150 -

Bibliography

[16] Drury JL, Mooney DJ. Hydrogels for tissue engineering: scaffold design variables and applications. Biomaterials. 2003;24:4337-51.

[17] Sasisekharan R, Ernst S, Venkataraman G. On the regulation of fibroblast growth factor activity by heparin-like glycosaminoglycans. Angiogenesis. 1997;1:45-54.

[18] Zisch AH, Lutolf MP, Hubbell JA. Biopolymeric delivery matrices for angiogenic growth factors. Cardiovasc Pathol. 2003;12:295-310.

[19] Yannas IV, Burke JF. Design of an artificial skin. I. Basic design principles. J Biomed Mater Res. 1980;14:65-81.

[20] Pachence JM. Collagen-based devices for soft tissue repair. J Biomed Mater Res. 1996;33:35-40.

[21] Dedhar S, Ruoslahti E, Pierschbacher MD. A cell surface receptor complex for collagen type I recognizes the Arg-Gly-Asp sequence. J Cell Biol. 1987;104:585-93.

[22] Olde Damink LHH, Dijkstra PJ, van Luyn MJA, van Wachem PB, Nieuwenhuis P, Feijen J. Cross-linking of dermal sheep collagen using a water-soluble carbodiimide. Biomaterials. 1996;17:765-73.

[23] Sheridan MH, Shea LD, Peters MC, Mooney DJ. Bioabsorbable polymer scaffolds for tissue engineering capable of sustained growth factor delivery. J Controlled Release. 2000;64:91-102.

[24] Peters MC, Polverini PJ, Mooney DJ. Engineering vascular networks in porous polymer matrices. J Biomed Mater Res. 2002;60:668-78.

[25] Cohen S, Yoshioka T, Lucarelli M, Hwang LH, Langer R. Controlled delivery systems for proteins based on poly(lactic/glycolic acid) microspheres. Pharm Res. 1991;8:713-20.

[26] Cleland JL, Duenas ET, Park A, Daugherty A, Kahn J, Kowalski J, et al. Development of poly-(,-lactide-coglycolide) microsphere formulations containing recombinant human vascular endothelial growth factor to promote local angiogenesis. J Controlled Release. 2001;72:13-24.

[27] Kim T-K, Burgess DJ. Pharmacokinetic characterization of 14C-vascular endothelial growth factor controlled release microspheres using a rat model. J Pharm Pharmacol. 2002;54:897-905.

[28] Murphy WL, Peters MC, Kohn DH, Mooney DJ. Sustained release of vascular endothelial growth factor from mineralized poly(lactide-co-glycolide) scaffolds for tissue engineering. Biomaterials. 2000;21:2521-7.

[29] Griffith LG. Polymeric biomaterials. Acta Mater. 2000;48:263-77.

[30] Martens P, Anseth KS. Characterization of hydrogels formed from acrylate modified poly(vinyl alcohol) macromers. Polymer. 2000;41:7715-22.

- 151 -

Bibliography

[31] Hassan C, Peppas N. Structure and applications of poly(vinyl alcohol) hydrogels produced by conventional crosslinking or by freezing/thawing methods. Advances in Polymer Science2000. p. 37-65.

[32] Langer R, Peppas NA. Advances in biomaterials, drug delivery, and bionanotechnology. AIChE J. 2003;49:2990-3006.

[33] Nomi M, Atala A, De Coppi P, Soker S. Principals of neovascularization for tissue engineering. Mol Aspects Med. 2002;23:463-83.

[34] Kannan RY, Salacinski HJ, Sales K, Butler P, Seifalian AM. The roles of tissue engineering and vascularisation in the development of micro-vascular networks: a review. Biomaterials. 2005;26:1857-75.

[35] Finkenzeller G, Torio-Padron N, Momeni A, Mehlhorn AT, Stark GB. In vitro angiogenesis properties of endothelial progenitor cells: a promising tool for vascularization of ex vivo engineered tissues. Tissue Eng. 2007;13:1413-20.

[36] Hughes SE. Functional characterization of the spontaneously transformed human umbilical vein endothelial cell line ECV304: Use in an in vitro model of angiogenesis. Exp Cell Res. 1996;225:171-85.

[37] Peters MC, Isenberg BC, Rowley JA, Mooney DJ. Release from alginate enhances the biological activity of vascular endothelial growth factor. J Biomater Sci Polym Ed. 1998;9:1267-78.

[38] Gu F, Amsden B, Neufeld R. Sustained delivery of vascular endothelial growth factor with alginate beads. J Controlled Release. 2004;96:463-72.

[39] Nakamura S, Kanatani Y, Kishimoto S, Nakamura S-i, Ohno C, Horio T, et al. Controlled release of FGF-2 using fragmin/protamine microparticles and effect on neovascularization. J Biomed Mater Res. 2008;9999:NA.

[40] Wissink MJB, Beernink R, Pieper JS, Poot AA, Engbers GHM, Beugeling T, et al. Binding and release of basic fibroblast growth factor from heparinized collagen matrices. Biomaterials. 2001;22:2291-9.

[41] Steffens GCM, Yao C, Prevel P, Markowicz M, Schenck P, Noah EM, et al. Modulation of angiogenic potential of collagen matrices by covalent incorporation of heparin and loading with vascular endothelial growth factor. Tissue Eng. 2004;10:1502- 9.

[42] Ishihara M, Obara K, Ishizuka T, Fujita M, Sato M, Masuoka K, et al. Controlled release of fibroblast growth factors and heparin from photocrosslinked chitosan hydrogels and subsequent effect on in vivo vascularization. J Biomed Mater Res. 2003;64A:551-9.

[43] Tan CS, Jejurikar A, Rai B, Bostrom T, Lawrie G, Grøndahl L. Encapsulation of a glycosaminoglycan in hydroxyapatite/alginate capsules. J Biomed Mater Res. 2008;9999:NA.

- 152 -

Bibliography

[44] Korff T, Kimmina S, Martiny-Baron G, Augustin HG. Blood vessel maturation in a 3-dimensional spheroidal coculture model: direct contact with smooth muscle cells regulates endothelial cell quiescence and abrogates VEGF responsiveness. FASEB J. 2001;15:447-57.

[45] Melero-Martin JM, Khan ZA, Picard A, Wu X, Paruchuri S, Bischoff J. In vivo vasculogenic potential of human blood-derived endothelial progenitor cells. Blood. 2007;109:4761-8.

[46] Richardson TP, Peters MC, Ennett AB, Mooney DJ. Polymeric system for dual growth factor delivery. Nat Biotechnol. 2001;19:1029-34.

[47] Yao C, Roderfeld M, Rath T, Roeb E, Bernhagen J, Steffens G. The impact of proteinase-induced matrix degradation on the release of VEGF from heparinized collagen matrices. Biomaterials. 2006;27:1608-16.

[48] Fujita M, Ishihara M, Simizu M, Obara K, Ishizuka T, Saito Y, et al. Vascularization in vivo caused by the controlled release of fibroblast growth factor-2 from an injectable chitosan/non-anticoagulant heparin hydrogel. Biomaterials. 2004;25:699-706.

[49] Pieper JS, Hafmans T, Wachem PBv, Luyn MJAv, Brouwer LA, Veerkamp JH, et al. Loading of collagen-heparan sulfate matrices with bFGF promotes angiogenesis and tissue generation in rats. J Biomed Mater Res. 2002;62:185-94.

[50] Brey EM, Uriel S, Greisler HP, McIntire LV. Therapeutic neovascularization: Contributions from bioengineering. Tissue Eng. 2005;11:567-84.

[51] Carmeliet P, Conway EM. Growing better blood vessels. Nat Biotech. 2001;19:1019-20.

[52] Bourke SL, Al-Khalili M, Briggs T, Michniak BB, Kohn J, Poole-Warren LA. A photo-crosslinked poly(vinyl alcohol) hydrogel growth factor release vehicle for wound healing applications. AAPS PharmSci. 2003;5:E33.

[53] Edelman ER, Mathiowitz E, Langer R, Klagsbrun M. Controlled and modulated release of basic fibroblast growth factor. Biomaterials. 1991;12:619-26.

[54] Smeds KA, Grinstaff MW. Photocrosslinkable polysaccharides for in situ hydrogel formation. J Biomed Mater Res. 2001;54:115-21.

[55] Kirker KR, Luo Y, Nielson JH, Shelby J, Prestwich GD. Glycosaminoglycan hydrogel films as bio-interactive dressings for wound healing. Biomaterials. 2002;23:3661-71.

[56] Whitelock JM, Iozzo RV. Heparan sulfate: A complex polymer charged with biological activity. Chem Rev. 2005;105:2745-64.

[57] Wissink MJB, Beernink R, Scharenborg NM, Poot AA, Engbers GHM, Beugeling T, et al. Endothelial cell seeding of (heparinized) collagen matrices: effects of bFGF

- 153 -

Bibliography pre-loading on proliferation (after low density seeding) and pro-coagulant factors. J Control Release. 2000;67:141-55.

[58] Nuttelman CR, Henry SM, Anseth KS. Synthesis and characterization of photocrosslinkable, degradable poly(vinyl alcohol)-based tissue engineering scaffolds. Biomaterials. 2002;23:3617-26.

[59] Massia SP, Hubbell JA. Covalently attached GRGD on polymer surfaces promotes biospecific adhesion of mammalian cells. Ann N Y Acad Sci. 1990;589:261-70.

[60] Schmedlen RH, Masters KS, West JL. Photocrosslinkable polyvinyl alcohol hydrogels that can be modified with cell adhesion peptides for use in tissue engineering. Biomaterials. 2002;23:4325-32.

[61] Rizzi SC, Ehrbar M, Halstenberg S, Raeber GP, Schmoekel HG, Hagenmuller H, et al. Recombinant protein-co-PEG networks as cell-adhesive and proteolytically degradable hydrogel matrixes. Part II: Biofunctional characteristics. Biomacromolecules. 2006;7:3019-29.

[62] Mann BK, Tsai AT, Scott-Burden T, West JL. Modification of surfaces with cell adhesion peptides alters extracellular matrix deposition. Biomaterials. 1999;20:2281-6.

[63] Pieper JS, Hafmans T, Veerkamp JH, van Kuppevelt TH. Development of tailor- made collagen-glycosaminoglycan matrixes: EDC/NHS crosslinking, and ultrastructural aspects. Biomaterials. 2000;21:581-93.

[64] Whalen GF, Shing Y, Folkman J. The fate of intravenously administered bFGF and the effect of heparin. Growth Factors. 1989;1:157-64.

[65] van de Witte P, Esselbrugge H, Dijkstra PJ, van den Berg JWA, Feijen J. A morphological study of membranes obtained from the systems polylactide-dioxane- methanol, polylactide-dioxane-water and polylactide-N-methyl pyrrolidone-water. Journal of Polymer Science Part B: Polymer Physics. 1996;34:2569-78.

[66] Yang S, Leong K-F, Du Z, Chua C-K. The design of scaffolds for use in tissue engineering. Part I. Traditional factors. Tissue Eng. 2001;7:679.

[67] Mooney DJ, Baldwin DF, Suh NP, Vacanti JP, Langer R. Novel approach to fabricate porous sponges of poly(-lactic-co-glycolic acid) without the use of organic solvents. Biomaterials. 1996;17:1417-22.

[68] Harris LD, Kim B-S, Mooney DJ. Open pore biodegradable matrices formed with gas foaming. J Biomed Mater Res. 1998;42:396-402.

[69] Shea LD, Smiley E, Bonadio J, Mooney DJ. DNA delivery from polymer matrices for tissue engineering. Nat Biotech. 1999;17:551-4.

[70] Burke JFMD, Yannas IVPD, Quinby WCJMD, Bondoc CCMD, Jung WKMD. Successful use of a physiologically acceptable artificial skin in the treatment of extensive burn injury. Ann Surg. 1981;194:413-28.

- 154 -

Bibliography

[71] Clark RAF, Ghosh K, Tonnesen MG. Tissue engineering for cutaneous wounds. J Invest Dermatol. 2007;127:1018-29.

[72] Yannas IV, Burke JF, Gordon PL, Huang C, Rubenstein RH. Design of an artificial skin. II. Control of chemical composition. J Biomed Mater Res. 1980;14:107-32.

[73] Vercruysse KP, Marecak DM, Marecek JF, Prestwich GD. Synthesis and in vitro degradation of new polyvalent hydrazide cross-linked hydrogels of hyaluronic acid. Bioconjug Chem. 1997;8:686-94.

[74] Li Q, Wang DA, Elisseeff JH. Heterogeneous-phase reaction of glycidyl methacrylate and chondroitin sulfate: Mechanism of ring-opening-transesterification competition. Macromolecules. 2003;36:2556-62.

[75] Benoit DSW, Durney AR, Anseth KS. The effect of heparin-functionalized PEG hydrogels on three-dimensional human mesenchymal stem cell osteogenic differentiation. Biomaterials. 2007;28:66-77.

[76] Jackson RL, Busch SJ, Cardin AD. Glycosaminoglycans: molecular properties, protein interactions, and role in physiological processes. Physiol Rev. 1991;71:481-539.

[77] Huang-Lee LLH, Cheung DT, Nimni ME. Biochemical changes and cytotoxicity associated with the degradation of polymeric glutaraldehyde derived crosslinks. J Biomed Mater Res. 1990;24:1185-201.

[78] van Luyn MJA, van Wachem PB, Olde Damink L, Dijkstra PJ, Feijen J, Nieuwenhuis P. Relations between in vitro cytotoxicity and crosslinked dermal sheep collagens. J Biomed Mater Res. 1992;26:1091-110.

[79] van Luyn MJA, van Wachem PB, Damink LHHO, Dijkstra PJ, Feijen J, Nieuwenhuis P. Secondary cytotoxicity of cross-linked dermal sheep collagens during repeated exposure to human fibroblasts. Biomaterials. 1992;13:1017-24.

[80] van Wachem PB, van Luyn MJA, Olde Damink LHH, Dijkstra PJ, Feijen J, Nieuwenhuis P. Biocompatibility and tissue regenerating capacity of crosslinked dermal sheep collagen. J Biomed Mater Res. 1994;28:353-63.

[81] Pieper JS, Oosterhof A, Dijkstra PJ, Veerkamp JH, van Kuppevelt TH. Preparation and characterization of porous crosslinked collagenous matrices containing bioavailable chondroitin sulphate. Biomaterials. 1999;20:847-58.

[82] Shu XZ, Liu Y, Luo Y, Roberts MC, Prestwich GD. Disulfide cross-linked hyaluronan hydrogels. Biomacromolecules. 2002;3:1304-11.

[83] Kirker KR, Luo Y, Morris SE, Shelby J, Prestwich GD. Glycosaminoglycan hydrogels as supplemental wound dressings for donor sites. J Burn Care Rehabil. 2004;25:276-86.

[84] Macieira-Coelho A, Berumen L, Avrameas S. Properties of protein polymers as substratum for cell growth in vitro. J Cell Physiol. 1974;83:379-88.

- 155 -

Bibliography

[85] Shu XZ, Liu Y, Palumbo F, Prestwich GD. Disulfide-crosslinked hyaluronan- gelatin hydrogel films: a covalent mimic of the extracellular matrix for in vitro cell growth. Biomaterials. 2003;24:3825-34.

[86] Li H, Liu Y, Shu XZ, Gray SD, Prestwich GD. Synthesis and biological evaluation of a cross-linked hyaluronan-mitomycin C hydrogel. Biomacromolecules. 2004;5:895- 902.

[87] Shu XZ, Liu Y, Palumbo FS, Luo Y, Prestwich GD. In situ crosslinkable hyaluronan hydrogels for tissue engineering. Biomaterials. 2004;25:1339-48.

[88] Thorslund S, Sanchez J, Larsson R, Nikolajeff F, Bergquist J. Functionality and stability of heparin immobilized onto poly(dimethylsiloxane). Colloids and Surfaces B: Biointerfaces. 2005;45:76-81.

[89] Christensen K, Larsson R, Emanuelsson H, Elgue G, Larsson A. Heparin coating of the stent graft - effects on platelets, coagulation and complement activation. Biomaterials. 2001;22:349-55.

[90] Kristensen EME, Rensmo H, Larsson R, Siegbahn H. Characterization of heparin surfaces using photoelectron spectroscopy and quartz crystal microbalance. Biomaterials. 2003;24:4153-9.

[91] Johnell M, Elgue G, Larsson R, Larsson A, Thelin S, Siegbahn A. Coagulation, fibrinolysis, and cell activation in patients and shed mediastinal blood during coronary artery bypass grafting with a new heparin-coated surface. J Thorac Cardiovasc Surg. 2002;124:321-32.

[92] Johnell M, Larsson R, Siegbahn A. The influence of different heparin surface concentrations and antithrombin-binding capacity on inflammation and coagulation. Biomaterials. 2005;26:1731-9.

[93] Ono K, Saito Y, Yura H, Ishikawa K, Kurita A, Akaike T, et al. Photocrosslinkable chitosan as a biological adhesive. J Biomed Mater Res. 2000;49:289-95.

[94] Ishihara M, Ono K, Ishikawa K, Hattori H, Saito Y, Yura H, et al. Enhanced ability of heparin-carrying polystyrene (HCPS) to bind to heparin-binding growth factors and to inhibit growth factor-induced endothelial cell growth. J Biochem. 2000;127:797-803.

[95] Ishihara M, Saito Y, Yura H, Ono K, Ishikawa K, Hattori H, et al. Heparin-carrying polystyrene to mediate cellular attachment and growth via interaction with growth factors. J Biomed Mater Res. 2000;50:144-52.

[96] van Dijk-Wolthuis WNE, Franssen O, Talsma H, van Steenbergen MJ, Kettenes- van den Bosch JJ, Hennink WE. Synthesis, characterization, and polymerization of glycidyl methacrylate derivatized dextran. Macromolecules. 1995;28:6317-22.

[97] Wang L-F, Shen S-S, Lu S-C. Synthesis and characterization of chondroitin sulfate-methacrylate hydrogels. Carbohydr Polym. 2003;52:389-96.

- 156 -

Bibliography

[98] Carmeliet P. Manipulating angiogenesis in medicine. J Intern Med. 2004;255:538- 61.

[99] Bryant SJ, Davis-Arehart KA, Luo N, Shoemaker RK, Arthur JA, Anseth KS. Synthesis and characterization of photopolymerized multifunctional hydrogels: Water- soluble poly(vinyl alcohol) and chondroitin sulfate macromers for chondrocyte encapsulation. Macromolecules. 2004;37:6726-33.

[100] Padera R, Venkataraman G, Berry D, Godavarti R, Sasisekharan R. FGF- 2/fibroblast growth factor receptor/heparin-like glycosaminoglycan interactions: a compensation model for FGF-2 signaling. FASEB J. 1999;13:1677-87.

[101] Wu ZL, Zhang L, Yabe T, Kuberan B, Beeler DL, Love A, et al. The involvement of heparan sulfate (HS) in FGF1/HS/FGFR1 signaling complex. J Biol Chem. 2003;278:17121-9.

[102] Noro K. Hydrolysis of polyvinyl acetate to polyvinyl alcohol. In: Finch CA, editor. Polyvinyl alcohol, properties and applications. New York: Wiley; 1973.

[103] Finch CA. Chemical properties of polyvinyl alcohol. In: Finch CA, editor. Polyvinyl alcohol, properties and applications. New York: Wiley; 1973.

[104] Martens PJ. Hydrogel synthesis from multifunctional poly(vinyl alcohol) macromers: experimental and theoretical approaches to understanding degrading networks [Ph.D.]: University of Colorado; 2002.

[105] Peppas NA. Hydrogels in Medicine and Pharmacy. Volume 1 Fundamentals. Boca Raton, FL, USA: CRC Press Inc; 1986.

[106] Anseth KS, Metters AT, Bryant SJ, Martens PJ, Elisseeff JH, Bowman CN. In situ forming degradable networks and their application in tissue engineering and drug delivery. J Control Release. 2002;78:199-209.

[107] Baroli B. Photopolymerization of biomaterials: issues and potentialities in drug delivery, tissue engineering, and cell encapsulation applications. J Chem Technol Biotechnol. 2006;81:491-9.

[108] Mühlebach A, Müller B, Pharisa C, Hofmann M, Seiferling B, Guerry D. New water-soluble photo crosslinkable polymers based on modified poly(vinyl alcohol). J Polym Sci, Part A: Polym Chem. 1997;35:3603-11.

[109] Martens P, Holland T, Anseth KS. Synthesis and characterization of degradable hydrogels formed from acrylate modified poly(vinyl alcohol) macromers. Polymer. 2002;43:6093-100.

[110] Cavalieri F, Miano F, D'Antona P, Paradossi G. Study of gelling behavior of poly(vinyl alcohol)-methacrylate for potential utilizations in tissue replacement and drug delivery. Biomacromolecules. 2004;5:2439-46.

- 157 -

Bibliography

[111] Martens PJ, Bryant SJ, Anseth KS. Tailoring the degradation of hydrogels formed from multivinyl poly(ethylene glycol) and poly(vinyl alcohol) macromers for cartilage tissue engineering. Biomacromolecules. 2003;4:283-92.

[112] Martens P, Blundo J, Nilasaroya A, Odell RA, Cooper-White J, Poole-Warren LA. Effect of poly(vinyl alcohol) macromer chemistry and chain interactions on hydrogel mechanical properties. Chem Mater. 2007;19:2641-8.

[113] Masters KSB, Leibovich SJ, Belem P, West JL, Poole-Warren LA. Effects of nitric oxide releasing poly(vinyl alcohol) hydrogel dressings on dermal wound healing in diabetic mice. Wound Repair Regen. 2002;10:286-94.

[114] Jen AC, Wake MC, Mikos AG. Review: Hydrogels for cell immobilization. Biotechnol Bioeng. 1996;50:357-64.

[115] Yasukawa T, Kimura H, Tabata Y, Miyamoto H, Honda Y, Ikada Y, et al. Targeted delivery of anti-angiogenic agent TNP-470 using water-soluble polymer in the treatment of choroidal neovascularization. Invest Ophthalmol Vis Sci. 1999;40:2690-6.

[116] Goosen MFA, Sefton MV, Hatton MWC. Inactivation of thrombin by antithrombin III on a heparinized biomaterial. Thromb Res. 1980;20:543-54.

[117] Rollason G, Sefton MV. Inactivation of thrombin in heparin-PVA coated tubes. J Biomater Sci Polym Ed. 1989;1:31-41.

[118] Brinkman E, van der Does L, Bantjes A. Poly(vinyl alcohol)-heparin hydrogels as sensor catheter membranes. Biomaterials. 1991;12:63-70.

[119] Smith BAH, Sefton MV. Thrombin and albumin adsorption to PVA and heparin- PVA hydrogels. I. Single protein isotherms. J Biomed Mater Res. 1992;26:947-58.

[120] Cholakis CH, Sefton MV. In vitro platelet interactions with a heparin-polyvinyl alcohol hydrogel. J Biomed Mater Res. 1989;23:399-415.

[121] Godo MN, Sefton MV. Characterization of transient platelet contacts on a polyvinyl alcohol hydrogel by video microscopy. Biomaterials. 1999;20:1117-26.

[122] Smith BAH, Sefton MV. Thrombin and albumin adsorption to PVA and heparin- PVA hydrogels. 2: Competition and displacement. J Biomed Mater Res. 1993;27:89-95.

[123] Lee C-T, Kung P-H, Lee Y-D. Preparation of poly(vinyl alcohol)-chondroitin sulfate hydrogel as matrices in tissue engineering. Carbohydr Polym. 2005;61:348-54.

[124] Varshney L. Role of natural polysaccharides in radiation formation of PVA- hydrogel wound dressing. Nuclear Instruments and Methods in Physics Research Section B: Beam Interactions with Materials and Atoms. 2007;255:343-9.

[125] Mawad D, Martens PJ, Odell RA, Poole-Warren LA. The effect of redox polymerisation on degradation and cell responses to poly (vinyl alcohol) hydrogels. Biomaterials. 2007;28:947-55.

- 158 -

Bibliography

[126] Yanagishita M, Hascall VC. Cell surface heparan sulfate proteoglycans. J Biol Chem. 1992;267:9451-4.

[127] Noonan DM, Fulle A, Valente P, Cai S, Horigan E, Sasaki M, et al. The complete sequence of perlecan, a basement membrane heparan sulfate proteoglycan, reveals extensive similarity with laminin A chain, low density lipoprotein-receptor, and the neural cell adhesion molecule. J Biol Chem. 1991;266:22939-47.

[128] Lindahl U, Kusche-Gullberg M, Kjellen L. Regulated diversity of heparan sulfate. J Biol Chem. 1998;273:24979-82.

[129] Kjellen L, Lindahl U. Proteoglycans: Structures and interactions. Annu Rev Biochem. 1991;60:443-75.

[130] Iozzo RV. Series Introduction: Heparan sulfate proteoglycans: intricate molecules with intriguing functions. J Clin Invest. 2001;108:165-7.

[131] Ishihara M, Ono K. Structure and function of heparin and heparan sulfate; Heparinoid library and modification of FGF-activities. Trends Glycosci Glycotechnol. 1998;10:223-33.

[132] Iozzo RV, San Antonio JD. Heparan sulfate proteoglycans: heavy hitters in the angiogenesis arena. J Clin Invest. 2001;108:349-55.

[133] Toida T, Huang Y, Washio Y, Maruyama T, Toyoda H, Imanari T, et al. Chemical microdetermination of heparin in plasma. Anal Biochem. 1997;251:219-26.

[134] Guerrini M, Naggi A, Guglieri S, Santarsiero R, Torri G. Complex glycosaminoglycans: profiling substitution patterns by two-dimensional nuclear magnetic resonance spectroscopy. Anal Biochem. 2005;337:35-47.

[135] Bernfield M, Gotte M, Park PW, Reizes O, Fitzgerald ML, Lincecum J, et al. Functions of cell surface heparan sulfate proteoglycans. Annu Rev Biochem. 1999;68:729-77.

[136] Petitou M, Casu B, Lindahl U. 1976-1983, a critical period in the history of heparin: the discovery of the antithrombin binding site. Biochimie. 2003;85:83-9.

[137] Knox S, Merry C, Stringer S, Melrose J, Whitelock J. Not all perlecans are created equal: interactions with fibroblast growth factor (FGF) 2 and FGF receptors. J Biol Chem. 2002;277:14657-65.

[138] Guglieri S, Hricovíni M, Raman R, Polito L, Torri G, Casu B, et al. Minimum FGF2 binding structural requirements of heparin and heparan sulfate oligosaccharides as determined by NMR spectroscopy. Biochemistry. 2008;47:13862–9.

[139] Guimond S, Maccarana M, Olwin BB, Lindahl U, Rapraeger AC. Activating and inhibitory heparin sequences for FGF-2 (basic FGF). Distinct requirements for FGF-1, FGF-2, and FGF-4. J Biol Chem. 1993;268:23906-14.

- 159 -

Bibliography

[140] Folkman J, Klagsbrun M, Sasse J, Wadzinski M, Ingber D, Vlodavsky I. A heparin-binding angiogenic protein – basic fibroblast growth factor – is stored within basement membrane. Am J Pathol. 1988;130:393-400.

[141] Flaumenhaft R, Moscatelli D, Saksela O, Rifkin DB. Role of extracellular matrix in the action of basic fibroblast growth factor: Matrix as a source of growth factor for long-term stimulation of plasminogen activator production and DNA synthesis. J Cell Physiol. 1989;140:75-81.

[142] Saksela O, Moscatelli D, Sommer A, Rifkin DB. Endothelial cell-derived heparan sulfate binds basic fibroblast growth factor and protects it from proteolytic degradation. J Cell Biol. 1988;107:743-51.

[143] Rapraeger AC, Krufka A, Olwin BB. Requirement of heparan sulfate for bFGF- mediated fibroblast growth and myoblast differentiation. Science. 1991;252:1705.

[144] Yayon A, Klagsbrun M, Esko JD, Leder P, Ornitz DM. Cell surface, heparin-like molecules are required for binding of basic fibroblast growth factor to its high affinity receptor. Cell. 1991;64:841-8.

[145] Ornitz DM, Yayon A, Flanagan JG, Svahn CM, Levi E, Leder P. Heparin is required for cell-free binding of basic fibroblast growth factor to a soluble receptor and for mitogenesis in whole cells. Mol Cell Biol. 1992;12:240-7.

[146] Kan M, Wang F, Xu J, Crabb JW, Hou J, McKeehan WL. An essential heparin- binding domain in the fibroblast growth factor receptor kinase. Science. 1993;259:1918- 21.

[147] Safran M, Eisenstein M, Aviezer D, Yayon A. Oligomerization reduces heparin affinity but enhances receptor binding of fibroblast growth factor 2. Biochem J. 2000;345:107-13.

[148] Roghani M, Mansukhani A, Dell'Era P, Bellosta P, Basilico C, Rifkin DB, et al. Heparin increases the affinity of basic fibroblast growth factor for its receptor but is not required for binding. J Biol Chem. 1994;269:3976-84.

[149] Segev A, Nili N, Strauss BH. The role of perlecan in arterial injury and angiogenesis. Cardiovasc Res. 2004;63:603-10.

[150] Gitay-Goren H, Soker S, Vlodavsky I, Neufeld G. The binding of vascular endothelial growth factor to its receptors is dependent on cell surface-associated heparin-like molecules. J Biol Chem. 1992;267:6093-8.

[151] Conrad E. Commentary on heparin-like glycosaminoglycans. Angiogenesis. 1997;1:16-9.

[152] Whitelock JM, Melrose J, Iozzo RV. Diverse cell signaling events modulated by perlecan. Biochemistry. 2008;47:11174-83.

[153] Graham L, Hayward I, Underwood P. Activation of platelet heparitinase by vascular cell lysates. Biochem Mol Biol Int. 1995;37:231-7.

- 160 -

Bibliography

[154] Sanderson RD, Yang Y, Suva LJ, Kelly T. Heparan sulfate proteoglycans and heparanase - partners in osteolytic tumor growth and metastasis. Matrix Biol. 2004;23:341-52.

[155] Tae G, Scatena M, Stayton PS, Hoffman AS. PEG-cross-linked heparin is an affinity hydrogel for sustained release of vascular endothelial growth factor. J Biomater Sci Polym Ed. 2006;17:187-97.

[156] Merrill EW, Salzman EW, Wong PS, Ashford TP, Brown AH, Austen WG. Polyvinyl alcohol-heparin hydrogel "G". J Appl Physiol. 1970;29:723-30.

[157] Hanthamrongwit M, Reid WH, Grant MH. Chondroitin-6-sulphate incorporated into collagen gels for the growth of human keratinocytes: the effect of cross-linking agents and diamines. Biomaterials. 1996;17:775-80.

[158] Vasudev SC, Moses LR, Sharma CP. Covalently bonded heparin to alter the pericardial calcification. Artif Cells Blood Substit Immobil Biotechnol. 2000;28:241-53.

[159] McGonigle JS, Tae G, Stayton PS, Hoffman AS, Scatena M. Heparin-regulated delivery of osteoprotegerin promotes vascularization of implanted hydrogels. J Biomater Sci Polym Ed. 2008;19:1021-34.

[160] Elisseeff J, Anseth K, Sims D, McIntosh W, Randolph M, Langer R. Transdermal photopolymerization for minimally invasive implantation. Proc Natl Acad Sci U S A. 1999;96:3104-7.

[161] Williams CG, Malik AN, Kim TK, Manson PN, Elisseeff JH. Variable cytocompatibility of six cell lines with photoinitiators used for polymerizing hydrogels and cell encapsulation. Biomaterials. 2005;26:1211-8.

[162] Goldenberg M, Mueller KFUS. Polyvinyl alcohol derivatives containing pendant (meth)acryloyl units bound through urethane groups and crosslinked hydrogel contact lenses made therefrom. US: Ciba-Geigy Corporation; 1987.

[163] Goldenberg M, Mueller KFUS. Polyvinyl alcohol derivatives containing pendant (meth)acryloyl units bound through urethane groups and crosslinked hydrogel contact lenses made therefrom. US: Ciba-Geigy Corporation; 1988.

[164] Styan KE, Martin DJ, Poole-Warren LA. In vitro fibroblast response to polyurethane organosilicate nanocomposites. J Biomed Mater Res. 2008;86A:571-82.

[165] Zar JH. Biostatistical Analysis. 4th ed. Upper Saddle River, N.J.: Prentice Hall; 1999.

[166] Minitab I. Minitab Statistical Software. Release 15 for Windows ed. State College, Pennsylvania2006.

[167] DeMerlis CC, Schoneker DR. Review of the oral toxicity of polyvinyl alcohol (PVA). Food Chem Toxicol. 2003;41:319-26.

- 161 -

Bibliography

[168] Lee W-F, Lin W-J. Preparation and gel properties of poly[hydroxyethylmethacrylate-co-poly(ethylene glycol) methacrylate] copolymeric hydrogels by photopolymerization. Journal of Polymer Research. 2002;9:23-9.

[169] Temenoff JS, Athanasiou KA, Lebaron RG, Mikos AG. Effect of poly(ethylene glycol) molecular weight on tensile and swelling properties of oligo(poly(ethylene glycol) fumarate) hydrogels for cartilage tissue engineering. J Biomed Mater Res. 2002;59:429-37.

[170] Barile FA. Introduction to in vitro cytotoxicology: mechanism and methods. Boca Raton: CRC Press; 1994.

[171] Bryant SJ, Nuttelman CR, Anseth KS. Cytocompatibility of UV and visible light photoinitiating systems on cultured NIH/3T3 fibroblasts in vitro. J Biomater Sci Polym Ed. 2000;11:439-57.

[172] van Dijk-Wolthuis WNE, Kettenes-van den Bosch JJ, van der Kerk-van Hoof A, Hennink WE. Reaction of dextran with glycidyl methacrylate: An unexpected transesterification. Macromolecules. 1997;30:3411-3.

[173] Bryant SJ, Hauch KD, Ratner BD. Spatial patterning of thick poly(2-hydroxyethyl methacrylate) hydrogels. Macromolecules. 2006;39:4395-9.

[174] Berchtold KA, Lovell LG, Nie J, Hacioglu B, Bowman CN. The significance of chain length dependent termination in cross-linking polymerizations. Polymer. 2001;42:4925-9.

[175] Burdick JA, Lovestead TM, Anseth KS. Kinetic chain lengths in highly cross- linked networks formed by the photoinitiated polymerization of divinyl monomers: A gel permeation chromatography investigation. Biomacromolecules. 2002;4:149-56.

[176] Hacioglu B, Berchtold KA, Lovell LG, Nie J, Bowman CN. Polymerization kinetics of HEMA/DEGDMA: using changes in initiation and chain transfer rates to explore the effects of chain-length-dependent termination. Biomaterials. 2002;23:4057- 64.

[177] Wu ZL, Zhang L, Beeler DL, Kuberan B, Rosenberg RD. A new strategy for defining critical functional groups on heparan sulfate. 2002;16:539-45.

[178] Hirsh J, Anand SS, Halperin JL, Fuster V. Guide to anticoagulant therapy: Heparin: A statement for healthcare professionals from the American Heart Association. Circulation. 2001;103:2994-3018.

[179] Nasser NJ, Sarig G, Brenner B, Nevo E, Goldshmidt O, Zcharia E, et al. Heparanase neutralizes the anticoagulation properties of heparin and low-molecular- weight heparin. J Thromb Haemost. 2006;4:560-5.

[180] Lapierre F, Holme K, Lam L, Tressler RJ, Storm N, Wee J, et al. Chemical modifications of heparin that diminish its anticoagulant but preserve its heparanase- inhibitory, angiostatic, anti-tumor and anti-metastatic properties. Glycobiology. 1996;6:355-66.

- 162 -

Bibliography

[181] Ono K, Ishihara M, Ishikawa K, Ozeki Y, Deguchi H, Sato M, et al. Periodate- treated, non-anticoagulant heparin-carrying polystyrene (NAC-HCPS) affects angiogenesis and inhibits subcutaneous induced tumour growth and metastasis to the lung. Br J Cancer. 2002;86:1803-12.

[182] Tripodi A, Mannucci PM. Activated partial thromboplastin time (APTT). New indications for an old test? J Thromb Haemost. 2006;4:750-1.

[183] Tollefsen DM, Majerus DW, Blank MK. Heparin cofactor II. Purification and properties of a heparin-dependent inhibitor of thrombin in human plasma. 1982. p. 2162-9.

[184] Turnbull JE, Fernig DG, Ke Y, Wilkinson MC, Gallagher JT. Identification of the basic fibroblast growth factor binding sequence in fibroblast heparan sulfate. J Biol Chem. 1992;267:10337-41.

[185] Moy FJ, Safran M, Seddon AP, Kitchen D, Bohlen P, Aviezer D, et al. Properly oriented heparin decasaccharide-induced dimers are the biologically active form of basic fibroblast growth factor. Biochemistry. 1997;36:4782-91.

[186] Melrose J, Ghosh P. Determination of the average molecular size of glycosaminoglycans by fast protein liquid chromatography. J Chromatogr A. 1993;637:91-5.

[187] Barbosa I, Garcia S, Barbier-Chassefiere V, Caruelle J-P, Martelly I, Papy-Garcia D. Improved and simple micro assay for sulfated glycosaminoglycans quantification in biological extracts and its use in skin and muscle tissue studies. Glycobiology. 2003;13:647-53.

[188] Meng W, Dong Ki B, Jin-Ho C, Inn-Kyu K, Kyo Han K. In vitro blood compatibility of heparin-immobilized polyurethane containing ester groups in the side chain. J Mater Sci-Mater Med. 2004;15:1079-87.

[189] Kang I-K, Kwon OH, Kim MK, Lee YM, Sung YK. In vitro blood compatibility of functional group-grafted and heparin-immobilized polyurethanes prepared by plasma glow discharge. Biomaterials. 1997;18:1099-107.

[190] Maccarana M, Casu B, Lindahl U. Minimal sequence in heparin/heparan sulfate required for binding of basic fibroblast growth factor. J Biol Chem. 1993;268:23898- 905.

[191] Palacios R, Steinmetz M. IL3-dependent mouse clones that express B-220 surface antigen, contain ig genes in germ-line configuration, and generate B lymphocytes in vivo. Cell. 1985;41:727-34.

[192] Vlodavsky I, Friedmann Y, Elkin M, Aingorn H, Atzmon R, Ishai-Michaeli R, et al. Mammalian heparanase: Gene cloning, expression and function in tumor progression and metastasis. Nat Med. 1999;5:793-802.

- 163 -

Bibliography

[193] Wang D-A, Varghese S, Sharma B, Strehin I, Fermanian S, Gorham J, et al. Multifunctional chondroitin sulphate for cartilage tissue-biomaterial integration. Nature Materials. 2007;6:385-92.

[194] Tae G, Kim YJ, Choi WI, Kim M, Stayton PS, Hoffman AS. Formation of a novel heparin-based hydrogel in the presence of heparin-binding biomolecules. Biomacromolecules. 2007;8:1979-86.

[195] Hoskins R, Robb ID, Williams PA, Warren P. Phase separation in mixtures of polysaccharides and proteins. J Chem Soc, Faraday Trans. 1996;92:4515-20.

[196] Senatore F, Shankar H, Chen JH, Avantsa S, Feola M, Posteraro R, et al. In vitro and in vivo studies of heparinized-collageno-elastic tubes. J Biomed Mater Res. 1990;24:939-57.

[197] Darwis D, Yoshii F, Makuuchi K, Razzak MT. Improvement of hot water resistance of poly(vinyl alcohol) hydrogel by acetalization and irradiation techniques. Journal of Applied Polymer Science. 1995;55:1619-25.

[198] Vlodavsky I, Miao H-Q, Medalion B, Danagher P, Ron D. Involvement of heparan sulfate and related molecules in sequestration and growth promoting activity of fibroblast growth factor. Cancer Metastasis Rev. 1996;15:177-86.

[199] Jumaa H, Hendriks RW, Reth M. B cell signaling and tumorigenesis. Annual Review of Immunology. 2005;23:415-45.

[200] Levy-Adam F, Abboud-Jarrous G, Guerrini M, Beccati D, Vlodavsky I, Ilan N. Identification and characterization of heparin/heparan sulfate binding domains of the endoglycosidase heparanase. J Biol Chem. 2005;280:20457-66.

[201] Reiland J, Sanderson RD, Waguespack M, Barker SA, Long R, Carson DD, et al. Heparanase degrades syndecan-1 and perlecan heparan sulfate: Functional implications for tumor cell invasion. J Biol Chem. 2004;279:8047-55.

[202] Nakajima M, Irimura T, Di Ferrante N, Nicolson GL. Metastatic melanoma cell heparanase. Characterization of heparan sulfate degradation fragments produced by B16 melanoma endoglucuronidase. J Biol Chem. 1984;259:2283-90.

[203] Bame KJ. Heparanases: endoglycosidases that degrade heparan sulfate proteoglycans. Glycobiology. 2001;11:91R-8.

[204] Hulett MD, Freeman C, Hamdorf BJ, Baker RT, Harris MJ, Parish CR. Cloning of mammalian heparanase, an important enzyme in tumor invasion and metastasis. Nat Med. 1999;5:803-9.

[205] Freeman C, Parish CR. A rapid quantitative assay for the detection of mammalian heparanase activity. Biochem J. 1997;325:229-37.

[206] Gilat D, Hershkoviz R, Goldkorn I, Cahalon L, Korner G, Vlodavsky I, et al. Molecular behavior adapts to context: heparanase functions as an extracellular matrix-

- 164 -

Bibliography degrading enzyme or as a T cell adhesion molecule, depending on the local pH. J Exp Med. 1995;181:1929-34.

[207] Ehrbar M, Rizzi SC, Schoenmakers RG, San Miguel B, Hubbell JA, Weber FE, et al. Biomolecular hydrogels formed and degraded via site-specific enzymatic reactions. Biomacromolecules. 2007;8:3000-7.

[208] Takagaki K, Kakizaki I. Degradation of Glycosaminoglycans. In: Kamerling JP, editor. Comprehensive Glycoscience. Oxford: Elsevier; 2007. p. 171-92.

[209] Whitelock JM, Murdoch AD, Iozzo RV, Underwood PA. The degradation of human endothelial cell-derived perlecan and release of bound basic fibroblast growth factor by stromelysin, collagenase, plasmin, and heparanases. J Biol Chem. 1996;271:10079-86.

[210] Farndale RW, Buttle DJ, Barrett AJ. Improved quantitation and discrimination of sulphated glycosaminoglycans by use of dimethylmethylene blue. Biochim Biophys Acta. 1986;883:173-7.

[211] Behzad F, Brenchley PEC. A multiwell format assay for heparanase. Anal Biochem. 2003;320:207-13.

[212] Freeman C, Parish CR. Human platelet heparanase: purification, characterization and catalytic activity. Biochem J. 1998;330:1341-50.

[213] Sigma-Aldrich. Product information sheet - Heparin sodium from porcine intestinal mucosa, Sigma prod. nos. H3393 and H9399. St Louis1996.

[214] Rosenberg RD, Jordan RE, Favreau LV, Lam LH. Highly active heparin species with multiple binding sites for antithrombin. Biochem Biophys Res Commun. 1979;86:1319-24.

[215] Pikas DS, Li J-p, Vlodavsky I, Lindahl U. Substrate specificity of heparanases from human hepatoma and platelets. J Biol Chem. 1998;273:18770-7.

[216] Pye DA, Gallagher JT. Monomer complexes of basic fibroblast growth factor and heparan sulfate oligosaccharides are the minimal functional unit for cell activation. J Biol Chem. 1999;274:13456-61.

[217] Sigma-Aldrich. Certificate of Analysis - Heparin sodium salt from porcine intestinal mucosa, Grade I-A, ≥180 USP units/mg. St Louis2005.

[218] Sigma-Aldrich. Certificate of Analysis - Heparin sodium salt from porcine intestinal mucosa, mol wt 4,000-6,000 Da St Louis2003.

[219] Goodger SJ, Robinson CJ, Murphy KJ, Gasiunas N, Harmer NJ, Blundell TL, et al. Evidence that heparin saccharides promote FGF2 mitogenesis through two distinct mechanisms. J Biol Chem. 2008;283:13001-8.

- 165 -

Bibliography

[220] Tans G, Rosing J, Thomassen M, Heeb M, Zwaal R, Griffin J. Comparison of anticoagulant and procoagulant activities of stimulated platelets and platelet-derived microparticles. Blood. 1991;77:2641-8.

[221] Oldberg A, Heldin CH, Wasteson A, Busch C, Hook M. Characterization of platelet endoglycosidase degrading heparin-like polysaccharides. Biochemistry. 1980;19:5755-62.

[222] Iozzo RV, Cohen IR, Grässel S, Murdoch AD. The biology of perlecan: the multifaceted heparan sulphate proteoglycan of basement membranes and pericellular matrices. Biochem J. 1994;302:625-39.

[223] Martens P, Bryant SJ, Anseth KS. Photopolymerization of poly(vinyl alcohol) and poly(ethylene glycol)-based macromers to produce cross linked, degradable hydrogels with controlled transport properties. Abstracts of Papers, 224th ACS National Meeting, Boston, MA, United States, August 18-22, 2002. 2002:MACR-014.

[224] Murphy CM, Haugh MG, O'Brien FJ. The effect of mean pore size on cell attachment, proliferation and migration in collagen-glycosaminoglycan scaffolds for bone tissue engineering. Biomaterials. 2010;31:461-6.

[225] Dagalakis N, Flink J, Stasikelis P, Burke JF, Yannas IV. Design of an artificial skin. Part III. Control of pore structure. J Biomed Mater Res. 1980;14:511-28.

[226] Bryant SJ, Cuy JL, Hauch KD, Ratner BD. Photo-patterning of porous hydrogels for tissue engineering. Biomaterials. 2007;28:2978-86.

- 166 -