Elastase Responsive Hydrogel Dressing for Chronic Wounds

EEllaassttaassee RReessppoonnssiivvee HHyyddrrooggeell DDrreessssiinngg ffoorr CChhrroonniicc WWoouunnddss

A thesis submitted to the University of Manchester for the degree of Doctor of Philosophy in the Faculty of Engineering

and Physical Sciences

2010

Nurguse Bibi

School of Materials

Contents

2

CCOONNTTEENNTTSS

Contents 2

List of Tables 7

List of Figures 8

Abstract 12

Declaration 13

Copyright Statement 14

Acknowledgements 15

Dedication 16

Abbreviations 17

Conference Presentations 24

CCHHAAPPTTEERR 11 II nnttrroodduuccttiioonn 25

1.1 Objective 28

1.2 Aims of the project 29

1.3 Scope of Thesis 30

CCHHAAPPTTEERR 22 LLii tteerraattuurree RReevviieeww –– WWoouunndd HHeeaall iinngg && MMaannaaggeemmeenntt ooff CChhrroonniicc WWoouunnddss 32

2.1 ANATOMY & PHYSIOLOGY OF THE SKIN 33

2.1.1 Anatomy of the skin 33

2.1.1.1 Epidermis 33

2.1.1.2 Dermis 35

2.1.2 Physiological function of the skin 35

2.2 WOUND HEALING 37

2.2.1 Acute and chronic wound healing 37

2.2.2 Phases of wound healing and the immune response 38

2.2.2.1 Hemostasis 38

2.2.2.2 Inflammation 41

2.2.2.3 Proliferation 44

2.2.2.4 Remodelling 46

Contents

3

2.2.3 Role of enzymes in wound healing 46

2.2.3.1 Serine proteases 48

2.2.3.2 Matrix Metalloproteinases 54

2.3 TYPES OF CHRONIC WOUNDS 62

2.4 GRADIENTS OF CHRONIC WOUNDS 62

2.4.1 Oxygen 63

2.4.2 Temperature 63

2.4.3 pH 63

2.5 ROLES OF ENZYMES IN CHRONIC WOUNDS 64

2.6 MANAGEMENT OF WOUND HEALING 67

2.6.1 Debridement of chronic wounds 68

2.6.2 Moist wound healing 68

2.6.3 Management of chronic wounds using hydrogel dressings 69

2.6.3.1 Passive/ Non-responsive hydrogel dressings 70

2.6.3.2 Active/ Responsive hydrogel dressings 71

2.6.3.3 Protease-modulating hydrogel dressings 75

2.7 SUMMARY 88

CCHHAAPPTTEERR 33 FFuunnccttiioonnaall iissaattiioonn ooff PPEEGGAA PPaarrttiicclleess 90

3.1 INTRODUCTION 91

3.2 OBJECTIVES 94

3.3 MATERIALS & METHODS 94

3.3.1 Materials 94

3.3.2 Methods 95

3.3.2.1 Fourier-transform infrared spectroscopy (FT-IR) 95

3.3.2.2 Fmoc solid phase peptide synthesis 96

3.3.2.3 Kaiser test 99

3.3.2.4 Fmoc loading 99

3.3.2.5 Isoelectric focusing and silver staining 101

3.3.2.6 Cleaving functionalised PEGA particles with proteases 102

3.3.2.7 Swelling 103

3.3.2.8 Statistics 104

Contents

4

3.4 RESULTS & DISCUSSION 104

3.4.1 FT-IR of PEGA particles 104

3.4.2 Homogenous loading of solid-phase peptide synthesis 109

3.4.3 Isoelectric point of elastases 112

3.4.4 Designing of the elastase responsive ECPs coupled to PEGA 114

3.4.4.1 Selective design of the ERS responsive to elastase 116

3.4.4.2 Selective hydrolysis of Fmoc-tripeptide-PEGA particles

by proteases 117

3.4.4.3 Selective hydrolysis of the CMR from Fmoc-tripeptide-

PEGA particles by proteases 122

3.4.4.4 Selective hydrolysis of Fmoc-dipeptide-PEGA particles

by proteases 124

3.4.5 Swelling analysis 129

3.4.5.1 Swelling behaviour of un-cleaved Fmoc-tripeptide-

PEGA particles 129

3.4.5.2 Swelling behaviour of cleaved Fmoc-tripeptide-PEGA

particles 135

3.5 CONCLUSION 139

CCHHAAPPTTEERR 44 SSeelleeccttiivvee EEnnttrraappmmeenntt ooff EEllaassttaassee iinnttoo PPEEGGAA PPaarr ttiicclleess 140

4.1 INTRODUCTION 141

4.2 OBJECTIVES 146


4.3.1 Materials 146

4.3.2 Methods 147

4.3.2.1 Fmoc SPPS and Kaiser test 147

4.3.2.2 Potassium phosphate buffer 147

4.3.2.3 Eznyme hydrolysis using HPLC 147

4.3.2.4 Dansyl chloride staining 148

4.3.2.5 SDS-Page 149

4.3.2.6 Accessibility of FITC-dextran 149

4.3.2.7 Accessibility FITC-elastase 150

Contents

5

4.3.2.8 Quantification of TPM micrographs 151

4.3.2.9 Fluorescence substrate assay 151



4.4.1 Selective hydrolysis of ECPs by elastase at high ionic strength as a

function of pH 153

4.4.2 Fluorescence Studies 154

4.4.2.1 Diffusion of elastase via dansyl chloride 154

4.4.2.2 Accessibility of a FITC-labelled dextran into PEGA

particles 163

4.4.2.3 FITC-elastase 168


4.5 CONCLUSION 176

CCHHAAPPTTEERR 55 RReemmoovvaall ooff FFiibbrroobbllaasstt EEllaassttaassee AAccttiivvii ttyy BByy CChhaarrggeedd PPEEGGAA PPaarrttiicclleess 177

5.1 INTRODUCTION 178

5.2 OBJECTIVES 180


5.3.1 Materials 181

5.3.2 Methods 181

5.3.2.1 Fmoc SPPS 181

5.3.2.2 Cell culture of human dermal fibroblast (HDF) 181

5.3.2.3 Activation of HDF cells by IL-1β to express

HDF-elastase 182

5.3.2.4 Elastase treated with functionalised PEGA particles 183




5.4.1 IL-1β induced HDF-elastase expression in HDF cells 184

5.4.2 Reduction of HDF-elastase activity by PEGA particles 195

5.5 CONCLUSION 200

Contents

6

CCHHAAPPTTEERR 66 CCoonncclluussiioonnss && FFuurrtthheerr SSttuuddiieess 202

6.1 CONCLUSIONS 203

6.1.1 Limitations of the studies 205

6.2 FURTHER STUDIES 207

CCHHAAPPTTEERR 77 RReeffeerreenncceess 209

AAPPPPEENNDDIIXX II 232

AAPPPPEENNDDIIXX IIII 237

AAPPPPEENNDDIIXX IIIIII 247

WWoorrdd CCoouunntt

Including references: 71, 993

Excluding references: 64, 306

List of Tables

7

LLIISSTT OOFF TTAABBLLEESS

Table 1. Layers of the epidermis. 34

Table 2. Layers of the dermis. 35

Table 3. Summary – The Physiological function of the skin. 36

Table 4. Human proteases which express elastinolytic activity. 51

Table 5. Systemic and alarm proteinase inhibitors to control HNE activity. 53

Table 6. Summary for the function of human matrilysins, MT-MMPs and other MMPs during in the wound healing process.

60

Table 7. Types of Chronic wounds. 62

Table 8. Application of protease-modulating hydrogel dressings for chronic wounds.

76

Table 9. Washing steps during SPPS. 96

Table 10. Colour observation of PEGA particles and surrounding solution after Kaiser test.

99

Table 11. Assignments of FTIR frequencies of unmodified PEGA(800 and 1900) particles.

106

Table 12. Types of conformational amide bands for FTIR frequencies.

108

Table 13. ECPs coupled to PEGA particles. 115

Table 14. Percentage decrease of HDF-elastase activity in SMF compared to DMEM + 10% FBS for both untreated and treated HDF cells with or without of IL-1β

190

Table 15. Statistical differences and similarities between the elastase activity of PPE in DMEM depending on both the concentration of FBS and temperature

191

Table 16. Statistical comparisons of HDF-elastase activity in DMEM (SFM, or supplemented with 10% or 25% FBS) using one-way ANOVA

194

Table 17. Substrate specificity of proteolytic proteases involved in the process of wound healing

232

Table 18. Passive hydrogel dressings designed to treat chronic wounds 237

List of Figures

8

LLIISSTT OOFF FFIIGGUURREESS

Figure 1. The molecular structure of PEGA particles. 29

Figure 2. A schematic diagram exhibit a deep chronic wound being healed after treatment with PEGA particles.

29

Figure 3. Swelling to collapse of PEGA particles. 30

Figure 4. Events of acute and chronic wound healing. 38

Figure 5. Formation of thrombin and fibrin. 39

Figure 6. Fibrinolytic pathway. 40

Figure 7. General mechanism of leukocyte adhesion to the endothelial cells, mediated by the cell-cell adhesion mechanism of selectins/ integrin.

42

Figure 8. The hydrolysis of peptide substrate via the serine proteases (thrombin, plasmin or elastase).

49

Figure 9. Multifunctional roles of elastase during the wound healing process. 54

Figure 10. The conformational domains of human MMPs. 55

Figure 11. Activation of MMPs via the ‘cysteine-switch’ mechanism. 56

Figure 12. Reciprocal interpretation for the role of plasmin, elastase and MMPs in achieving ECM degradation during the wound healing process.

61

Figure 13. The‘vicious elastase cycle’ causes extensive tissue/ ECM degradation promoting chronic inflammation via a positive feedback mechanism within chronic wounds.

66

Figure 14. Mode of action for OxyzymeTM hydrogel dressing. 73

Figure 15. Mechanism of Cadesorb. 83

Figure 16. Chemical structure of the MMP-inhibitor, bisphosphonate. 86

Figure 17. Bisphosphonate-functionalised hydrogels synthesised via Schotten-Baumann reaction.

87

Figure 18. Mechanism for the functionalisation of unmodified PEGA particles with enzyme cleavable peptides (ECPs) using Fmoc SPPS.

97

Figure 19. Chemical structures of uncharged (neutral) and charged Fmoc-ECPs coupled to PEGA particles.

98

Figure 20. Standard curve for the absorbance (301nm) of Fmoc group. 100

Figure 21. Standard curve for the area of Fmoc group (301 nm) via HPLC 102

Figure 22. FTIR spectrum of unmodified PEGA(800 and 1900) particles and the molecular structure of PEGA.

105

Figure 23. Mechanism of ninhydrin with free amino groups on PEGA particles via imine formation.

110

List of Figures

9

Figure 24. Colorimetric observations of the Kaiser test. 111

Figure 25. Total net charge of elastase specifically PPE (EC 3.4.21.36) within the pH range of 1 – 14.

113

Figure 26. Schematic illustration of designing elastase responsive ECPs that accommodate the environment of chronic wounds both above and below pH 8.31.

114

Figure 27. Selective hydrolysis of neutral ECPs (Fmoc-dipeptides) coupled to PEGA800 particles by elastase.

117

Figure 28. Schematic illustration comparing the charges of thermolysin and elasatse within the pH range of 6.0 – 9.0.

118

Figure 29. A chemical reaction displaying the selective hydrolysis of Fmoc-tripeptide-PEGA particles by a protease.

118

Figure 30. Selective hydrolysis of Fmoc-X-Ala-Ala-PEGA(800,1900) particles by thermolysin and elastase under the influence of pH.

119

Figure 31. Selective hydrolysis of the CMR from Fmoc-X-Ala-Ala-PEGA(800,1900) particles by thermolysin and elastase.

123

Figure 32. The chemical reaction of cleaving of Fmoc-dipeptide-PEGA particles (RA, EA, GA) by proteases.

125

Figure 33. Selective hydrolysis of Fmoc-X-Ala-PEGA(800,1900) particles by thermolysin and elastase at different pH values.

126

Figure 34. Swelling behaviour of functionalised Fmoc-X-Ala-Ala-PEGA1900 particles as a function of pH (a: 7.0, b: 8.0 and c: 9.0) and ionic strength (ranging from 0.001M – 0.2M) in potassium phosphate buffer.

131

Figure 35. Swelling behaviour of functionalised Fmoc-X-Phe-Phe-PEGA1900 particles as a function of pH (a: 7.0, b: 8.0 and c: 9.0) and ionic strength (ranging from 0.001M – 0.2M) in potassium phosphate buffer.

132

Figure 36. Fmoc-tripeptide-PEGA1900 particles (a) cleaved by protease to generate cleaved products (b-c).

135

Figure 37. Swelling behaviour of cleaved product (+)Ala-PEGA1900 and the percentage decrease/increase in swelling when compared to the un-cleaved Fmoc-X-Ala-Ala-PEGA1900 particles.

136

Figure 38. Swelling behaviour of cleaved product (+)Phe-PEGA1900 and the percentage decrease/increase in swelling of the cleaved Fmoc-X-Phe-Phe-PEGA1900 particles.

137

Figure 39. Jablonski diagram and Stokes shift of fluorescence. 142

Figure 40. Jablonski diagram comparing the absorption of two-photon against one-photon.

144

Figure 41. Chemical reaction of dansyl chloride with cleaved PEGA particles. 148

Figure 42. Chemical reaction of staining elastase with the base fluorescein molecule, FITC.

150

Figure 43. The mechanism for the cleaving of the fluorescence-quenched substrate: MeOSuc-Ala-Ala-Pro-Val-AMC by elastase.

152

List of Figures

10

Figure 44. Selective hydrolysis of Fmoc-X-Ala-Ala-PEGA1900 particles and Fmoc-X-Phe-Phe-PEGA1900 particles by elastase at high ionic strength under the influence of pH.

154

Figure 45. TPM micrographs (top) and the average fluorescence graph (bottom) depicting the fluorescence labelling after un-cleaved and cleaved Fmoc-X-Phe-Phe-PEGA1900 particles (RFF, EFF and GFF) were stained with dansyl chloride at pH 8 (0.1 M).

155

Figure 46. TPM micrographs (top) and the average fluorescence (bottom) illustrating the fluorescence observed after un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles (RAA, EAA and GAA) were stained with dansyl chloride at pH 8 (0.1 M).

156

Figure 47. The selective diffusion of elastase into Fmoc-X-Ala-Ala-PEGA1900 particles (RAA, EAA and GAA) followed by the selective hydrolysis of the corresponding ECPs in potassium phosphate buffer at pH 8 and 0.001M monitored over the course of 180 minutes.

159

Figure 48. TPM micrographs demonstrating the diffusion of elastase after Fmoc-X-Ala-Ala-PEGA1900 particles were cleaved with elastase for 3 hours at pH 7 and pH 9 (0.1 M).

160

Figure 49. Treating both un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles (0.1 M, pH 7.0) with Kaiser test.

161

Figure 50. Comparing the pixel fluorescence intensity at pH 7 (0.1 M) after unmodified and ionic PEGA1900 particles were stained with dansyl chloride.

162

Figure 51. TPM micrographs and pixel intensity graphs demonstrating the penetration of a 20 kDa FITC-labelled dextran into both un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles at 0.1 M (pH 8).

165

Figure 52. TPM micrographs (left and middle) and pixel intensity graphs (right) demonstrating the accessibility and diffusion of FITC-labelled 20 kDa dextran into unmodified PEGA1900 particles at an ionic strength of 0.1 M and 0.001 M for the duration of 10 minutes.

166

Figure 53. TPM micrographs and pixel intensity graphs demonstrating the penetration of a 20 kDa FITC-labelled dextran into both un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles at 0.001M (pH 8).

167

Figure 54. TPM micrographs (top) and the average fluorescence graph (bottom) depicting the real-time penetration of FITC-elastase into Fmoc-X-Ala-Ala-PEGA1900 particles in PBS buffer (0.15 M and pH 7.4).

169

Figure 55. TPM micrographs (top) and the average fluorescence graph (bottom) depicting the real-time penetration of FITC-elastase into GAA particles at 0.1 M (pH 7.0).

170

Figure 56. Optimisation of the fluorescence substrate assay using Michaelis-Menten kinetics.

172

Figure 57. Rates of residual elastase activity of PPE after sample fluids containing elastase were treated with Fmoc-X-Ala-Ala-PEGA particles at 0.1 M and at various pH values: pH 7 (a), pH 8 (b) and pH 9 (c).

174

Figure 58. Photomicrographs of HDF cultures (passage 10) showing the effect of IL-1β on the proliferation of HDFa cells after 1 day and 7 days

185

List of Figures

11

growth in DMEM + 10% FBS..

Figure 59. Rate of HDF-elastase activity expressed by HDF cells in DMEM + 10% FBS.

186

Figure 60. Photomicrographs of lysing HDF cells depending on the surrounding media after 5 days growth.

187

Figure 61. Rate of HDF-elastase activity expressed by HDF cells in SFM. 189

Figure 62. Variation of elastase activity in DMEM supplemented with or without FBS against temperature.

191

Figure 63. Rate of HDF-elastase activity expressed by HDF cells in DMEM + 25% FBS.

192

Figure 64. Comparisons between the initial rate for HDF-elastase activity in DMEM + 25% FBS versus both SFM and DMEM + 10% FBS.

194

Figure 65. Rates of residual elastase activity remaining in sample fluids (at physiological pH 7.4 and ionic strength 0.15M) after the removal of functionalised PEGA particles (RAA, EAA, GAA).

196

Figure 66. Rates of residual HDF-elastase activity remaining in sample fluids after treatment with functionalised PEGA particles (RAA, EAA, GAA) swollen in DMEM + 25% FBS.

199

Figure 67. Diffusion of an enzyme into PEGA particles via a ‘ring-effect’. 248

Figure 68. Separation of elastase by SDS-page via 1-D electrophoresis. 249

12

AABBSSRRAACCTT

Chronic wounds are a major financial and clinical burden causing the deaths of millions per year. Over expression of elastase is well documented as the main culprit that delays the normal wound repair process within chronic wounds. The aim of this thesis is to design a responsive chronic wound dressing based on the hydrogel polymer, PEGA (polyethylene glycol acrylamide) in the form of particles to mop-up excess elastase by exploiting polymer collapse in response to elastase hydrolytic activity within sample fluids mimicking the environment of chronic wounds.

PEGA particles were functionalised with enzyme cleavable peptides (ECPs) containing charged residues. Upon cleavage the charge balance changes, causing polymer swelling and consequent elastase entrapment. The pH range of chronic wounds is reported in the range of 5.45 – 8.65. Due to its pI which is around 8.3, within this range elastase exist both in its cationic and anionic forms. To accommodate a hydrogel dressing that could selectively entrap excess elastase both in its cationic and anionic, oppositely charged ECPs were designed. In its cationic form, elastase was found to have a high preference of cleaving ECPs and penetrating into PEGA particles bearing negative charges. In contrast, in its anionic form the opposite effect was observed, wherein elastase preferred to cleave ECPs and penetrate PEGA particles bearing positive charges. The diffusion, accessibility and entrapment of elastase into functionalised PEGA particles was explored using various fluorescence microscopy techniques. Removal of the charged residue by elastase showed a reduction in particle swelling causing the pores of PEGA particles to become restricted. In this manner, cleaved PEGA particles prevented the accessibility of molecules with a molecular weight as low as 20 kDa into the cleaved PEGA particles. Since elastase has a molecular weight of 25.9 kDa the collapsing of the pores within PEGA particles entrapped elastase inside the interior of cleaved PEGA particles. In its cationic form (at pH 7.4) elastase was found to penetrate and become trapped more into both negative and positive PEGA particles compared to neutral particles. The negative particles were shown to trapped cationic elastase within 2 minutes compared to the positive particles. In contrast, the neutral particles failed to retain and encapsulate elastase as the fluorescence inside the neutral particles was found to decrease. Coinciding with these observations, after sample fluids containing elastase were treated with functionalised PEGA particles, the residual elastase activity in sample fluids was reduced more by the charged PEGA particles compared to neutral particles. The cell culture studies demonstrated that the elastase activity observed in human dermal fibroblasts (HDF) was also reduced more by the charged particles compared to the neutral particles. However, the positive particles were found to significantly reduced HDF-elastase activity compared to both the negative and neutral PEGA particles. Overall, this thesis exemplifies that on the basis of charge selective cleaving of ECPs coupled to PEGA particles can be exploited to selectively remove excess proteases such as elastase from sample fluids mimicking the environment of chronic wounds.

13

DDEECCLLAARRAATTIIOONN

No portion of the work referred to in this dissertation has been submitted in support of

an application for another degree or qualification of this or any other university or other

institution of learning.

14

CCOOPPYYRRIIGGHHTT SSTTAATTEEMMEENNTT

i. The author of this thesis (including any appendices and/or schedules to this thesis)

owns certain copyright or related rights in it (the “Copyright”) and s/he has given The University of Manchester certain rights to use such Copyright, including for administrative purposes.

ii. Copies of this thesis, either in full or in extracts and whether in hard or electronic

copy, may be made only in accordance with the Copyright, Designs and Patents Act 1988 (as amended) and regulations issued under it or, where appropriate, in accordance with licensing agreements which the University has from time to time. This page must form part of any such copies made.

iii. The ownership of certain Copyright, patents, designs, trade marks and other

intellectual property (the “Intellectual Property”) and any reproductions of copyright works in the thesis, for example graphs and tables (“Reproductions”), which may be described in this thesis, may not be owned by the author and may be owned by third parties. Such Intellectual Property and Reproductions cannot and must not be made available for use without the prior written permission of the owner(s) of the relevant Intellectual Property and/or Reproductions.

iv. Further information on the conditions under which disclosure, publication and

commercialisation of this thesis, the Copyright and any Intellectual Property and/or Reproductions described in it may take place is available in the University IP Policy (see http://documents.manchester.ac.uk/DocuInfo.aspx?DocID=487), in any relevant Thesis restriction declarations deposited in the University Library, The University Library’s regulations (see http://www.manchester.ac.uk/library/aboutus/ regulations) and in The University’s policy on Presentation of Theses.

15

AACCKKNNOOWWLLEEDDGGEEMMEENNTTSS

I would like to acknowledge and thank first and foremost Professor Rein Ulijn, Dr Julie

Gough and Dr David Farrar for giving me the opportunity to carryout this project. Rein

and Julie are further thanked for the continuous support, understanding and help during

this project; together with their inspiration to expand my knowledge on developing this

idea.

EPSRC and Smith & Nephew are gratefully acknowledged for sponsoring this project

which also assisted in helping me to attend various conferences to present my research to

an international audience. The Biochemical Society is thanked for the full exclusive travel

fund along with the MSCWU team who selected me to attend the Microspectroscopy

FEBS advance course (Netherlands). The MRS committee are thanked for the EPSRC

travel grant that enable me to attend the 2007 MRS Spring Meeting in San Francisco.

Robert Fernandez (FLS) is especially thanked for helping me to obtained images from 2-

photon microscope. Within School of Materials: Olwen Richert is thanked for her help

and kindness of personal and academic related matters; Francis Carabine and Shirley are

thanked for their help throughout the biomaterial labs; and Polly Crook for technical

advice on utilising equipment for studying wet-chemistry of PEGA particles. All

members of the biomaterial group are thanked for the friendship and advice throughout

this research, especially: Rachael Saunders (fluorescence plate reader), Mi Zhou (cell

culturing of HDF), Brian Cousins (providing an insight into SPSS) and Alison Patrick

(assistance with software packages, help and advice). The following people are thanked

including those mentioned above as all them in their own way helped to keep my spirits

up: Honglei Qu, Tom McDonald, Simon Todd, Rumana Rashid, Andrew Hirst, Claire

Tang, Ayeesha Mujeeb, Deepak Kalasker, Vineetha Jayawarna, Naheed Ali, Roukaya

Belkharchouche including members from E15. Karen Morgan, Professor Terry Brown

and Dr Jaleel Miyan are thanked for their kindness and continuous professional advice

since my undergraduate studies at UMIST.

I would like to thank Dr Sarah Herrick and Dr Iain Gibson for the interesting discussion

during my viva of which I thoroughly enjoyed!

Finally, and not forgotten I would like to thank my family and friends for their

continuous encouragement. My family are graciously thanked for their constant

inspiration and for their unfailing love and support; I definitely would have not got this

far without your honourable help throughout my entire career.

16

I dedicate this thesis to my father, mother, brother & sisters;

Life is absolutely meaningless without you…

Abbreviations

17

AABBBBRREEVVIIAATTIIOONNSS

α Alpha

α1-PI antiproteinase inhibitor

α2-MG α2-macroglobulin

β Beta

λ Wavelength

↑ Increase

+ve Positive

–ve Negative

1st First

2nd Second

3D Three dimensional

3rd Third

ACN Acetonitrile

ADAMs A disintegrin and metalloprotease

Ag Silver

AgNO3 silver nitrate

a.k.a Also known as

Ala Alanine (A)

AMC 7-amino-4-methyl coumarin

ANOVA Analysis of variance

Arg Arginine (R)

Asn Asparagine (N)

Asp Aspartic acid (D)

AT III Anti-thrombin III

bFGF Basic fibroblast growth factor

BM Basement membrane

BNF British National Formulary

BP(s) Bisphosphonate(s)

Ca Calcium

CAMs Cell adhesion molecules

CHAPS 3-[(3-Cholamidopropyl)dimethylammonio]-1-propanesulfonate

CMC Carboxymethylcellulose

CMR Charge modified residue: Arg(+) (positive); Glu(–) (negative) and neutral (Gly)

CO2 Carbon dioxide

Abbreviations

18

COOH Carboxyl group

Cys Cysteine (C)

CWFs Chronic wound fluids

DAG Diacylglycerol

DFUs Diabetic foot ulcers

dH2O Deionised water

DH2O Distilled water for cell culture

DHBA 2,3-dihydroxybenzoic acid

DHB-MS Modified gelatin microspheres

DIC Di-isopropylcarbodimide

DMEM Dulbecco’s modified eagle medium

DMF Dimethylformanide

DNA Deoxyribronucleic acid

Dnp Dinitrophenyl

Dpa N3-dnp-L-2,3-diaminopropionic acid

EAA Fmoc-X-Ala-Ala-PEGA, where X: Glu(–)

ECM Extracellular matrix

E.coli Eschericia coli

ECL Enzyme cleavable linkers

ECPs Enzyme cleavable peptides

EDTA Na2+ Diaminoethanetetra – acetic acid disodium salt

EFF Fmoc-X-Phe-Phe-PEGA, where X: Glu(–)

EGF Epidermal growth factor

ESL-1 E-selectin ligand-1

ERS Enzyme recognition sequence

ES Enzyme substrate

EU Europeon Union

FBS Fetal bovine serum

FDPs Fibrin degradation products

FITC Fluorescein-isothiocyanate

Fmoc/ FMOC Fluorenylmethoxycarbonyl

FW First Water

FTIR Fourier-transform infrared spectroscopy

GAA Fmoc-X-Ala-Ala-PEGA, where X: Gly

GAGs Glycosaminoglycans

GFF Fmoc-X-Phe-Phe-PEGA, where X: Gly

Glu Glutamic acid (E)

Abbreviations

19

Gly Glycine (G)

GMAWCP The Global Market for Advanced Wound Care Products

GPCR(s) G-protein coupled receptor(s)

H+ Hydrogen ions

H2O Water

H2O2 Hydrogen peroxide

HCl Hydrochloric acid

HDF Human dermal fibroblast

HEMA Hydroxylethyl methacrylate

His Histidine (H)

HME Human macrophage elastase

HNE Human neutrophil elastase

HOBt Hydroxybenzotriazole

HPLC High Performance Liquid Chromatography

hr-1 Per hour

HRT Hydration Response Technology

IEF Iso-electric focusing

Ig Immunoglobulin

IgSF Immunoglobulin Super family

IL Interleukin

IL-1 Interleukin-1

IL-1α Interleukin-1 alpha

IL-1β Interleukin-1 beta

IL-2 Interleukin-2

IL-6r Interleukin-6 receptor

IL-8 Interleukin-8

Ile Isoleucine (I)

IR Infrared

J&J Johnson & Johnson

K1–4 Mini-plasminogen, angiostatin

KCN Potassium cynanide

K. pneumoniae Klebsiella pneumoniae

KGF Keratinoctye growth factor

KM Michaelis-Menten constant

Leu Leucine (L)

LNE Leukocyte neutrophil elastase

Lys Lysine (K)

Abbreviations

20

LVDP Leucine – Valine – Aspartic acid – Proline

MeOH Methanol

MeOSuc-AAPV-AMC Methyl succiyl-alanyl-alanyl-proline-valine-amino methyl coumarin

MeOSuc-AAPV-pNA Methyl succiyl-alanyl-alanyl-proline-valine-petidyl nitroanilide

Met Methionine (M)

MI Metal ionogen, same as PHI

MMP(s) Matrix metalloproteinase(s)

MMP-1 Collagenase-1, interstitial collagenase

MMP-2 Gelatinase A, 72-kDa gelatinase, Type IV collagenase

MMP-3 Stromelysin-1, Transin, CAP, proteoglycanase

MMP-7 Matrilysin-1

MMP-8 Collagenase-2, neutrophil collagenase

MMP-9 Gelatinase B, 92-kDa gelatinase, Type V collagenase

MMP-10 Stromelysin-2, Transin-2

MMP-11 Stromelysin-3, Furin motif

MMP-12 Macrophage metalloelastase, metalloelastase, Human macrophage elastase (HME)

MMP-13 Collagenase-3, Rat interstital collagenase

MMP-14 Membrane-type MMP-14 (MT-MMP-14), MT1-MMP



MMP-20 Enamelysin



MMP-26 Matrilysin-2

MMP-28 Epilysin

MS Microspheres

MT-MMPs membrane-type MMPs

Na Sodium

Na2CO3 Sodium carbonate

NaOH Sodium hydroxide

NaAMPS Sodium salt of 2-acrylamido-2-methylpropanesulphonic acid

NaSPA Acrylic acid (3-sulphopropyl) ester sodium salt)

NE Neutrophil elastase

NH2 Amino group or amine

NSD Not significantly different, p > 0.05

OH Hydroxyl group

Abbreviations

21

ORC Oxidised regenerated cellulose

OtBu tert-butyl ester

P1–P1′ scissile P1–P1′ bond

PAs Plasminogen activators

P. aeruginosa Pseudomonas aeruginosa

PAIs Plasminogen activator inhibitors

PAs Plasminogen activators

Pbf, pbf pentamethyldihydrobenzofuran-5-sulfonyl

PBS, DPBS Phosphate buffer saline

PC Protein C

PDAF Platelet-derived angiogenic factor

PDGF Platelet-derived growth factor

PDGF-β,β Platelet-derived growth factor-beta, beta

PEG Polyethylene glycol

PEGA Poly (ethylene) glycol acrylamide

Phe Phenylalanine (F)

pH Potential hydrogen

pHEMA Poly(hydroxylethyl methacrylate)

PHI Polyhydrated ionogens

pI Isoelectric point

PIP Phosphoinositol phosphate

PKC Protein kinase C

PLC Phospholipase Cβ2

PMN Polymorphonuclear or polymononuclear

pNA Petidyl-4-nitroanilide

PPE Porcine pancreatic elastase or elastase (Pancreatic from porcine pancreas)

PPG Polypropylene glycol

Pro Proline (P)

Pro-MMPs Proenzyme-MMPs or zymogen of MMPs

PSGL-1 P-selectin glycoprotein ligand -1

PVA Poly(vinyl alcohol)

R1, R2, R3 Amino acid side chains

R2–NH Deprotonated amino acid

R2–NH2 Amine product

RAA Fmoc-X-Ala-Ala-PEGA, where X: Arg(+)

RBCs Red blood cells

Abbreviations

22

RFF Fmoc-X-Phe-Phe-PEGA, where X: Arg(+)

RFU Relative fluorescence units

RGD Arginine – Glycine – Aspartic acid

ROS Reactive oxygen species

RNS Reactive nitrogen species

rpHPLC Reverse phase high performance liquid chromatography

RV Release vesicles

RXKR Furin recognition motif

S&N Smith and Nephew

S. aureus Staphylococcus aureus

So Ground state

S1, S2 High energy state, excitated state

SD Significantly different, p < 0.05

SDS Sodium dodecyl sulphate

SDS-Page Sodium dodecyl sulphate – polyacrylamide gel electrophoresis

SE Standard error

Ser Serine (S)

SEM Standard error of the mean

SFM Serum free medium

sIPNs Semi-interpenetrating networks

SLPI secretory leukocyte elastase inhibitor

SP Signal peptide

SPPS Solid phase peptide synthesis

SPSS Statistical package for social science

S-SEBS Sulfonated styrene – ethylene – butylenes – styrene

TFA Trifluoroacetic acid

TGF-α Transforming growth factor-α

TGF-β Transforming growth factor-β

TI Tetrahedral intermediate

Thr Threonine (T)

TIMPs Tissue inhibitors matrix proteinases or tissue metalloproteases inhibitors (e.g. TIMP-1, -2, -3, -4)

TNF Tumour necrosis factor

TNF-α Tumour necrosis factor-alpha

TNFr Tumour necrosis factor receptor

TM Trademark

tPA Tissue-type plasminogen activator

Abbreviations

23

TPM Two-photon microscopy

Trp Tryptophan (W)

Tyr Tyrosine (Y)

UKSB UK Society for Biomaterials

UMIST University of Manchester Institute of Science & Technology

uPA Urinokinase-type plasminogen activator

US United States

USA United States of America

UK United Kingdom

UV Ultraviolet

UV/VIS Ultraviolet/ visible spectroscopy

VAC® Vacuum-Assisted Closure

Val Valine (V)

VEGF Vascular endothelial growth factor

VLUs Venous leg ulcers

Vmax Maximum velocity

WBCs White blood cells

Zn Zinc

,

24

CCOONNFFEERREENNCCEE PPRREESSEENNTTAATTIIOONNSS

Surface Science of Biologically Important Interfaces meeting (2007). Removal of Chronic Wound Proteases Using Enzyme-Responsive Hydrogel Dressing (poster presentation), University of Manchester, Manchester, UK. Materials Research Society (Spring Meeting, 2007). PEG based enzyme-responsive hydrogel particles for treatment of chronic wounds (oral presentation), San Francisco, USA. UKSB (28th - 29th July 2007). PEG based enzyme-responsive hydrogel particles for treatment of chronic wounds (poster presentation), University of King’s College, London. UK. FEBS advanced course: Microspectroscopy - Imaging Biochemical Dynamics in Living cells (2006). PEG based polymer hydrogel beads for removing proteases in chronic wounds (oral presentation), University of Waginengen (MSCWU), Waginengen, Netherlands, EU. UK Polymer Forum Conference (11th - 12th September 2006). PEG based polymer hydrogel beads for removing proteases in chronic wounds (poster presentation), University of Manchester, Manchester. UK. UKSB (28th - 29th June 2006). Responsive PEGA Beads for Removing Proteases in Chronic wounds (poster presentation), University of Manchester, Manchester. UK. Royal Society of Chemistry (RSC): Biomaterials Chemistry (18th January 2006). Responsive Hydrogel Beads for Selective Removal of Proteases in Chronic Wounds (poster presentation). University of Sheffield, Sheffield. UK. 19th European Conference on Biomaterials: ESB (11th-15th September 2005). Designing PEGA beads as a dressing to treat non-healing wounds (poster presentation), Sorrento. Italy. EU. UK Society for Biomaterials (UKSB, 21st -22nd June 2005). Enzyme Responsive PEGA beads for the Management of Wound Care (poster presentation), University of Nottingham, Nottingham. UK.

CHAPTER 1: Introduction

25

CCHHAAPPTTEERR 11

II nnttrroodduuccttiioonn


26

Wound healing is a complex biological process consisting of four overlapping phases by

which the skin or mucous membrane heals itself after an injury (Baranoski and Ayello

2004; Jones et al. 2004; Tortora and Grabowski 1993; Wysocki 1996) whether its an

internal or external break in the body tissue. Several proteins, cytokines, growths factors,

different types of cells, components of extracellular matrix (ECM) and proteolytic

enzymes are all involved in conjunction with the immune response to ensure proper

healing of the wound and that each phase of wound healing process is timely controlled

to completion to restore the integrity of the injured tissue (Westerhof and Vanscheidt

1994; Wysocki 1996; Siedler and Schuller-Petrovic 2002).

Non-healing chronic wounds take ‘time to heal’ as they do not undergo the timely

controlled wound healing process instead eventually become deadlocked during the

inflammation/ proliferation phase (Ayello et al. 2004; Morrison et al. 2004) and are

unable to progress to the final remodelling phase of the wound healing process; and

subsequently the wound struggles to close. This causes substantial trauma, which

decreases both the mobility and quality of life of millions of people which consequently

increases the risk of amputation and mortality of chronic wound sufferers each year. The

patients most at risk are the elderly and those who suffer from clinical diseases/

conditions (e.g. diabetes, heart disease, lung disease, circulation disorders, liver disease,

cancer, arthritis, autoimmune disorders, obesity) and skin ulcers (foot, pressure,

vascular/leg, arterial). Skin ulcers are the largest and more frequent types of chronic

wounds (Eaglstein and Falanga 1997; Stadelmann et al. 1998).

As the population ages, the number of patients suffering from chronic wounds is

reported to increase each year: 2 million for pressure ulcers and between 600,000 – 2.5

million for leg and foot chronic ulcers (Stadelmann et al. 1998). This causes a significant

clinical burden on both the healthcare/ resources required and the financial cost of

treating chronic wounds (Wysocki 1996; The Global Market for Advanced Wound Care

Products (GMAWCP) 2008). In UK alone the wound care treatment for chronic wounds

was estimated to cost the NHS £2.3 – 3.1 billion in 2005 – 2006 (Posnett and Franks

2008), in the US $10 billion dollars was estimated in 2007 (for each unit cost of wound

dressing) and the largest wound care market in 2007 was Europe with a total estimated

cost of healing the wound as US $2.2 billion (GMAWCP 2008). Wound care has been


27

estimated to rise to US $12.5 billion in 2012 in the US (GMAWCP 2008) and globally

between $13 – > $15 billion annually (Fonder et al. 2008; Kannan 2008). The length of

hospitalisation faced by chronic suffers is another medical problem that increases the

cost of treatment as wound care physicians/ clinicians are unable to detect the correct

prognosis to effectively treat patients as the pathophysiology of chronic wounds is not

fully understood as of yet.

However, in recent years, at the molecular level substantial research has reported that

various biochemical factors of elevated-expression of proteolytic activity and the

decrease activity of their inhibitors are the underlying problem of deadlocking the healing

of chronic wounds. The imbalance of various proteases: MMPs, plasmin, thrombin, and

elastase (Westerhof and Vanscheidt 1994; Wysocki 1996; Tarnuzzer and Schultz 1996;

Stadelmann et al. 1998; Trengove et al. 1999; Yager et al. 1996, 1997; Yager and

Nwomeh 1999; Cullen et al. 2002b; Greener et al. 2005; Xue et al. 2006; Schultz et al.

2005a) in chronic wound fluids (CWFs) are able to degrade various cellular factors:

growth factors, cytokines, their receptors; and the newly formed ECM and its proteins

(e.g. elastin, fibronection, collagen). This accordingly perturbs the normal tissue repair

and the wound becomes stuck during the healing process.

Excess elastase has been reported as the main culprit for perpetuating the over-

expression of MMPs in chronic wounds during inflammation due to the imbalance of

elastase and elastase inhibitors (Fleck and Chakravarthy 2007). The prime function of

elastase is to degrade elastin, however it also play a role in activating latent MMPs to

active MMPs (Zhu et al. 2001; Fleck and Chakravarthy 2007) which in turn initiates the

cascade of active MMPs to activate other latent MMPs (Zhu et al. 2001). This ‘vicious

cycle’ of elastase is further propagated in chronic wounds by degrading the tissue

metalloproteases inhibitors (TIMPs) which inhibit and control MMPs activity. This

imbalance between MMPs and TIMPs causes extensive turnover and degradation of the

newly formed collagen of the ECM (Zhu et al. 2001; Fleck and Chakravarthy 2007).

Numerous wound dressings are available (Hess 2002); despite widespread scientific

perception about wound dressings, globally most of the available traditional and modern

dressings are passive meaning they are non-responsive. Currently, the research and

innovation within wound companies and university discoveries of wound care products


28

is shifting from the conventional non-responsive wound dressings to generating

advanced wound dressings of biocompatible polymers. In doing so, such advanced

wound dressings contain a responsive element that will tackle and control the underlying

biochemical mechanisms, pathological and cellular turnover caused by high proteolytic

activity in order to facilitate wound healing of a deadlocked wound by restoring the

imbalance of proteases to normal levels.

Despite significant advances in developing new advanced wound dressings, traditional

dressings are still used. This is because the complexity and lack of knowledge by wound

care clinicians/ physicians of the new advance wound dressings hinders wound care

clinicians/ physicians to make use of them (Nobel 2006) since the pathophysiology of

chronic wounds is still poorly understood. In recent years, various studies have

demonstrated inhibiting the expression of proteolytic activity by sophisticated wound

dressings such as PromogranTM which has been shown to be more effective (Johnsons &

Johnsons, Cullen et al. 2002a; Nobel 2006; Smeets et al. 2008), but there are built-in risks

associated with this dressing as it is made of bovine which is of animal origin (Eming et

al. 2008). Therefore, there is a necessity and a growing interest to increase the availability

of responsive advanced wound dressings of biocompatible synthetic origin under moist

healing conditions because this is accepted as the most effective way of healing wounds

(Winter 1962). Much attention has been contributed to developing responsive hydrogel

dressings as these materials play an important role in protecting wounds from bacterial

infiltration, dehydration, controls trauma and promotes moist wound healing, but most

importantly is biologically biocompatible to the body.

1.1 Objective

The main objective of this thesis is to develop a responsive hydrogel polymer based on

poly(ethylene) glycol acrylamide (PEGA) particles (figure 1) that will selectively mop-up

and entrap excess elastase into the hydrogel material in order to improve the

management of chronic wounds (figure 2).


29

NH

O n

NHO

NHO

*

*

NH2

O

n

n

CH3

CH3

O

O C

H2

CH

2

O

xl

n

NH2

CH2

CH2

O

CH3

xl

Figure 1. The molecular structure of PEGA particles. PEGA consists of a polyacrylamide backbone (n, red) with PEG crosslinks (black and blue). The free amine groups (green) are used to couple enzyme cleavable peptides (ECPs) using solid phase peptide synthesis (SPPS). Depending on the type of PEGA, the molecular weight of the bio-inert PEG chains (xl, blue) within the matrix of the hydrogel vary: PEGA1900 has a longer PEG chain as opposed to PEGA800; consequently the loading (mmol/g) of PEGA1900 is less compared to PEGA800.

Figure 2. A schematic diagram exhibit a deep chronic wound being healed after treatment with PEGA particles (light solid circles) that have been are designed to selectivity mop-up excess proteases presence (dark solid circles with pink colour inside) in chronic wounds.

1.2 Aims of the project

The aims of the project are split into three sections. Initially the research involved

functionalising the matrix of PEGA particles with Fmoc-peptide substrates which are

referred to as enzyme cleavable peptides (ECP) so that the hydrogel polymer mimics

elastase substrates involved in normal wound healing. Elastase then recognises and

cleaves the peptide sequences of the ECP; and the charges of the ECP control the

accessibility of elastase into the interior of PEGA particles. The penetration of elastase

into interior of PEGA particles is determined by swelling of the PEGA particles

depending on pH and ionic strength. Once inside the interior of PEGA particles elastase

eventually becomes entrapped. This is achieved by the pores of PEGA particles which

Chronic wound

Remove particles with

entrapped protease

Particles added to chronic wound

↑ Healing of chronic wound


30

collapse upon selective removal of charged groups within the matrix of the PEGA

particles (figure 3) thus removing the elastase from the sample fluid. Finally, elastase-

type activity expressed by fibroblast cells was shown to decrease in the presence of

PEGA particles. To address the aims, the scope of the experimental study is split into

three individual chapters (see section 1.3).

Figure 3. Swelling to collapse of PEGA particles. PEGA particles are modified by the incorporation of charges (for example negative) and this causes a proteases of opposite charge (in this case, positive) to be attracted and then diffuse into the PEGA particle. The protease selectively recognises and cleaves the ECPs coupled to PEGA particles. Removing the charge of the ECPs causes the pores of PEGA particles to collapse thus entrapping the proteases within the particle.

1.3 Scope of Thesis

Chapter 2 reviews the literature; firstly the chapter reviews the physiological anatomy

(structure and function) of the skin and the fundamental biology for the process of

wound healing in the context of acute healing in order to understand chronic wound

healing and the two categories are differentiated enabling the reader to understand the

pathophysiology of chronic wounds compared to acute wounds. This chapter gives an

account of the role of proteases in wound healing which includes showing the broad

selectively of wound proteases and most importantly determining the charge of each

protease depending on its pI value. After discussing chronic wounds the chapter focuses

on giving an account of reports of high levels of proteases (e.g. elastase) as the underlying

cause of chronic wounds. Finally, the ending of this chapter is devoted to the

management of wound healing and the discussion in this thesis focuses on hydrogel

dressings (both non-responsive and responsive) that are available/ researched for

chronic wounds as well as the recent developments of protease-modulating hydrogels

design to tackle elevated levels of proteases observed in chronic wounds.

SPPS

Protease (+ve)

PEGA particles swells with the incorporation of

negative charges

PEGA particles collapses upon removal of charge, protease is trapped (pink)

Unmodified PEGA particles


31

Following chapter 2, the experimental chapters are presented as three individual chapters.

To lead the reader in a logical way each chapter has their own introduction, methods and

materials, results and discussion as well as conclusions. The reader should note that

there is overlap of certain methods in all three experimental chapters and these have

been cross-referenced to the corresponding chapter to enable the reader to easily locate

the method in question.

Chapter 3 outlines the use of stepwise solid phase peptide synthesis (SPPS) to

functionalise PEGA particles with enzyme cleavable peptides responsive to elastase using

Fmoc-chemistry (HOBt/ DIC). The functionality of modified PEGA particles is further

shown to be controlled by monitoring the swelling of the PEGA particles as a function

of pH and ionic strength. Next, chapter 4 outlines the accessibility and entrapment of

elastase into PEGA particles by exploiting the use of various fluorescence techniques

using both indirect/ direct approaches. Finally, chapter 5 takes advantage of using

human dermal fibroblasts to explore the expression of elastase-like by these cells and

demonstrates that a decrease in elastase-like activity is observed in the presence of the

PEGA responsive hydrogel particles. Subsequently conclusions and further studies are

presented in chapter 6.

CHAPTER 2: Literature Review – Wound Healing & Management of Chronic Wounds

32

CCHHAAPPTTEERR 22

LLii tteerraattuurree RReevviieeww -- WWoouunndd HHeeaall iinngg && MMaannaaggeemmeenntt ooff CChhrroonniicc WWoouunnddss


33

2.1 ANATOMY & PHYSIOLOGY OF THE SKIN 2.1.1 Anatomy of the skin

Structurally, the human skin consists of three layers: the outer, thinner layer is composed

of epithelium called the EPIDERMIS (a.k.a cuticle; Cohen et al. 1992). The epidermis is

attached to the inner, thicker layer consisting of connective tissue known as the

DERMIS (a.k.a corium; Cohen et al. 1992). Beneath the dermis is the

SUBCUTANEOUS layer (a.k.a. superficial fascia or hypodermis or subcutis), which

consists of aveolar and adipose tissues (Baranoski and Ayello 2004; Tortora and

Grabowski 1993). Fibres from the dermis extend down into the subcutaneous layer and

anchor the skin to it. The subcutaneous layer, in turn, attaches to underlying tissues and

organs, and therefore separates the skin from the muscle of the body wall beneath. The

epidermis and dermis are the two major layers of the skin (Baranoski and Ayello 2004;

Cohen et al. 1992; Tortora and Grabowski 1993), which are now described in detail.

2.1.1.1 Epidermis

The epidermis is the most superficial layer of the skin, composed of stratified squamous

epithelium (layered cells) and contains four principle types of cells of the epidermal cells:

(1) Keratinocytes: these produce the protein keratin, (2) Melanocytes: these produce the

pigment melanin, (3) Langerhans cells: these arise from the bone marrow and interact

with helper T cells in immune responses, and (4) Merkel cells: these make contact with

the tactile disc (and ending of a sensory neuron) (Tortora and Grabowski 1993).

There are two distinct regions of the epidermis (i) Upper area – where cells convert from

living to dead; and (ii) Cell renewal area – allows living cells to be renewed via mitosis.

The two regions are composed of five layers, from the deepest to the most superficial,

are: stratum basale, stratum spinosum, stratum granulosum, stratum lucidum, stratum

corneum (Baranoski and Ayello 2004; Tortora and Grabowski 1993); further details of

these layers are expressed in table 1.


34

Table 1. Layers of the epidermis

Layers of the epidermis

Description Function

Stratum germination (basale layer)

• Actively living layer of epidermis

• Deepest layer of the epidermis

• Its lower surface is attached to the dermis (from which it receives nutrient fluid from the blood vessels)

• Contains stem cells, which multiply, and produce keratinocytes – which push up towards the surface and become part of the more superficial layers

• Melanocytes (melanin-forming cells) are formed within this layer

• Contains tactile (Merkel) discs

• Cells are in active growth

• Stem cells are capable of continuing cell division (mitosis) – development of new cells occurs and gradual displacement of older cells towards surface of skin

• Melanin: protects skin against damaging ultraviolet (UV) light & changes in skin colour when exposed to the sun

• Tactile disc function in the sensation of touch

Stratum spinosum

(prinkle cell layer)

• Actively living layer of the epidermis

• Contains 8 – 10 rows of polyhedral cells that fit closely together

• Composed of newly formed epidermal cells, called keratinocytes. These cells are joined together by prickly desmosomes (outgrowths), hence the name prickle layer

• Keratinisation is the change of living cells containing a nucleus into layers of flat cells composed of hard durable protein keratin

• Long projections of melanocytes extend among the keratinocytes, which take in melanin by phagocytosis of these melanocyte processes

• Connecting of the keratinocytes allows each cell to receive nourishment from the tissue fluid or protoplasm

• Cells start to die via the Keratinisation process – keratinocytes are in early stages of producing the a tough fibrous protein called keratin

Stratum granulosum (granular layer)

• Non-living layer of epidermis

• Consist of 3 – 5 rows of flattened cells

• Cells become flattened and nucleus disintegrates, there is a loss of fluids which contributes to the development of keratohyalin (a precursor of keratin)

• Breakdown of nucleus – the cells are no longer able to carry out vital metabolic reactions

• Final stages of keratinisation take place in this layer

• Keratohyalin – transforms the cells into keratin (a protein found in outer layer of epidermis)

• Cells die due to the breakdown of nucleus and cell contents

Stratum lucidum

(transparent layer)

• Non-living layer of epidermis

• Lies between the outer Stratum corneum and the inner Stratum granulosum

• Consist of 3 – 5 layers of small, tightly packed (flat) transparent dead cells containing droplets of keratohyalin and no nucleus

• Appears in the thickest areas of skin, the soles of feet and palm of the hands

• These cells contain an epidermal fatty substance resembling bees’ wax

• The droplets keratohyalin (dead cells) is eventually transformed into keratin

• Epidermal fatty substance

– functions as a barrier zone controlling the transmission of water through the skin, and

– helps prevent the skin from cracking and becoming open to bacterial infection

Stratum corneum

(horny layer)

• outer layer – non-living layer of epidermis

• cells have no nucleus

• Consist of 25 – 30 layers of mature squames – keratinised epithelial cells tightly packed together (flat), dead cells completely filled with keratin

• Each squame is roughly disc shaped – has a tough protein wall and packed with keratin

• All molecules of keratin are aligned parallel to the surface of the skin and parallel to each other

• This layer protects the living cells within - serves as an effective barrier against UV light and heat waves, bacteria, and many chemicals

• The generations of squames - they gradually migrate towards the outer surface

• Eventually, cells of superficial layers are continuously shed and replaced by cells from deeper strata

(Baranoski and Ayello 2004; Hess 2002; Tortora and Grabowski 1993)


35

2.1.1.2 Dermis

The second principal part of the skin, the dermis lies beneath the epidermis and is the

tough elastic layer (composed of connective tissue containing collagen and elastic fibres)

of the skin. The dermis has higher water content than any other region of the skin and

provides nourishment to the epidermis. It is composed of two layers; the outermost

layer just beneath the epidermis is the papillary layer (Baranoski and Ayello 2004; Cohen

et al. 1992; Tortora and Grabowski 1993). Beneath it and forming the bulk of the dermis

is the reticular layer (Baranoski and Ayello 2004; Cohen et al. 1992; Tortora and

Grabowski 1993), which has most of the protein fibres (refer to table 2).

Table 2. Layers of the dermis

Dermis layers Description Function

Papillary 1ayer

• situated at the interface of the dermis with the epidermis

• undulating (wavy) tissue

• consisting of areolar connective tissue containing fine elastic fibres

• upwards projections are called dermal papillae, they contain blood & lymph capillaries and nerve endings

• highly active & important area of skin

• increases surface area of reproductive cells – dermal papillae

• provides living layers of epidermis with vessels which supply nourishment and remove cellular waste

• nerve endings perform skin’s sensory function

Reticular layer

• situated beneath papillary layer

• dense and fibrous (tough and elastic collagen fibres)

• protein fibres are made from fibroblast cells contained in a ‘ground substance’

• contains main components of the skin: spaces between the fibres are occupied by adipose tissue, hair follicles, nerves, oil glands, and ducts of sweat glands

• attached to underlying organs i.e. bone and muscle

• this layer varies in thickness depending on differences in thickness of skin

• protects and repairs injured tissue

• fibres allow skin to bend and fold over underlying muscle activity:

– Collagen: gives skin its strength and resilience

– Elastin: allows skin to stretch easily and quickly regain its shape, and general tone of skin (smoothness to epidermis)

– Reticulum: keeps all the other structures in place


2.1.2 Physiological function of the skin

The skin is the largest organ of the human body in surface area and weight. It consists of

different tissues that are joined to perform specific activities. It provides a tough, flexible

covering and has many essential functions (as depicted in the table 3).


36

Table 3. Summary – The physiological function of the skin

Function Description for role of the skin Temperature regulation

Body temperature is controlled by heat loss through the skin and by sweating:

• High temperature - evaporation of sweat from skin surface lowers elevated body temperature to normal.

• Low temperature - production of sweat is decreased, therefore conserves heat

Protection Skin covers the body providing a physical barrier that protects underlying tissues from:

• Physical abrasion

• Bacterial invasion

• Dehydration

• UV radiation

Sensation Skin is very sensitive:

• It contains abundant nerve ending and receptors, which detect stimuli - enabling the feelings of touch, pressure, pain and temperature

Excretion

Skin aids in the removal of waste products from the body:

• Sweat also allows the excretion of small amounts of salts and several organic compounds

Immunity Some cells of epidermis are essential components of the immune system

• Skin provides a warning system (1st line defence mechanism) against outside invasion i.e. foreign invaders

• Visual indication (redness/ irritation) shows the skin is intolerant to either an external or internal stimuli

Blood reservoir

Dermis of skin contains extensive networks of blood vessels

• Skin blood flow may increase – aids to dissipate heat from the body, or

• Skin blood vessels may constrict (narrow) ∴ more blood is able to circulate to contracting muscles.

Nutrition Skin enables fat to be stored:

• An energy reserve Synthesis of Vitamin D:

• Achieved by the activation of a precursor molecule in the skin by UV rays of sunlight

• Enzymes then modify the molecule, finally producing calcitriol (active form of vitamin D)

• Calcitriol – enables the homeostasis of body fluids by the absorption of calcium in foods

• Vitamin D is an hormone as its produced in one location of the body, then transported by

the blood to function in another location ∴ the skin is considered to behave as an endocrine organ

Moisture control

Skin controls the level and movements of moisture from within the deeper layers of the skin


In order for the skin to function effectively, the skin must be cared for both internally

and externally as it plays an important role in maintaining homeostasis of the body. One

way in which this is achieved, is by wound healing (Tortora and Grabowski 1993).


37

2.2 WOUND HEALING

Various stimuli can damage the skin, and the body has an extensive defence system,

where it undergoes several mechanisms to maintain normal structure and function

(Tortora and Grabowski 1993). Wound healing can be considered as the process by

which a wound regenerates and repairs itself.

2.2.1 Acute and chronic wound healing

Wound healing can be classified into two forms: acute (normal) and chronic (abnormal)

wound healing (Boyce 1996; Jones et al. 2004). Acute wounds can be considered as

normal wounds, which are the end result of traumatic injury or surgery (Boyce 1996;

Hess 2002; Jones et al. 2004) and can be defined as “a disruption in the integrity of the

skin and underlying tissues that progress through the healing process in a timely and

uncomplicated manner” (Bates-Jensen and Wethe as cited in Jones et al. 2004) via

primary intention (Hess 2002; Jones et al. 2004). On the other hand, chronic wounds

heal by secondary intention (Hess 2002; Jones et al. 2004) and can be defined as “any

interruption in the continuity of the body’s tissue that requires a prolonged time to heal,

does not heal, or recurs” (Wysocki 1996) and “fails to progress through a normal,

orderly, and timely sequence of repair or wounds that pass through the repair process

without restoring anatomic and functional results” (Lazarus as cited in Jones et al. 2004),

and therefore resulting in the formation of a chronic wound. A third way of repairing

the skin is via tertiary intention (delayed primary) where healing of skin involves closure

by using sutures, staples or adhesive skin closures.

Wounds of the skin can be classified into different types according to the layers of the

skin involved: (1) Superficial wounds – the epidermis layer is only involved; (2) Partial

thickness wounds – involve the dermis; and (3) Full thickness skin wounds – cut deep

into the subcutaneous layer (Collins et al. 1992). It can be said that chronic wounds are

usually the result of partial or full thickness wounds. However, superficial wounds can in

turn result into a chronic wound only when the wound healing is disrupted.


38

2.2.2 Phases of wound healing and the immune response

The fundamental basis of acute wound healing needs to be addressed to understand

chronic wounds (Wysocki 1996). Both wound-healing processes undergo similar

sequences of events, which occur simultaneously to repair and restore tissue integrity of

the injury (Jones et al. 2004; Wysocki 1996). Wound healing is a complex cascade

consisting of four timely controlled and overlapping processes of biochemical and

cellular events that are divided into four stages as follows: (1) Hemostasis (coagulation);

(2) Inflammation; (3) Proliferation; and (4) Remodelling (Jones et al. 2004) as depicted in

figure 4. The progression of the healing process is essentially controlled by the

signalling events from growth factors and cytokines.

Figure 4. Events of acute and chronic wound healing. (Figure modified from Baranoski and Ayello 2004; Boyce 1996).

2.2.2.1 Hemostasis

Hemostasis can be referred to as acute inflammation (Cohen et al. 1992) or the initiation

phase (Collins et al. 1992) that begins immediately after an injury. An injury causes

PROLIFERATION Anabolic reactions

Mitosis → cell proliferation Various proteases i.e. MMPs

Angiogenesis Epithelisation

HEMOSTASIS Platelet Activation

→ Blood coagulation (complement cascade)

INFLAMMATION Catabolic reactions

Recruitment of leukocytes (neutrophils, macrophages)

→ phagocytose bacteria and breakdown damaged tissue

REMODELLING Synthesis & ECM remodelling of collagen

Activation of collagenases → degradation of collagen

fibrinolysis

Vasoconstriction

NORMAL SKIN

INJURY

HEALED SKIN

Delayed healing

↓ CHRONIC WOUND

Impeded phase


39

excessive bleeding, which is stopped by the formation of a clot (thrombus) over the

wound; which is controlled by the process of blood clotting (coagulation) - a complex

cascade of enzymatic reactions that occur via two processes: (1) the classical (intrinsic or

contact activation) pathway and (2) the alternative (extrinsic or tissue factor pathway)

pathway. Both pathways eventually combine to activate the complement pathway, and

this releases complement split products that are essential mediators required for

inflammation.

In response to the depositing of the epidermal layer, the blood coagulation cascade is

activated and this process is mediated by the release of platelets (thrombocytes), clotting

factors (a variety of enzymes), erythrocytes, and leukocytes (Wysocki 1996; Boyce 1996;

Lobmann et al. 2005). Ruptured blood vessels expose the ruptured collagen to matrix

proteins, which activate platelet aggregation. The activated platelets release growth

factors, such as the platelet-derived growth factor (PDGF) and the basic fibroblast

growth factor (bFGF) (Silver and Christiansen 1999; Lobmann et al. 2005) that initiate

the initial phase of hemostasis by causing vascular constriction of ruptured blood vessels

to stop bleeding.

Figure 5. Formation of thrombin and fibrin.

This is then followed by vasodilation causing the plasma to exit from the capillaries and

into the wound. When the platelets interact with the injured tissue, this results in the

release of thrombin (a serine protease), which in turn catalyses the conversion of the

soluble plasma protein, fibrinogen into insoluble fibrin strands and fibrinopeptides

(Cohen et al. 1992; Kuby 1997) as summarised by figure 5. The blood plasma becomes

sticky and the fibrin strands crisscross one another like a mesh to form a hemostatic

blood clot (a.k.a the fibrin clot, fibrin plug, fibrin blood clot or blood clot) over the

Prothrombin (inactive enzyme)

THROMBIN (active enzyme)

FX

FXa (thrombokinase)

CLOT

Fibrinogen (soluble)

Fibrinopeptides + FIBRIN (insoluble)


40

wound that traps red blood cells and activated platelets. The activated platelets release

cytokines and growth factors that provide signals to further propagate the clot (Cohen et

al. 1992) to stick to blood cells and attach to fibronectin and vitronectin (plasma

proteins).

In subsequent phases of wound healing, the fibrin clot serves as a provisional matrix that

allows inflammatory cells to attach, proliferate and migrate into the wound site (Kerstein

1997; Martin 1997). When the blood vessels have sealed, the temporary fibrin clot is

removed from the wounded area by the activation of fibrinolytic pathway (Kuby 1997;

Silver and Christiansen 1999) a.k.a the plasminogen or plasminogen/ plasmin system; an

enzymatic process summarised by figure 6.

Figure 6. Fibrinolytic pathway. Fibrinolysis in the blood is essentially regulated by tPA, whereas in tissues its regulated by uPA. Abbreviations: PAs (plasminogen activators): tPA (tissue-type plasminogen activator) and uPA (urolinase-type plasminogen activator); ↑ (increase); and WBCs (white blood cells). Figure tailored from Kudy 1997; Silver and Christiansen 1999; Liekens et al. 2001.

The fibrin clot is lysed by plasmin (a serine protease) into fibrin degradation products i.e.

chemoattractants and cellular building blocks (Cohen et al. 1992) that are reabsorbed by

the body and utilised during subsequent phases of the wound healing process. In

addition, when the activated platelets degranulate, their α-granules release various

cytokines and growth factors, such as: PDGF, platelet-derived angiogenic factor (PDAF),

transforming growth factor-β (TGF-β), and epidermal growth factor (EGF) (Silver and

Christiansen 1999; Lobmann et al. 2005), transforming growth factor-α (TGF-α) and

vascular endothelial growth factor (VEGF) (Morison et al. 2004) that are largely (1)

chemoattractants for inflammatory cells (neutrophils, monocytes and macrophages) and

(2) mitogens for the non-inflammatory cells (fibroblasts and endothelial cells) that are

Plasminogen (inactive enzyme)

PLASMIN (active enzyme)

Activation of Complement

System

FIBRIN CLOT (insoluble)

Products of fibrin degradation

↑ Vascular permeability

↑ WBCs (neutrophil)

↑ chemotaxis

Endothelial cell factors (PAs: uPA, tPA) Blood clotting factors


41

involved in subsequent phases of wound healing. The initial release of the growth

factors from the platelets is essentially crucial for successful progression of the

subsequent phases of wound healing (Singer and Clark 1999; Lobmann et al. 2005).

Various vasodilatory substances (such as serotonin, histamines) are produced by

endothelial cells, which also aid in the initiation of the inflammation stage of wound

healing process (Boyce 1996).

2.2.2.2 Inflammation

The inflammation phase is also known as the defence or reaction phase (Hess 2002).

This stage is usually personified by redness, swelling, heat and pain (Hess 2002; Lobmann

et al. 2005) that typically last for 1 – 7 days (Lobmann et al. 2005). The inflammation

phase is exemplified by vasodilation and increased permeability for the migration and

activation of white blood cells (WBCs, mast cells, neurophils, granulocytes, monocytes,

lymphocytes) from the blood into the tissue (as summarised in figure 7). This is

mediated by chemotaxis and various growth factors (Boyce 1996) which is vital for

immune surveillance and inflammation.

The inflammation stage begins 6 hours after injury (Lobmann et al. 2005) and mainly

involves catabolic reactions. The first leukocytes to migrate into the provisional matrix

of the wound are neutrophils. During this diapesis of leukocytes, the leukocytes adhere

to endothelial cell adhesion molecules and then migrate across the vascular endothelium.

Endothelial cell adhesion molecules and their counter-receptors on leukocytes (e.g.

neutrophils, monoctyes etc.) generate intracellular signals. These signals are controlled

by the release of growth factors and chemoattractants from leukocytes and red blood

cells to activate the cell-cell adhesion mechanism.

The selectin family consist of three different single chain transmembrane receptors: E-,

P-, and L-selectin. These receptors initiate leukocyte: capturing, rolling and tethering (i.e.

weak adhesion) during the inflammation process. However, the integrins are involved in

firmer adhesion before leukocyte emigration from blood vessels into the target tissues.

Integrins are heterodimetric transmembrane glycoproteins, composed of non-covalently

associated α and β subunit, each a type-1 membrane protein.


42

Leukocytes GPCR (G-protein coupled receptors)

RBC (red blood cells)

Chemokine Proteolycan Endothelial cells

Figure 7. General mechanism of leukocyte adhesion to the endothelial cells, mediated by the cell-cell adhesion mechanism of selectins/ integrin. The selectins enable weak adhesion whereas the integrins enable strong adhesion. All leukocytes undergo this general mechanism of cell-cell adhesion once they migrate into provisional matrix during wound healing. (Figure modified from Silver and Christiansen 1999).

Initially, the P- and E-selectins bind to their counter-receptors (P-selectin glycoprotein

ligand-1 (PSGL-1), E-selectin ligand-1 (ESL-1) respectively) on the leukocyte membrane,

which weakly tethers leukocyte to the endothelium and initiates rolling along the

endothelium cell. As the leukocyte cell rolls, the epithelium cells produce and secrete

chemotactic cytokines, such as interleukin-2 (IL-2), which activates L-selectin to shed

from the leukocytes soon after its activation (Springer 1994) to attract the leukocyte to

endothelium. Then, the chemokines bind to their counter-receptors, G-protein coupled

receptors (GPCRs) at the leukocyte surface, and induces an ‘inside-out’ signalling cascade

which induces G-protein/tyrosine kinase activity resulting in phospholipase Cβ2 (PLC)-

mediated phosphoinositol phosphate (PIP) hydrolysis. The subsequent release of

diacylglycerol (DAG) results in protein kinase C (PKC) activation and IP3-mediated

release of [Ca2+]. During cell-cell adhesion, these signals up-regulate cell surface

Ca2+

Selectins Integrins & IgSF Rolling Sticking Extravasation Chemotaxis

Activation by chemokine Activation of

endothelial cells Activation of leukocytes

LUMEN

TSSUE SPACE Site of injury of

wound

Chemotaxis


43

expression of the α/β2-integrin on the leukocyte to gain affinity for the immunoglobulin

superfamily (IgSF) cell adhesion molecules (CAMs) on the endothelial cells and therefore

the activation of the α/β2-integrin causes its conformational change from an inactive to

an active form. This is further enhanced by binding of divalent cations to α-subunit of

integrin.

The α/β2-integrins then bind to the Leu-Val-Asp-Pro (LVDP) domains of fibronectin

on the endothelial cells of the endothelium – inducing the outside-in signalling

transduction pathways. Such signals stabilise adhesion causing the leukocyte to firmly

adheres to the endothelial cell (Springer 1994), and is able to transmigrate between the

endothelial cells, crossing the blood vessel wall into the damage/infected tissue. In doing

so, the leukocytes mop up debris, damaged tissue, foreign substances and attack any

pathogens to control the infection.

Neutrophils have the ability to phagocytose bacteria, ECM and degrade degenerating

connective tissue by secreting destructive proteases (Morison et al. 2004; Theilgaard-

Moench et al. 2004) such as elastase. At 24 hours, the levels of neutrophils reach their

peak (Cohen et al. 1992; Bowler 2002) and begin to decrease as the monocytes migrate

into the wound (Cohen et al. 1992; Lobmann et al. 2005). These monocytes also

undergo phagocytosis and then differentiate into macrophages (Cohen et al. 1992;

Lobmann et al. 2005) between 48 – 72 hours and persist for a few days. The

macrophages suppress bacterial growth, debride the wounded area of cellular debris,

necrotic tissue (Kuby 1997; Hess 2002) and the damaged matrix is broken down by

proteolytic enzymes (i.e. collagenase and elastase) secreted by macrophages. Also, during

wound healing, the macrophages mediate the conversion of macromolecules to amino

acids and sugars (Hess 2002).

Macrophages further release growth factors (such as transforming growth factor beta,

TGF-β) and chemoattractants/ cytokines (such as interleukin-1, IL-1; tumour necrosis

factor, TNF) to recruit cells involved in the proliferation phase such as fibroblasts

(develop from mesenchymal cells) and endothelial cells to the site of injury around 3 – 4

day post-wounding just before the inflammatory phase ends (Cohen et al. 1992; Hess

2002). Eventually after a couple of days, fibroblasts become the predominant cell type.


44

This initiates the proliferation phase of wound healing process under the influence of

macrophages.

2.2.2.3 Proliferation

The proliferation stage lasts for approximately 24 days (Hess 2002). This phase involves

various overlapping processes: angiogenesis, granulation, epithelisation and wound

contraction in order for the epithelial cells beneath the scab to grow and replace the

provisional matrix with new collagen molecules that are matured in the remodelling

phase.

Angiogensis is also refer to as ‘neovascularisation’ and this process is initiated by the

secretion of angiogenic factors that are secreted by macrophages and these become

activated by low oxygen tension and the migration of endothelial cells. Angiogensis is

the process by which new blood vessels are formed from the endothelial cells (Cohen et

al. 1992; Tortora and Grabowski 1993; Morison et al. 2004). The process of angiogenesis

occurs simultaneously with the clonal expansion of fibroblast and the migration of

endothelial cells into the provisional matrix as fibroblasts and epithelial cells both require

oxygen, therefore the process of angiogenesis is important for the remaining phases of

wound healing.

During hypoxia (low oxygen levels) macrophages and platelets release angiogenic factors

which chemotactically recruit endothelial cells into the provisional matrix. The new

blood vessels are formed from the endothelial cells of undamaged blood vessels which

develop pseudopodia enabling them to push through the ECM and into the wound site.

The migration of endothelial cells into the provisional matrix is mediated by proteolytic

enzymes (i.e. collagenases and the plasminogen activators) to lyse the clot (for further

details on fibrinolysis see haemostasis phase and role of enzymes in wound healing) and

the ECM (Cohen et al. 1992; Hess 2002; Morison et al. 2004). The process continues

until the blood vessels are repaired and tissue oxygenation is restored (Morison et al.

2004) then the migration of endothelial cells is decreased.

Granulation tissue fills the open wound approximately 2 – 5 days post-wounding (during

the inflammatory phase) and the wound continues to fill and grow until the wound bed is

covered. Granulation tissue is characterised by the presence of macrophages, fibroblasts,


45

inflammatory cells, endothelial cells, new blood vessels, myofibroblasts, new provisional

ECM and fibronectin (Hess 2002; Morison et al. 2004).

Fibroblasts from the undamaged tissue migrate into the wound site. The growth factors

secreted by fibroblasts enable epithelial cells to migrate to the wound site. For about 2 –

4 weeks, the main function of fibroblasts is to rapidly synthesise collagen by randomly

depositing and laying down immature collagen type III fibres (Tortora and Grabowski

1993; Cohen et al. 1992; Silver and Christiansen 1999; Morison et al. 2004; Baranoski and

Ayello 2004). The growth factors and fibronectin also encourage proliferation,

migration across the wound bed, and production of ECM molecules by fibroblasts.

Eventually the production of immature collagen (type III) levels off as it equals its

destruction by collagenases and other factors (Baranoski and Ayello 2004; Morison et al.

2004). Granulation completes when the levels of fibroblast decrease and at the end of

this process fibroblast undergo apoptosis and the granulation tissue consist mainly of

type III collagen (Stadelmann et al. 1998).

Within a week, growth factors activate fibroblasts to differentiate into myofibroblasts

which cause the wound to contract (Baranoski and Ayello 2004). Under the influence of

fibronectin and growth factors myofibroblasts move along the fibronectin-fibrin clot in

the provisional ECM to migrate to the wound edges. At this stage the myofibroblasts

attach to each other as well as adhering to fibronectin and collagen molecules of the

ECM at the wound margins. In doing so, the actin in myofibroblasts contract

(Stadelmann et al. 1998) causing the edges of the wound to pull together sealing the open

wound from the external environment (Hess 2002). This triggers the epithelialisation

stage, which is the final step of the proliferation phase. The epithelialisation stage is

characterised by the presence of keratinocytes from the wound edges, hair follicles, sweat

glands, and sebaceous glands. The epithelial cells migrate across the granulation tissue to

form a thin barrier between the wound and the environment. Keratinocytes migrate

across the granulation tissue of the wound bed until epithelial cells from either side meet

in the middle, and the edges of the wound align to close the epidermal layer (Cohen et al.

1992; Bradley et al. 1999). The completion of the epithelisation results in the formation

of a scar (Hess 2002). By doing so the keratinocytes stop migrating and the

myofibroblasts stop contracting and this stops the proliferation phase and initiates the

remodelling phase.


46

2.2.2.4 Remodelling

The wound enters the remodelling phase (final stage of wound healing) by the migration

of keratinocytes. The remodelling stage is sometimes referred to as the maturation phase

(Hess 2002; Tortora and Grabowski 1993) and during this stage the randomly organised

collagen type III fibers are remodelled and matured into organised layers of collagen type

I and the number of fibroblasts present within the wound bed decrease; and finally the

blood vessels are restored to normal (Tortora and Grabowski 1993; Hess 2002). During

this phase, the thickness of the newly formed epidermis returns to 80% of the original

strength (Silver and Christiansen 1999; Hess 2002) by improvement of tensile strength

and cellular organisation causing the wound to contract which in turn causes the scab to

fall off. Depending on the extent of injury, remodelling of the wound (i.e. collagen

matrix) continues for weeks to months and even years as the scar tissue is not strong

compared to the tissue it replaces.

2.2.3 Role of enzymes in wound healing

The composition of the ECM is influenced by the infiltration and migration of various

cells that have numerous immunological functions during the wound healing process.

This regulation for the infiltration and migration of the immune cells is effectively

orchestrated by signals received from both soluble and complexed ECM cytokines and

chemokines which produce ECM degrading, modifying and synthesising proteases

(Alexander and Werb 1989; Goetzl et al. 1996). Accordingly, the wound healing process

is tightly controlled by these proteolytic enzymes which are present within wound

exudates and these enzymes have the ability to cleave numerous substrate specificities as

shown in table 17 (appendix I). The activity of these proteases for a given substrate is

controlled at the transcriptional and translational level by regulating enzyme activation

and inhibition to cause a balance between tissue synthesis and degradation. Any

disruption in the control pathways can lead to the increase of protease activity within

wounds and this over expression of various proteolytic enzymes within wounds can lead

to the development of chronic wounds (Westerhof and Vanscheidt 1994; Morison et al.

2004) as discussed in section 2.5 (see below).


47

Proteases are classified depending on their catalytic mechanism into two types: (1)

endopeptidases – which degrade proteins by cleaving peptide bonds within proteins; (2)

exopeptidases – degrade proteins by hydrolysing the terminal amino and carboxyl ends

of proteins (Wysocki 1996; Westerhof and Vanscheidt 1994). The exopeptidases remove

amino acids either individually, or as dipeptidylpeptides (Westerhof and Vanscheidt

1994). However, the endopeptidases are further classified according to the mechanistic

role of their active site: (1) serines (represent trypsin and elastase); (2) cysteines (e.g.

papain and lysosomal proteases); (3) aspartics (exemplify by pepsin and cathepsin); and

(4) the MMPs which contain a metal ion (Westerhof and Vanscheidt 1994) e.g.

collagenase, gelatinase, and stromelysin (Wysocki 1996) and membrane-type MMPs (MT-

MMPs).

Essentially there are two important families of proteases involved in wound healing: the

serine proteases and the MMPs (Morison et al. 2004; Xue et al. 2006; Vaday and Lider

2000) during the process of matrix degradation and tissue remodelling (Wysocki 1996).

Proteases have a high specificity of recognising seven or more amino acids that are

flanked on either side of the scissile P1–P1′ bond of peptide sequences (Schechter and

Berger 1967) of any particular substrate. The broad substrate specificities for the serine

proteases and MMPs involved in wound healing within the body are summarised by table

17 (appendix I). To control the cleaving or the activation of the broad substrate

specificities by each protease, the proteases are initially secreted as their inactive

zymogens1 (except for elastase, Vaday and Lider 2000) which are eventually activated by

other mediators and proteases when required during any phase of the wound healing

process. Nevertheless, once elastases are synthesised they are intracellularly stored within

inflammatory cells (i.e. neutrophil, monocytes and macrophage) in their active forms

(Vaday and Lider 2000) and in view of that it is crucial that activity of elastases is

stringently controlled. Herein, the description for the roles of the proteases involved in

wound healing is limited to elastase (neutrophil) and MMPs which as previously

encounter in chapter 1 are proteases that are reported as being elevated in chronic

wounds. At the end of this section, figure 16 summarises the functions of these

proteases within acute wound healing.

1 also known as pro-enzymes, latent/inactive enzymes


48

2.2.3.1 Serine proteases

During the wound healing process, serine proteases play an important role during

haemostasis (coagulation and fibrinolytic systems), inflammation, proliferation and

degradation of extracellular proteins. Thrombin, plasmin and neutrophil elastase (NE)

are all serine proteases. Serine proteases are a large family of enzymes that contain a

highly reactive serine residue (hence the family name) within the conserved catalytic triad

(or charge-relay system): His57–Asp102– Ser195 (chymotrypsin numbering) of the active

site. All serine proteases share a general catalytic mechanism via an acyl-enzyme

intermediate as highlighted by figure 8.

The serine residue of the active site behaves as the main nucleophile and the histidine

residue behaves as a general-acid/base catalyst which interacts with the side chain of an

aspartate residue (Copeland 2000). The proficiency of the serine protease is highly

marked due to the close vicinity and explicit interactions of both the serine and histidine

residues, since the nitrogen lone pair of the histidine side chain stabilises the highly

nucleophilic hydroxyl group of the serine residue (Voet and Voet 1995; Copeland 2000).

The role of the aspartate residues is unclear (Copeland 2000).

On penetration of a peptide substrate into the active site the mechanism is initiated and

the peptide substrate binds to the enzyme (in this case thrombin, plasmin or NE); and

the catalysis occurs in several steps. Firstly, a nucleophilic ‘addition’ attack at the

carbonyl carbon of the peptide substrate is initiated when the lone pair of the histidine

residue removes a proton from the serine residue of the catalytic triad As a result, a

high-energy unstable acylated tetrahedral intermediate (TI), the Michealis-Menten

complex (Voet and Voet 1995) is formed in the oxyanion hole of the enzyme. The

oxygen atom of the ester leaves the unstable acylated TI and the complex forms a highly

stable acyl enzyme with the serine residue as well as yielding an amine product (an amino

acid, R2–NH2). The reaction of this step is driven forward by the histidine residue which

behaves as a general acid to donate its proton to the deprotonated amino acid (R2– NH).

On departure of the amine product (R2–NH2) from the active site it is replaced with a

water molecule which is then deprotonated by the histidine residue which behaves as a

general base again. Via nucleophillic addition, the carbonyl carbon of the acyl-enzyme is

attacked by the water molecule to generate a deacylated TI. The restoration of the


49

carbonyl carbon dissociates the deacylated TI to yield an acid product and the free serine

protease is regenerated whereby the lone pair of the nitrogen atom of the histidine side

chain again stabilises the serine oxygen.

H OH

C

R1

O

NH

R2

O

NH2

O

NH2

N N HO

NH2

O

O

NH2

N N H

CON

H

R2

R1

HO

NH2

O

O

NH2

N N

CO

NH2

R2

R1

O

ONH2

O

HO

NH2

O

O

NH2

N N

CO

R1

H

O

H

H

O

NH2

O

O

NH2

N N

CO

R1

H

O

H

H OH O

NH2

O

NH2

N N

O

ONH2

O

C

OH

O

R1

..:

Acylation TI

+

..

Deacylation TI

:

[Asp]

[His]

Serine protease

Peptide substrate

Amine product

[Ser] [Ser]

[Ser]

[His]

[His]

Acyl enzyme

:

[Ser]

[His]

:

Hydration

[Ser]

[His]

[Asp]

[His]

Serine protease

[Ser]

+

Acid product

..

+

......

..........

..... .....

Figure 8. The hydrolysis of peptide substrate via the serine proteases (thrombin, plasmin or elastase). In the first instance the reaction proceeds by an acylation step to yield an amine product (an amino acid, R2–NH2) and an acyl-enzyme intermediate; followed by both an hydration and deacylation step. By doing so, an acid product is yielded and the serine protease (e.g. thrombin, plasmin or elastase) is regenerated. The tetrahedral intermediates are abbreviated by TI; R1 and R2 are amino acids of a peptide substrate. Details of the mechanism are given in the text. Figure adapted from Farley et al. 1997; Stryer 1995.

It should be borne in mind that although serine proteases share a general mechanism of

catalysis, they are highly specific in their ‘substrate specificity’ and this unique distinction

amongst the proteases of this family is maintained by the preference of the enzyme to

recognise and hydrolyse particular scissile P1–P1′ bonds of peptide substrates depending

on certain amino acid residues (Schechter and Berger 1967; Lauwers and Scharpe 1997).


50

From table 17 (appendix I) it can be seem that even though thrombin, plasmin and NE

are able to cleave or activate the same substrates during the wound healing process their

chosen substrate specificity of the scissile bond differs. Elastases are stated to cleave

peptide bonds after small hydrophobic residues such as Val~X and Ala~X (Lauwers and

Scharpe 1997; Stryer 1995; Purich and Allison 2002). The preference of these residues by

elastases depends on the type of elastase for example human NE (HNE) preferentially

cleaves adjacent to valine residues at the P1 position (Owen and Campbell 1999; Purich

and Allison 2002) whereas porcine pancreatic elastase (PPE) prefers to cleave after

alanine residues (Purich and Allison 2002). The substrate specificity of thrombin and

plasmin is trypsin-like except it is more restricted compared to trypsin (Barrett et al.

2004). Both proteases have similar preference for hydrophilic positively charged residues

within peptide substrates and inhibitors; thrombin prefers to cleave Arg~X bonds (for

examples of full sequences, see table 17 in appendix I) whereas plasmin prefers to cleave

Lys~X and Arg~X bonds (Barrett et al. 2004).

Besides their main roles, thrombin, plasmin and NE have other functions that are

important during the wound healing process. Earlier the prime function of thrombin

and plasmin was briefly covered in section 2.2.2.1 as proteases that control the proper

regulation of both the coagulation and plasminogen pathways during the haemostasis

phase to control, stop and restore vascular bleeding upon tissue injury. Although

thrombin and plasmin are reported as being elevated in chronic wounds their

multifunctional roles will not be discussed within this section as the wound healing

process of chronic wounds is reported as being deadlocked during the inflammation/

proliferation phase (Ayello et al. 2004a; Morison et al. 2004) as mentioned in section 2.5.

As NE plays an important role during the inflammation phase, its multifunctional roles

are now discussed.

Elastase is a broad term that characterises proteases which have the ability to degrade

and solubilise elastin via elastolysis, and such proteases are said to have elastinolytic

activity as depicted in table 4 (Farley et al. 1997; Owen and Campbell 1995). As elastases

have multifunctional roles during tissue repair, confusion can arise when understanding

the subclass of the type of elastase that is synthesised/ secreted by cells as generally

enzymes are grouped according to their substrate specificity, native structure, the

catalytic active site that they possess or even pH (Owen and Campbell 1995). Neutrophil


51

elastase (NE, entry 1 in table 4) and macrophage elastase (MMP-12, entry 6 in table 4) are

commonly defined as the two major elastases that regulate the wound healing process.

NE functions during the early stages (i.e. haemostasis and especially inflammation);

whereas the MMP-12 functions during the final stages (proliferation and tissue

remodelling) of the wound healing process. This section discusses the roles of NE,

whereas the next section (2.2.3.2) covers the role of MMP activity, hence MMP-12

during normal wound healing.

Table 4. Human proteases which express elastinolytic activity

Entry Protease Cell Type Catalytic family Active site Optimum pH

1 Elastase Neutrophil

2 Cathepsin G Neutrophil

3 Proteinase 3 Neutrophil

Serine protease

Catalytic triad Asp-His-Ser

Neutral

(pH 7 – 9)

4 Collagenase Neutrophil

5 Gelatinase Neutrophil

6 Metalloproteinase Macrophage

Metalloproteinase

Zn2+

coordinated to amino acids

Neutral

(pH 7 – 9)

7 Cathepsin B, L, S Macrophage Cysteine proteinase Cys, His Acidic

(pH 5 – 6)

(Adapted from Farley et al. 1997; Owen and Campbell 1995; Barrick et al. 1999)

In response to inflammatory stimuli of various cytokine signals, neutrophils are the first

inflammatory cells attracted to the wound area to combat and disinfect the wound from

microbial infection and proteolyse the damage tissue by releasing caustic proteases such

as NE that are stored within the peroxidise positive primary lysosome-like, azurophilic

granules of neutrophils. NE (EC 3.4.21.37) is sometimes referred to as human NE

(HNE), leukocyte NE (LNE) and polymorphonuclear (PMN) elastase. It is a

glycoprotein of 218-residues consisting of two asparaginyl N-linked carbohydrate side

chains with four disulfide bridges and the catalytic triad His41–Asp88–Ser173 of NE is

located within the centre of the reactive site (Takahashi et al. 1988; Lee and Downey

2001) and has a preference of hydrolysing Val~X bonds (e.g. X-Ala-Ala-Pro-Val~Y)

more than Ala~X.

During the wound healing process, NE is considered as the most influential and

destructive serine protease as it is proficient at simultaneously carrying out a range of

functions alongside its principal function of degrading the highly cross-linked fibrous


52

macromolecule of the ECM, elastin into soluble peptides at neutral pH (Takahashi et al.

1988, Prager et al. 1991; Baxter 1994; Owen and Campbell 1995). Initially NE facilitates

the host defence mechanism of the innate immunity in conjunction with wound

debridement by digesting the cell wall of gram-negative bacteria (Takahashi et al. 1988;

Farley et al. 1997; Chua and Laurent 2006) and eventually phagocytosing and engulfing

the bacteria into neutrophils where they are lysed and then digested by the potent active

NE encapsulated within neutrophils (Owen and Campbell 1995; Chua and Laurent

2006).

After phagocytosis, NE has the ability to directly or indirectly speed up tissue damage by

digesting and destroying the majority of the proteins of both the connective tissue and

the ECM. It is able to cleave proteins of both the coagulation and complement pathways

(Takahashi et al. 1988) including endogenous inhibitors such as antithrombin III (AT III)

to increase fibrin deposits and in this fashion supports the pro-coagulant activity during

the coagulation pathway (Prager et al. 1991). On the other hand, NE modulates

thrombin activity in order to decrease the stimulation of platelets to reduce the clotting

of fibrinogen (Brower et al. 1985; 1987) by degrading factor XIII (Wohner 2008) in this

way factor XIIIa is no longer generated to convert insoluble fibrin into crosslinked fibrin

monomers. Additionally NE has the ability to inactivate both the extrinsic and intrinsic

coagulation pathways by inhibiting coagulation factors such as factors VII, VIII, IX and

XII (Wohner 2008). During the fibrinolytic pathway, in parallel to plasmin, NE has the

capacity of directly digesting the fibrin clot into FDPs (Kluft 2003-2004) together with

providing an alternative pathway which indirectly facilitates the dissolution of the fibrin

clot (Wohner 2008). In the alternative pathway, NE directly cleaves 4 kringle domains of

the native plasminogen structure to activate the conversion of plasminogen into mini-

plasminogen (Wohner 2008) which is a.k.a K1-4 or angiostatin (Hoffman et al. 1998).

Mini-plasminogen is readily converted by PAs (i.e. uPA or tPA) into mini-plasmin which

is more proficient at digesting cross-linked fibrin in contrast to plasmin because mini-

plasmin is less likely to be attacked and inhibited by α2–antiplasmin (Wohner 2008). In

this manner, the stability of the fibrin clot varies amongst both pathways i.e. the FDPs

created by elastase-mediated hydrolysis are persistently released as much longer strands in

contrast to plasmin-mediated hydrolysis which rapidly dismantles the fibrin clot and

liberates associated FDPs (Wohner 2008).


53

In addition to the above, NE is proficient in the turnover of many other ECM

components/proteins, inflammatory mediators and membrane molecules such as

cleaving fibrin, fibronectin, laminin, vitronectin, proteoglycans, collagens and gelatin

(Baxter 1994; Dovi et al. 2004; Xue et al. 2006; Vaday and Lider 2000; Wohner 2008)

alongside other substrates as summarised in table 17 (appendix I). Furthermore, it

controls the conversion of cytokines into active/ inactive forms (Dovi et al. 2004) and

similarly regulates the activation of pro-MMPs to active MMPs (Zhu et al. 2001; Fleck

and Chakravarthy 2007). In doing so, NE possess many functions such as activating:

platelets, leukocytes (i.e. monocytes/macrophages), cytokine secretion and the

complement process (Barrett et al. 2004); as well as efficiently assisting in the migration

of neutrophils (Farley et al. 1997) across the wound bed seeing as active NE is non-

covalently translocated within the plasma membrane of inactive/active neutrophils

(Vaday and Lider 2000; Champagne et al. 1998) but mainly by active neutrophils (Owen

and Campbell 1995). The expression of membrane-bound NE is individually or

sequentially upregulated by TNF-α or via NE activating IL-8 (Vaday and Lider 2000;

Chua and Laurent 2006). Unlike free NE that is present in the extracellular space,

membrane-bound NE is not inhibited by antiproteinase inhibitors because it is resistant

to them (Owen and Campbell 1995). These multiple functions enables NE to properly

systematise the reconstruction and remodelling of the connective tissue (Farley et al.

1997) or ECM during tissue repair. Overall the direct and indirect cascade reactions

mediated by NE are summarised in figure 9.

Table 5. Systemic and alarm proteinase inhibitors to control HNE activity

Class Inhibitor MW (kDa) Synthesised/ secreted Type of Inhibition

Systemic α1-antiproteinase (α1-PI) 52 Hepatocytes � plasma Irreversible

Systemic α2-macroglobulin (α2-MG) 725 Hepatocytes � plasma Partial

Alarm SLPI 11.7 Epithelial, mast,

neutrophils, macrophages

Reversible

Alarm Elafin 9.9 Epithelial, keratinocytes Reversible

(Tailored from Barrick et al. 1999; Fitch et al. 2006; Barrett et al. 2004).

The secretion and cellular biochemical functions of NE activity are stringently controlled

at various levels by endogenous systemic or alarm proteinase inhibitors such as: α1-

antiproteinase inhibitor (α1-PI; previously known as α1-antitrypsin), α2-macroglobulin (α2-


54

MG), secretory leukocyte elastase inhibitor (SLPI) and elafin (Barrick et al. 1999; Fitch et

al. 2006; Barrett et al. 2004) as summarised in table 5.

From the systemic inhibitors, α1-PI is the most potent irreversible inhibitor as it inhibits

~92% of elastase activity by covalently binding to NE to form an NE-α1-PI complex and

in this way NE is no longer available to bind any of it substrates (Fitch et al. 2006;

Barrett et al. 2004; Medina et al. 2005; Janoff 1985). The remaining systemic inhibition is

achieved by α2-macroglobulin which structurally inhibits NE by partially hindering its

conformation. The alarm inhibitors, SLPI and elafin inhibit elastase activity reversibly

(Fitch et al. 2006; Barrett et al. 2004).

Figure 9. Multifunctional roles of elastase during the wound healing process. Reactions of activation (maroon and red arrows), conversion (black arrow) and inhibition (blue text and arrows). Abbreviations: SLPI (secretory leukocyte elastase inhibitor); α1-PI (α1-antiproteinase inhibitor); IL (interleukin); TNF–α (tumour necrosis factor alpha); α2-MG (α2-macroglobulin); TGF–β (transforming growth factor beta), MMPs (matrix metalloproteinases), TIMPs (tissue inhibitors matrix proteinases); ECM (extracellular matrix), FDP (fibrin degradation products). Figure tailored from the text within this chapter and Lauwers and Sharpe 1997. For a detail description of this figure see text within this section.

2.2.3.2 Matrix metalloproteinases

MMPs (also known as matrixins) constitute a large family of zinc-dependant and calcium

activating endopeptidases and over 20 proteases of this family have been identified

(Nagase and Woessner 1999; Barrick et al. 1999; Eming et al. 2008) and each MMP is

structurally and functionally related (Woessner and Nagase 2002). Initially MMPs were

ELASTASE (active)

Pro-elastase (inactive) Activates cells

(e.g. neutrophils, macrophages, kerinocytes)

Bacterial phagocytosis

↑ Chemokines/ cytokines (e.g. IL-1β; IL-8; TNF–α)

α1-PI

MMPs

ECM DEGRADATION

Pro-MMPs

TIMPs

Plasminogen

Mini-plasminogen

Activate/ Inactivate coagulation process

(e.g. thrombin, factors XIII, VII, VIII, IX, XII)

α1-PI α2-MG

SLPI & Elafin

Mini-plasmin

PAI

PAs

TGF – β (inactive)

(Fibrinolysis)

Fibrin Clot FDP

TGF – β (active)

Angiogenesis

Inactivate AT III, α2- antiplasmin, CI inactivator


55

classified according to their substrate specificity into various subgroups such as:

collagenases. gelatinases, stromelysins, matrilysins, membrane-type MMPs and a

heterogeneous subgroup termed as other MMPs. Given that the substrate specificity of

some MMPs overlaps (Somerville et al. 2003) they are now further categorised into

various subtypes depending on the structural domains contained within their structure as

summarised by figure 10 (Nagase and Woessner 1999; Somerville et al. 2003; Cawston

and Wilson 2006).

Figure 10. The conformational domains of human MMPs. MMPs are structurally related as they contain 3 distinct domains: a SP (for secretion from cells), a highly conserved N-terminal pro-domain (which specifies their latency) and a catalytic domain containing 3 histidine residues bound to Zn2+ and 2Ca2+ ions within the active site of MMPs (authorises enzymatic activity). The smallest MMPs are group A (MMP-7, -26); groups C (MMP-2) and D (MMP-9) both contain 3 fibronectin type II repeats enabling them to bind gelatin, collagen and laminin; the type V collagen-like domain within MMP-9 makes it structurally and functionally distinctive from MMP-2. The hemopexin domain (groups B – G) determines substrate specificity; whereas the furin recognition motif (RXKR) within the MMPs of groups E – H permits intracellular activation by furin-like proteinases. Furin-recognition MMPs are either secreted (group E) or anchored to the cell surface (groups F, G, H) either by a glycosylphosphatidyl (GPI) domain (MMP-17, -25) or via a transmembrane (type I)/ cytoplasmic domain (MMP-14-16, -24). Exclusively, group H (MMP-23) contains an N-terminal transmembrane (type II) within its SP, a Cys/Pro-rich (cysteine/proline) domain with either a C-terminal interleukin-1 (IL-1)/ immunoglobulin-like (Ig) domain. Figure adapted from Nagase and Woessner 1999; Woessner and Nagase 2002; Visse and Nagase 2003; Somerville et al. 2003; Cawston and Wilson 2006.


56

In relation to section 2.6.3.3 (below) various researchers have designed protease-

modulating wound dressings (with or without inhibitors) to catalytically inactive the

enzymatic activity of MMPs within chronic wounds. At this stage, it is crucial to

understand the activation of MMPs which is structurally achieved by the relationship of

the pro-domain with the catalytic domain. Once the MMPs are secreted by cells

extracellulary (except for furin-like MMPs, as mention below) the signal peptide (SP) is

cleaved off during translation (Toriseva and Kähäri 2009) by endopeptidases; and the

MMPs are initially secreted as their zymogens usually termed as pro-MMP-X (where X is

a number). As the pro-MMPs are catalytically inactive, before they can exert their

function they are activated via ‘cysteine-switch’ mechanism (Van Wart and Birkdahl-

Hansen 1990) as summarised by figure 11.

Figure 11. Activation of MMPs via the ‘cysteine-switch’ mechanism. The active site of MMPs consists of 3 histidine (H) residues ligated to the catalytic Zn2+ (3His-Zn2+) or Ca2+ ions (not shown) within the catalytic domain. Potential substrates are unable to access the active site as it is blocked by the pro-domain and the thiol-Zn2+ interaction (i.e. Cys-S-Zn2+). MMPs are activated by modifying or interrupting the thiol-Zn2+interaction either chemically by reactive oxygen species (ROS) or physiologically by proteinases (such as elastase, plasmin, uPA, furin and other active MMPs) produces an intermediate MMP. Consequently the cleavage of the N-terminal pro-domain (blue arrow) by proteinases or by autolysis (e.g. via MMP intermediate itself) produces an active MMP. Figure adapted from Somerville et al. 2003; Lint and Libert 2006.


57

The N-terminal pro-domain contains a highly conserved Pro-Arg-Cys-Gly-X-Pro-Asp

(where X is a Val/Asn; PRCGXPD) sequence (Nagase and Woessner 1999; Somerville et

al. 2003; Toriseva and Kähäri 2009) at positions 173 – 179 (Woessner and Nagase 2002).

This sequence preserves the latency of pro-MMPs because the unpaired thiol (sulphydryl)

group of the cysteine residue within the PRCGXPD sequence is coordinated opposite

the catalytic Zn2+ ion that is bound to 3 histidine (His) residues (i.e. 3His-Zn2+) of the

highly conserved zinc binding sequence: His-Glu-X-Gly-His-X-X-Gly-X-X-His-Ser

(HEXGHXXGXXHS, where X is any amino acid) at the active site within the catalytic

domain (Nagase and Woessner 1999; Somerville et al. 2003; Toriseva and Kähäri 2009).

Since the thiol-Zn2+ interaction (shown as Cys-S-Zn2+ in figure 11) blocks all proteolytic

activity, during the “cysteine-switch” mechanism the thiol-Zn2+ interaction initially

undergoes modification or interruption either chemically by reactive oxygen species

(ROS) or by a variety of active proteases (such as elastase, plasmin, uPA, furin and other

MMPs) and as a result an MMP intermediate is produced (Somerville et al. 2003; Lint

and Libert 2006; Ra and Parks 2007; Toriseva and Kähäri 2009).

Subsequently within the active site, the thiol group is replaced by water and the activation

process is completed after the pro-domain is proteolytically cleaved off directly by

proteases (Somerville et al. 2003; Lint and Libert 2006) or autolysed by the MMP

intermediate itself (Ra and Parks 2007; Toriseva and Kähäri 2009). Consequently a

biologically active MMP is formed (figure 11), permitting the penetration of potential

substrates into the exposed active site of MMPs; along with 3His-Zn2+, water molecules

and the stabilising Ca2+ binding sides within the catalytic domain enable MMPs to

undergo enzymatic activity of various substrates as summarised by table 17 (appendix I).

Essentially, the activation of furin-like MMPs (containing the furin recognition motif, e.g.

MMP-11, -23 and -28 and MT-MMPs) is achieved intracellularly by furin (an convertase)

during the secretory pathway (Somerville et al. 2003; Ra and Parks 2007; Toriseva and

Kähäri 2009) of the Golgi-body.

During the wound healing process, MMPs are the normal physiologic mediators that are

collectively controlled to preclude substantial matrix degradation (Wysocki 1996) of

majority of the ECM components. The MMPs specifically involved in wound healing

are the collagenases, gelatinases and stromelysins (Morison et al. 2004) including elastases


58

(e.g. MMP-12) and each have a broad substrate specificity as summarised in table 17

(appendix I). Majority of the MMPs are involved in re-epithelisation, remodelling and

the migration of various cells during the healing process (Greener et al. 2005). The

function of the collagenases (i.e. MMP-1, -8 and –13) is to specifically degrade

connective tissue (Lijnen 2002; Morison et al. 2004) and initiate the splitting of the triple

helix of the native fibrillar collagens (Lobmann et al. 2005; Yager and Nwomeh 1999) or

more precisely they are involved in normal and pathologic remodelling of collagen. Each

collagenase is specific to the type of collagen produced by granulocytes, macrophages,

epidermal cells, and fibroblasts (Westerhof and Vanscheidt 1994). During inflammation

fibroblasts, the epidermal and endothelial cells secrete MMP-1 (Lobmann et al. 2005) and

table 17 (appendix I) displays all the collagenases that are able to degrade: gelatin and the

various collagen types (I, II, III, and X). Compared to MMP-1, MMP-8 prefers to cleave

type I collagen more rapidly than type III collagen (Purich and Allison 2002).

Additionally, MMP-1 regulates the migration of keratinocyte and re-epithelialisation,

MMP-8 inhibits apoptosis and anti-inflammation; whereas MMP-13 enables endothelial

migration (Xue et al. 2006). Each collagenase is specific in the sequence it recognises and

hydrolyses, for examples of the types of sequences recognised by collagenases see entries

4, 5 and 6 in table 17 (appendix I)

Once the collagenases have completed the initial cut into the helical structure of collagen,

then the gelatinases, MMP-2 (gelatinase A, 72-kDa) and MMP-9 (gelatinase B, 92 kDa)

can further degrade the partially denatured collagens i.e. type I and III (Lobmann, R., et

al., 2005) by binding the collagen via the three fibronectin type II repeats within the

catalytic domain of their structure (Visse and Nagase 2003). The gelatinases also mediate

the digestion of the non-fibrillar collagen types IV, V, VII and X (Birkedal-Hasen 1995

as cited in Lobmann et al. 2005) including the basement membrane (BM) (Morison et al.

2004), gelatin and laminin (Visse and Nagase 2003). Of all the MMPs, MMP-2 is the

most common protease as it is continuously produced during the wound healing process

since it is expressed by various cells (e.g. fibroblasts, keratinocytes, endothelial cells and

monocytes; Lobmann et al. 2005) and it is activated by various membrane-type MMPs (in

particular, MMP-14, -16, -24 and -25; Xue et al. 2006) during various stages of the

healing process. Specifically, MMP-2 assists in keratinocyte migration, anti-inflammation

and activates pro-MMPs-1, -9 and -13 (Xue et al. 2006). Compared to MMP-2, MMP-9

is expressed and synthesised by keratinocytes, monocytes and macrophages (Lobmann et


59

al. 2005) to maintain inflammation and assists in neutrophil migration (Xue et al. 2006).

In contrast to collagenase, the gelatinases recognise and hydrolyse sequences containing

Gly~Ile bonds (see entry 7: MMP-2 and entry 8: MMP-9 in table 17; appendix I)

The stromelysins: MMP-3 (Stromelysin-1), MMP-10 (Stromelysin-2) and MMP-11

(Stromelysin-3) are known to degrade ECM proteoglycans (Morison et al. 2004;

Lobmann et al. 2005). MMP-3 and MMP-10 have a broad specificity to cleave many

substrates such as: collagen type III, IV; gelatin, elastin, MMP-8 (table 17, appendix I).

Both MMP-3 and MMP-10 enable cell migration; additionally MMP-3 facilitates within

the fibrinolytic pathway by directly mediating the degradation of fibrin into FDPs (Kluft

2003-2004) as well as hydrolysing α2-antiplasmin (Medina et al. 2005); finally MMP-3 has

the ability to activate pro-MMP-1, -7, -8, -9, and -13; and it recognises and hydrolyses

following sequences: Arg-Arg-Lys-Pro-Val-Glu~Z-Trp-Arg-Lys and Arg-Pro-Leu-

Ala~X-Trp-Arg-Ser (see entry 9 in table 17, appendix I) whereas MMP-10 facilitates re-

epithelialisation (Xue et al. 2006) and cleaves sequences Asp-Val-Gly-His~Phe-Ser-Ser-

Phe and Gly-Pro-His-Leu~Leu-Val-Glu-Ala (see entry 10 in table 17, appendix I). Anti-

proteolytic activity is inhibited by MMP-11 as it is able to hydrolyse the Ala350 – Met351

bond of the α1-proteinase inhibitor (Purich and Allison 2002) also cleaves sequences

containing X-Ala~Met-Y (see entry 11 in table 17, appendix I).

As mentioned earlier, NE functions during the early stages of the wound healing process;

after neutrophils have completed their function and die; macrophages are then recruited

from the blood circulation (Mutsaers et al. 1997). Macrophages secrete another type of

elastase, human macrophage elastase (HME) or more commonly MMP-12. MMP-12

functions during the final stages of the wound healing process and it continues with

phagocytosing bacteria and has a broad substrate specificity of digesting various proteins

of the ECM (summarised in table 17, appendix I). Its prime substrate is elastin (Chen

2004); in conjunction with its elastolytic activity, MMP-12 regulates the successful

migration of macrophages (Visse and Nagase 2003) alongside encouraging the efficient

proliferation of epithelial cells (Xue et al. 2006), plus has the ability to self regulate its

own activation via autolytic processing and by doing so it amplifies various proteolytic

processes during tissue repair by activating MMP-2 and MMP-3 (Chen 2004). MMP-12

recognises and hydrolyses the following sequences: Leu-Val-Glu-Ala~Leu-Tyr~Leu-


60

Val-CysO3H-Gly; Arg-Pro-Leu-Ala~Leu-Trp-Arg-Ser; Arg-Pro-Phe-Glu~Val-Lys-Asp-

Thr; and Gly-Ala-Met-Phe~Leu-Glu-Ala-Ile (see entry 12 in table 17, appendix I).

In recent years, matrilysins, membrane-type MMPs (MT-MMPs) and other MMPs have

been identified as playing a role in the wound healing process (Xue et al. 2006). The

function of each of these MMPs has been summarised in table 6 and since these

proteases have not been reported in chronic wounds (except MMP-19; Xue et al. 2006)

their function and substrate specificity will not be discussed in here; however they have

been extensively reviewed elsewhere (Visse and Nagase 2003; Barrett et al. 2004).

Table 6. Summary for the function of human matrilysins, MT-MMPs and other MMPs during in the wound healing process.

Proteases EC Function in wound healing process

Matrilysins

MMP-7 (Matrilysin-1)

MMP-26 (Matrilysin-2)

3.4.24.23

3.4.24.

Directly mediates fibrin digestion. Controls inflammation, re-

epithelialisation, and inhibits apoptosis

Possible role in cell migration

Membrane-type MMPs

MMP-14 (MT1-MMP)

MMP-15 (MT2-MMP

MMP-16 (MT3-MMP)

MMP-24 (MT4-MMP)

MMP-25 (MT6-MMP)

3.4.24.

3.4.24.

3.4.24.

3.4.24.

3.4.24.

Keratinocyte survival and activates pro-MMP2 and MMP-2;

directly mediates fibrin digestion.

Anti-apoptosis

Activates MMP-2

Activates MMP-2

Maintains migration of neutrophils, activates MMP-2

Other MMPs

MMP-19

MMP-20 (Enamelysin)

MMP-28 (Epilysin)

3.4.24.

3.4.24.

3.4.24.

Assists in macrophages migration within the wound bed

Controls enamel formation

Enables epithelial cell proliferation

(Xue et al. 2006; Kluft 2003/2004; Purich and Allison 2002).

MMPs are tightly regulated during transcription (gene expression and secretion), post-

transcription level by activating the MMP zymogens by proteolytically cleaving the pro-

domain (as described above) and finally by controlling the proteolytic activity of the

active MMP by inhibitors. At the extracellular level, TIMPs are the endogenous

inhibitors that inhibit MMP activity and four members of this multigene family have

been identified as TIMP-1, -2, -3 and -4 (Visse and Nagase 2003; Dasu et al. 2003;

Stamenkovic 2003). Physiologically each of the TIMPs form irreversible complexes with

MMPs (Dasu et al. 2003) by directly binding to the Zn2+ ion at the active site within the

catalytic domain (Toy 2005). In tissue fluids/ circulation, the inhibition of MMP activity

is inactivated by the non-specific inhibitor, α2–MG in a ‘bait and trap’ mechanism


61

(Somerville et al. 2003). When sites within α2–MG are cleaved, this enables α2–MG to

bind and form a cocoon around the MMP forming an irreversible complex which is

attacked and removed by scavenger receptors (Somerville et al. 2003; Stamenkovic 2003)

and the active MMP is segregated from its substrates (Somerville et al. 2003).

The combined mechanism of plasmin, elastase and MMPs to degrade components of the

ECM with the intention of making space for cells to efficiently migrate across the ECM

tissue space in order to facilitate the synthesis of a functional ECM during tissue repair is

summarised by figure 12.

Figure 12. Reciprocal interpretation for the role of plasmin, elastase and MMPs in achieving ECM degradation during the wound healing process. After leukocytes have migrate across the endothelial cells into the tissue space, activated endothelial and epithelial cells secrete pro-inflammatory mediators which activates the transcription of MMP, in addition to causing leukocytes to secrete pro-MMPs, MMPs/ADAMs (a disintegrin and metalloprotease), elastase, heparanase and uPA/tPA. Elastase alongside plasmin activates pro-MMPs into active MMPs and all simultaneously degrade/ modify components of ECM with the intention of producing a functional ECM. Reactions of activation (black arrows), enzymatic activation (red arrows), conversion (blue arrows), secretion (green arrows), positive feedback (+ve) and negative feedback (-ve). Figure adapted from Silver and Christiansen 1999; Vaday and Lider 2000; and the figures/ text within this chapter.


62

2.3 TYPES OF CHRONIC WOUNDS

Chronic wounds (a.k.a abnormal wounds, Boyce 1996) become disrupted in the

inflammatory and proliferation stages and for that reason wound closure is incomplete as

the wounds struggles to undergo re-epithelialisation within the required time (Ayello et

al., 2004; Morison et al. 2004). Chronic wounds are the result of various pathological

conditions such as diabetes mellitus, poor circulation and nutrition, immunodeficiency’s

(Hess 2002). There are other factors that delay healing such as: infection, pressure,

trauma, edema, necrosis (Hess 2002). There are many types of chronic wounds (as

summarised in table 7) and as it can be seen some of these chronic wounds are the result

of an ulcer which is characterised as inflamed and painful open sores on the skin/

mucous membrane that cause devastation to the tissue surface and are usually referred to

as dermal ulcers (Hess 2002; Morison et al. 2004).

Table 7. Types of Chronic wounds

Ulcers Other chronic wounds

Arterial ulcers

Diabetic ulcers (foot)

Pressure sore ulcers

Trauma ulcers

Vascular ulcers (venous, stasis)

Burn wounds (2nd, 3rd degree)

Sickle cell ulcers

Surgical wounds

Tunnelling wounds

Various authors have written detailed descriptions (including the definitions and

classifications) of such wounds (Cohen et al. 1992; Collins et al. 1992; Eaglstein and

Falanga 1997; Hess 2002; Baranoski and Ayello 2004; Morison et al. 2004). By

understanding the precise pathogenesis of chronic wounds it will enable researchers to

understand the multidisciplinary approach of wound management that can be used to

heal the aetiology of such wounds.

2.4 GRADIENTS OF CHRONIC WOUNDS

In chronic wounds there are a number of gradients: oxygen, temperature and pH. The

variation of these gradients can affect the pathophysiology of chronic wounds.


63

2.4.1 Oxygen

The level of oxygen plays a crucial role for skin repair as it is highly demanded. Blood

circulation and the atmosphere are the only two sources that supply oxygen to a chronic

wound (Church 2001). An oxygen gradient is present between the blood circulation

and the wound area, as hypoxia is usually the stimulus that causes the activation of

angiogenesis to maintain haemostasis by the development of new blood vessels. Church

(2001) reports that it’s difficult to characterise the micro-environmental dynamics of

oxygen distribution.

2.4.2 Temperature

In most environments, the edge of the skin is much cooler compared with core

temperature of the body (Church 2001; Xia et al. 2000) which is at 37oC. Many factors

(i.e. the blood supply to the wound, whether or not the wound is covered or exposed,

whether or not the wound is synthetically heated) can significantly cause temperature

variations, that is gradients of chronic wounds (Church 2001). For example “necrotic

tissue is ‘cold’, and will remain more or less at ambient temperature” (Church 2001).

Bello et al. averagely reported the wound temperature of an in vivo wound to be on

average 3.1oC below the core temperature (Bello et al. as cited in Xia et al. 2000). For

over a week Xia et al., maintained their experiments at 33.9oC + 0.1 and later the

temperature of their cell culture was modified to 33.0oC to accommodated the

temperature in the range of 32 – 33oC (see citations within Xia et al. 2000).

2.4.3 pH

The effect of pH on wound healing has only been studied by a few researches.

Variations in pH within wounds is known to affect protease activity. Church (2001)

reported that the pH of chronic wounds is low where necrosis is taking place and its

normal where there is viable cellular activity. This statement contradicts Leveen (1973)

observations who showed that acidification of wound surfaces increases wound healing

by increasing the amount of oxygen at the wound bed. It has been reported that an

acidic pH wound fluid is more beneficial for antibacterial activity (Varghese et al. 1986).

Nevertheless, Chai (1992) reported that the pH of chronic wounds (more precisely

granulation of burn) varies with the amount of bacteria present within the wound. They


64

reported that when the number of Escherichia coli (E.coli) or Staphylococcus aureus (S.

aureus) is above 10 (7 organisms)/gm of granulation tissue the pH of the wound is 6.7;

and when the number of Pseudomonas aeruginosa (P. aeruginosa) is 10 (8 organisms)/gm

of granulation tissue then the pH is 8.0 of the wound.

It has been suggested that the pH of human wounds typically ranges between pH 6.0 –

8.0 (Ayello et al. 2004). Dissemond et al. (2003) demonstrated that the pH of chronic

wounds ranges between 5.45 – 8.65 and similarly this range corresponds to the data given

by Prager et al. (1994) who reported that the range of pH is from pH 5.3 – 8.4 for burn-

wound exudate. Prager et al. (1994) found that by increasing the pH causes an increase

in protease activity. In doing so, chronic wound proteases efficiently have a tendency to

degrade proteins at neutral to slightly alkaline pH (i.e. pH 7 – 8, Greener et al. 2005). It

has been accounted that in order to facilitate wound healing the pH of the wound

environment should be lowered to 4 (Schultz et al. 2005b) or greater than 4 and in

sequence to escape degradation of newly formed ECM the pH must be lower than 7

(Greener et al. 2005) making the wound environment acidic (Schultz et al. 2005b). It

has been stated, that protease activity can be reduced by changing the pH of the wound

bed (Greener et al. 2005) using topical protease-modulating wound care products such as

Cadesorb (for further details of this hydrogel dressing, see section 2.6.3.3).

2.5 ROLES OF ENZYMES IN CHRONIC WOUNDS

It is not clearly understood why the healing process is delayed for chronic wounds

(Stroock and Cabodi 2006). Nevertheless, there has been increasing evidence at both the

biochemical and molecular level signifying the over expression of various proteases such

as neutrophil elastase, MMPs, plasmin and thrombin to cause the healing of chronic

wounds to become delayed in comparison to acute wounds (Wysocki et al. 1993;

Wysocki 1996; Westerhof and Vanscheidt 1994; Grinnell and Zhu 1994, 1996; Bullen et

al. 1995; Harris et al. 1995; Yager et al. 1996, 1997; Yager and Nwomeh 1999; Vaalamo

et al. 1997; Tarnuzzer and Schultz 1996; Stadelmann et al. 1998; Trengove et al. 1999;

Cullen et al. 2002; Morison et al. 2004; Edwards et al. 2004; Ayello and Cuddigan 2004;

Greener et al. 2005; Schultz et al. 2005a). As encountered in section 2.2.3, the chronic

wound proteases (i.e. NE and MMPs) were reported as having multifunctional roles


65

during acute wound healing where they either regulate or contribute to various protective

mechanisms to collectively restore tissue integrity after an injury; and their progression is

properly regulated via signalling events from growth factors and cytokines/ chemokines.

Essentially, NE is the fundamental predicament that disturbs and deadlocks the normal

tissue repair of chronic wounds since elastase activity is found to be much higher in

CWFs compared to acute wound fluids. For example Trengove et al. (1999) reported

that the concentration of NE in CWFs was in the range of 5.9 – 344 µg/ml (median of

99.5 µg/ml) for 6 patients (46.15 %) suffering from chronic wounds (total: 13 patients);

whereas the wound fluid from acute wounds, all showed NE activity to be less than 1

µg/ml (total: 14 patients). Similarly, Rao et al. (1995) reported chronic wounds to have a

10 – 40 fold increase of elastase activity compared to acute wounds and the authors of

this study as well as other others, report elastase to be exclusively responsible for the

degradation of fibronectin (Grinnell and Zhu 1994, 1996; Rao et al. 1995) and growth

factors (Yager et al. 1997). This is because there is an imbalance between elastase and its

endogenous inhibitors (Rao et al. 1995; Fleck and Chakravarthy 2007) and for that

reason elastase activity has been reported to be uninhibited due to low levels of α2-

macroglobulin (Yager et al. 1997) and essentially α1-PI in chronic wounds because they

are highly degraded by excess chronic wound proteases (Grinnell and Zhu 1994, 1996;

Rao et al. 1995) such as elastase (see figure 13). In doing so, the uncontrolled high levels

of elastase cause immense tissue or ECM destruction (see below).

Pro-MMPs are secreted by many cells (such as neutrophils, monocytes/ macrophages

and fibroblasts/ keratinocytes). Surplus elastase activity is the main culprit for

propagating the overexpression of MMPs in chronic wounds because elastases (such as

NE or MMP-12) activate the conversion of pro-MMPs into active MMPs which

consecutively induces the cascade of active MMPs to activate other pro-MMPs (Zhu et

al. 2001; Fleck and Chakravarthy 2007). Un-interruptedly, the activation cascade of

MMPs spirals out of control seeing as the ‘vicious cycle’ of elastase disseminates further

whereby the endogenous inhibitors of MMPs i.e. TIMPs (or α2-MG) become targets of

elastase as they are highly degraded resulting in an imbalance between MMPs and TIMPs

(or α2-MG). Since the protective tissue mechanisms are overwhelmed by high proteolytic

activity, subsequently the activity of elastase and MMP becomes out of control and

causes substantial degradation and turnover of both the pre-existing and newly formed


66

collagen of the ECM (Zhu et al. 2001; Fleck and Chakravarthy 2007) as well as degrading

vital ECM components that are required for normal wound healing e.g. ECM proteins/

their receptors and numerous mediators i.e. growth factors, cytokines/ chemokines and

their receptors (Grinnell and Zhu 1994; Rao et al. 1995; Yager et al. 1996; 1997; Snyder

2005; Morison et al. 2004).

Figure 13. The‘vicious elastase cycle’ causes extensive tissue/ ECM degradation promoting chronic inflammation via a positive feedback mechanism within chronic wounds. Neutrophils secrete high amounts of elastase which in chronic wounds elevated levels of elastase degrade antiproteinases (e.g. α1-PI) and TIMPs causing an imbalance in the equilibrium between (1) elastase levels and elastase inhibitors; and (2) MMPs and TIMPs. Elastase activates pro-MMPs into active MMPs, which in turn degrade α2-antiplasmin causing a rise in plasmin activity. Plasmin has a much lower proteolytic activity than elastase, because elastase activates the conversion of plasminogen into mini-plasminogen which inihibits angiogenesis as well as being converted to mini-plasmin to promote fibrinolysis alongside plasmin. Excess levels of elastase and MMP causes extensive ECM degradation resulting in the formation of a non-functional ECM as well as generating chemotactic products which recruit more neutrophils via a positive feedback mechanism to secrete more elastase and then whole vicious cycle begins again promoting extensive chronic inflammation within chronic wounds. Reactions of activation (black arrows), elastase reaction (dark red arrows); secretion (green arrows), inhibition (blue lines), and degradation (dotted black arrows). Figure adapted from Fleck and Chakravarthy 2007; Moseley et al. 2004).

NE activates macrophages which persistently secrete pro-inflammatory cytokines (e.g.

IL-8) to accelerate the influx of neutrophils by stimulating a cascade of reactions causing

neutrophils and macrophages to release elevated levels of elastase (i.e. NE and MMP-12,

respectively) which continually contribute to the ‘vicious elastase cycle’ of promoting

wound chronicity caused by immense tissue damage (Dovi et al. 2004; Fleck and

Chakravarthy 2007) and consequently over time, this hinders the formation of a


67

functional ECM resulting in a non-functional ECM, cell necrosis and apoptosis.

Accordingly, during chronic inflammation the degradation products (achieved through

the destruction of ECM; and elastase inhibitors i.e. α1-PI) possess chemotactic properties

which enable them to activate and recruit more neutrophils via a positive feedback

mechanism. In doing so, neutrophils secrete excessive amounts of elastase (Menke et al.

2007) and the whole ‘vicious cycle’ of elastase commences again as schematically

summarised by figure 13. Given that neutrophils are constantly present, the wound

healing process of chronic wounds persistently becomes stuck during the inflammation/

proliferation phase (Ayello et al. 2004; Morison et al. 2004).

Additionally, high levels of MMPs (specifically MMP-3, see section 2.2.3.2) degrade α2-

antiplasmin and therefore plasmin levels become elevated within chronic wounds. As a

consequence high plasmin levels facilitate in the conversion of pro-MMPs into active

MMPs and therefore enchances the brutal breakdown of ECM by MMPs as well as

causing immense fibrinolysis of the fibrin clot. On the contrary of plasmin being

elevated in chronic wounds, its concentration is much lower in contrast to elastase

(Cullen et al. 2002a; Hoffman et al. 1998) because NE promotes the conversion of

plasminogen to mini-plasminogen (i.e. K1-4 or K1-3, in which K = kringle) this

therefore reduces the level of intact plasminogen available in chronic wounds and

subsequently reduces the conversion of plasminogen into plasmin (Hoffman et al. 1998).

Mini-plasminogen is a specific potent inhibitor termed as angiostatin which inhibits

angiogensis in VLUs (Hoffman et al. 1998) as summarised by figure 13. Even though

thrombin is said to be elevated within chronic wounds (Schultz et al. 2005a) its function

within chronic wounds remains unclear. Nevertheless, removal of the ‘vicious elastase

cycle’ by reducing elastase activity within chronic wounds will hinder chronic

inflammation and stop the uncontrolled positive feedback mechanism that exists within

chronic wounds.

2.6 MANAGEMENT OF WOUND HEALING

The management of chronic wounds is highly researched as chronic wounds are a major

health issue since the number of patients suffering from chronic wounds increases each

year resulting in 2 million patients suffering from pressure ulcers and between 600,000 –

2.5 million suffering with chronic leg and foot ulcers (Stadelmann et al. 1998) and


68

eventually causing the deaths of millions per year. The fundamental management of

speeding up the healing of chronic wounds is not completely understood because the

pathophysiology of such wounds is still being investigated.

Essentially the wound care market is subdivided into two areas: traditional and advanced

wound dressings. The wound healing in presence of the traditional dressings functions

in dry surroundings whereas the advance dressings function within moist surroundings

(GMAWCP 2008). Moist surroundings are more acceptable than traditional dressings;

however traditional dressings are still used today (GMAWCP 2008). There are many

wound products available for the treatment of chronic wounds. Nevertheless, there is an

essential need to advance the wound care market due to the trauma that is still faced by

chronic wound sufferers.

2.6.1 Debridement of chronic wounds

A wound is initially debridement which is the removal of damaged tissue and it is a

means of facilitating the healing of a wound to become more efficient. In the wound bed,

on occasion debridement is crucial as damaged tissue may become infiltrated with

bacterial growth which behaves as a physical barrier to healing. It should be borne in

mind that the original methods of treating the wound are still of existence. There are

many methods that are used in the clinical practice to debride a wound and these include:

surgical, enzymatic, autolytic, mechanical and biological. In terms of enzymatic

debridement, the water-soluble collagenase preparation is the best proteinase used and it

functions to breakdown collagen (native or damaged) into gelatin (Falanga 2002).

2.6.2 Moist wound healing

Once the wound has been debrided then an appropriate wound dressing i.e. moist

dressing is chosen to allow the wound to heal within a moist environment. In 1962

George Winter (from Smith & Nephew, Inc.) introduced the concept of moist wound

healing as it was shown that the epithelisation of wounds occurs much faster under

occlusive dressing as such dressings uphold a moist wound area (Winter 1962; Winter

and Scales 1963). A moist environment is important for the infiltration and hence the

migration and movement of cells, growth factors, proteases, oxygen and delivery of


69

nutrients across the wound surface. This way the rate of epithelisation and angiogenesis

can be reached which then leads to the final stage of collagen synthesis and remodelling.

There are many moisture-maintaining dressings available: wounds of light – moderate

drainage dressings such as occlusive films, hydrogels, and hydrocolloids are usually

preferred. Whereas wounds of moderate – heavy exudates absorbent dressings such as

foams and alginates are usually selected as well as hydrogels. To date considerable

attention has been devoted in designing and developing moist wound dressings.

Several authors have described the range of wound products including wound dressings

to treat chronic wounds (Cohen et al. 1992; Collins et al. 1992; Hess 2002; Baranoski and

Ayello 2004). However, in this thesis the discussion will now summarise the hydrogels

dressings (passive, active and protease-modulating) to treat chronic wounds.

2.6.3 Management of chronic wounds using hydrogel dressings

Hydrogels are sometimes referred to as: stimulated/ responsive, smart, intelligent

hydrogels (Hoffman 2001; Galaev 1995; Galaev and Mattiasson 1999; Kim and Park

1998; Jeong and Gutowska 2002; Peppas et al. 2000; Ulijn 2006) because these polymers

essentially possess unique properties that allows them to change their dimension,

conformation or physical properties from a swell to collapse (shrink) state (or visa-versa)

or solution-to-gel transitions due to changes in the environmental conditions such as pH,

temperature, ionic strength, solvent, light (Kim and Park 1998; Hoffman 2001; Jeong and

Gutowska 2002; Jeong et al. 2002; Peppas et al. 2000; Ju et al. 2001; Ulijn et al. 2007a).

Hydrogels are defined as cross-linked 3D gels that have water (> 90%) as their dispersion

medium and therefore swell but not dissolve in aqueous solvents (Wang et al. 1993), and

are known to be useful biocompatible biomaterials for a wide range of biomedical

applications such as drug delivery, inflammation, tissue engineering (cartilage

replacement), skin grafts, bioseparation and wound dressings (Kim and Park 1998;

Hoffman 2002). In reference to wound healing, hydrogels are essentially important as

they possess ‘ideal’ properties of a dressing (Morgan 2002) such as: non-adherent

(therefore reduce pain when removed), maintain a moist environment, rehydrate the


70

wounds, maintain a cool and soothing effect, enable autolytic debridement (Hess 2002;

Baranoski and Ayello 2004) and able to absorb wound exudates.

2.6.3.1 Passive / Non-responsive hydrogel dressings

Table 18 (appendix II) summarises the various types of ‘passive’ hydrogel dressings that

are used to treat various chronic wounds. The table depicts the manufacturers that

developed the dressings, the management of action and finally the indications and

contraindications for each type of dressing. All of the hydrogel dressings are non-

responsive because they are designed to create a physical barrier to cover the wounds. In

doing so these hydrogel dressings enable the exchange of fluids and gases, enable

autolytic debridement, soak up fluid from the wound (i.e. hydrate the wound bed) and

essentially provide a moist healing environment.

Some of the dressings mentioned in table 18 (appendix II) are used as a primary dressing

(e.g. Aquasorb, entry 6; CarrGauze pads and strips with Acemannan, entry 9;

Dermagram hydrophilic and zinc saline dressings, entries 17-18; DermaSyn and

DermaGauze, entry 19; MPM gel pad, entry 30) and on the other hand, some dressings

have to be secured by a second dressing (e.g. Aquasite hydrogels: amorphous,

impregnated gauze, impregnated non-woven and sheet, entry 5). Two dressings are said

to be acidic (e.g. Bioloex wound gel, entry 7 and Phyto Derma wound gel, entry 35) to

promote healing. In addition, two dressings are formulated as a combination of

hydrogel/alginate i.e. NU-GEL (entry 33) and Purilon (entry 36) (Morgan 2002).

The advantage of incorporating alginate into hydrogel wound dressings such as Purilon

gel and NU-GEL enables both dressings to soak up significant amounts of fluids ~ 15 to

20 times more compared to their own molecular weight (Jones et al. 2006). The

absorption of NU-GEL is more than Purilon gel (23% and 18% respectively) whereas

the opposite effect is seen for the ability to donate moisture, Purilon donates more than

NU-GEL i.e. 11% and 3%, respectively (Jones and Vaughan 2005). Additionally, the

NU-GEL dressing can function under acidic conditions since the alginate component of

the dressing is modified with propylene glycol (Augst et al. 2006).


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In ‘The Global Market for Advanced Wound Care Products 2008’ report, IntraSite and

Purilon gels were recorded as the leading hydrogels used in the wound market. Intrasite

gel (Smith & Nephew, entry 28: table 18 – appendix II) is composed of

carboxymethylcellulose (CMC) polymer (2.3%), propylene glycol (20%) and water

(77.7%) whereas Purilon gel (Coloplast Corporation, entry 36: table 18 – appendix II) is

composed of CMC, calcium alginate and water. Their mechanistic actions are very

similar for both dressings as they consist of the hydrated CMC (cross-linked) material

which has the ability to absorb and donate fluids and subsequently facilitate the

rehydration needs that enable autolytic debridement of necrotic tissue (Thomas and Hay

1994) on top of maintaining a moist environment to improve wound healing. Both

hydrogels have very similar absorption properties i.e. 14% for IntraSite gel and 18% for

Purilon gel (Jones and Vaughan 2005). The moist environment is further increased as

both IntraSite and Purilon gels have a high water content which not only ensures

efficient exchange of water and oxygen molecules but also donates water molecules to

the wound surface (Jones and Milton 2000). Purilon gel is able to donate 11% moisture

compared to IntraSite gel which only donates 6% moisture (Jones and Vaughan 2005).

For both hydrogels the management of action is indicated for pressure ulcers, moreover

Purilon can be use to treat leg ulcers, whereas Intrasite can be used to treat tunnelling

wounds and partial/full-thickness wounds (as indicated in appendix II).

2.6.3.2 Active/ Responsive hydrogel dressings

In recent years, researchers have focused on designing more effective and economical

wound dressings. The current research is based on designing and developing responsive

(active) wound dressings that stimulate the microenvironment to activate chemical

reactions and biochemical processes within wounds to promote wound healing. It is

highly beneficial that such dressings also employed antimicrobial properties to remove

pathogens by transporting antibiotics, silver (Ag+) or iodine to inhibit infection; over the

course of time provide information for the wellbeing of the wound and essentially to

activate wound healing (Stroock and Cabodi 2006). The aim is moving towards

producing natural or synthetic dressings that will tackle more than one biochemical

problem e.g. managing elevated levels of proteases and free radicals; controlling the

release of therapeutic factors into non-healing wounds to restore the lack of protease

inhibitors, growth factors, various ECM proteins, cells and oxygen levels; and


72

transforming the pH of chronic wounds to normal levels. In doing this, the attempt is to

produce cost-effective wound products that will speed-up the healing of deadlocked

chronic wounds with various physiological problems.

This section briefly gives some examples of the recent advances for active dressing

whereas the next section extensively reviews protease-modulating dressings that are

primarily marketed/ designed or being investigated for the management of elevated

proteases found in chronic wounds.

Recently, Stroock et al. (2006) reviewed a new active hydrogel dressing to treat chronic

wounds. The design of the dressing is based on the concept of vacuum-assisted closure

(VAC®, Kinectic Concepts Inc.) that was developed by Morykwas (Morykwas et al.

1993). The mechanistic action of VAC® is based on a foam (made from polyurethane or

poly(vinyl alcohol) (PVA)) that is placed within a wound, protected with a plastic sheet

and then applying a vacuum (pulled down to ~ 635 torr) to the surface volume of the

wound in accordance to the sponge in order to withdraw wound fluids from wounds.

Under the plastic, compression of the foam delivers active mechanical stress to the

tissues and the wound fluids from tissue are withdraw and transported to the outlet tube.

It is suggested that this approach enables wound healing but the healing mechanism is

unclear. Stroock and colleagues state they intent to use the VAC approach to produce a

microporous poly(hydroxyethyl methacrylate) (pHEMA) sponge with pores of fixed

micro- and macro-geometry (i.e. pores ~20 µm in a 500 µm thick layer). The hydrogel

wound dressing is completed by covalently bonding the sponge to a silicone backing

(termed as PDMS) by acrylate-terminated silanes in doing so this produces strong fluidic

connections to the sponge. By using a model wound bed they have shown to achieve a

consistency of mass transfer of fluid (Stroock and Cabodi 2006). The main disadvantage

of this approach is that the wound dressing is designed to remove everything from the

wounds, and this may not aid in wound healing.

Insense Limited (UK) have marketed a moist oxygenating hydrogel dressing, OxyzymeTM

designed to tackle hypoxia (see section 2.4.1 above) in which the imbalance of oxygen

levels are restored to normal within chronic wounds such as: VLUs, DFUs, pressure and

arterial ulcers including surgical and radiation wounds (Ivins et al. 2007; Queen et al.

2007; Eaton 2008).


73

Figure 14. Mode of action for OxyzymeTM hydrogel dressing. The imbalance of oxygen within chronic wounds is restored by the application of OxyzymeTM which is an active double dressing utilising oxidative biological mechanisms to catalyses the conversion of the atmosphere oxygen (insoluble) into dissolved oxygen (green). Dissolved oxygen prevents hypoxia and oxygen levels are restored to normal. Figure adapted from Davis 2007.

This enzyme-activated hydrogel dressing is made from polysulphonate synthetic polymer

and > 65% water (Eaton 2008); and constructed together by two hydrogel sheets: (1) a

wound contact hydrogel which contains glucose incorporated with iodide ions (< 0.04%

w/w) and has to be applied directly over the wound; (2) a secondary hydrogel sheet with

immobilised glucose oxidase is placed over the wound contact hydrogel (Insense

Limited; Ivins et al. 2007; Davis 2007; Davis et al. 2009). Merging of the two hydrogel

sheets in the correct order activates the function of OxyzymeTM to mimic the

biochemical ‘respiratory burst’ process observed in leukocytes (Ivins et al. 2007) and the

mechanism of OxyzymeTM action is summarised by figure 14. Glucose and atmospheric

oxygen bind to glucose oxidase which in turn catalyses the oxidation of glucose to

gluconic acid and hydrogen peroxide (H2O2). H2O2 has two roles: firstly it kills anaerobic

bacteria and secondly at the wound edge it acts as an oxygen shuttle in which

endogenous H2O2 (in situ) remaining in OxyzymeTM behaves as a soluble water carrier

(Thorn et al. 2005) that diffuses through OxyzymeTM to react with the iodide ions to

actively release dissolved oxygen (Ivins et al. 2007; Davis et al. 2009) and iodine (Davis

2007) into the wound environment. In doing so, the oxygen levels within chronic

wounds are restored preventing hypoxia; additionally the dissolved oxygen has the ability

Oxygen

11 22

Secondary dressing:

Immobilised glucose

oxidase hydrogel

Primary dressing: Glucose

and iodide containing

wound contact hydrogel

Exuding wound

Oxygen

Glucose

Glucose

Oxidase

Hydrogen

peroxide

Iodide ions

Oxygen Iodine

33

44

3. Binding of oxygen to glucose oxidase catalyses the oxidation of

glucose to gluconic acid (not shown) and hydrogen peroxide

(purple) within the dressing.

4. Hydrogen peroxide binds with the immobilised iodide ions within

the dressing to generate dissolved oxygen and iodine. Oxygen is

delivered to the wound (green arrows) and the iodine combats

bacterial infection at the wound surface.

1. Combining of the hydrogels

dressings enables glucose (orange)

from the bottom dressing to diffuse

into the top enzyme dressing and

this activates glucose oxidase (blue)

in the top dressing.

2. Atmospheric oxygen (green)

migrates into the dressing.


74

of killing anaerobic bacteria via respiratory burst which is enhanced by iodine as it

behaves as a potent antimicrobial agent that produces a hostile environment for

microbial growth.

Man-made hydrogel release vehicles (RV) have been designed to control the release of

growth factors for wound applications to promote tissue formation within chronic

wounds. Currently, the non-sterile Regranex® hydrogel (Johnson & Johnson) made of

becaplermin (0.01% i.e. 100 µg per 1g), sodium CMC, sodium chloride, sodium acetate

trihydrate, glacial acetic acid, water for injection, including preservatives (i.e.

methylparaben, propylparaben, and m-cresol) and a stabiliser (L-lysine hydrochloride).

From this ingredient, Becapleman is the active component within Regranex® and it is a

mimic of the human PDGF-β,β. It is formulated as a human recombinant PDGF-β,β

generated by recombinant DNA technology in yeast in the form of a homodimer

(containing two identical polypeptide chains connected together by disulphide bridges)

with a molecular weight of 25 kDa. The function of Regranex® is to correct the

imbalance of endogenous growth factors within the wound bed of ulcers by delivering

Becapleman i.e. PDGF-β,β to facilitate chemotactic behaviour by recruiting and

proliferating cells to smooth the progress of granulation tissue formation. However, it is

suggested that such a dressing is inadequate in its treatment due to a prolonged

inflammatory phase (Cullen et al. 2002b). To combat this problem, over the years other

hydrogel RVs have been fabricated to release growth factors for wound healing

applications. Bourke et al. demonstrated a continual release of the PDGF-β,β up to 3-4

days from a UV photo-crosslinked PVA hydrogel with hydrophilic PVA fillers. In this

manner these hydrogels are considered to be more beneficial than contemporary wound

applications as they do not have to be changed frequently (Bourke et al. 2003).

Recently, Bader et al. showed the diffusive release of keratinoctye growth factor (KGF)

and glucose from semi-interpenetrating network (sIPN) hydrogels. These biodegradable

sIPNs were fabricated as a binary hydrogel to mimic the ECM by covalently photo-

crosslinking poly(ethylene glycol) diacrylate with unmodified gelatin. KGF was

successfully released from the sIPNs hydrogels via unidirectional desorption and had a

diffusion coefficient of 4.86 × 10−9 ± 1.86 × 10−12 cm2/s. On the other hand glucose

was unable to be released from the sIPNs, but the transport of glucose was achieved by a

flow mechanism through sIPN hydrogels with a diffusion coefficient of 2.25


75

× 10−6 ± 1.98 × 10−7 cm2/s (Bader et al. 2009). Bader et al. speculate that these sIPNs

are efficient hydrogels for releasing drugs, growth factors and nutrients to proliferating

cells during wound healing.

2.6.3.3 Protease-modulating hydrogel dressings

The failure of Regranex® to work effectively is mainly the result of elevated levels of

proteases present in chronic wound fluids (Cullen et al. 2002b) that cause tremendous

problems to various biochemical pathways within chronic wounds. Although the

pathophysiology of chronic wounds is still not clearly understood, there has been a

gradual increase of wound dressings that enter the wound care market to address and

overcome the biochemical problem of elevated proteases commonly observed in chronic

wounds. The wound dressings that adjust protease activity within chronic wounds

commonly fall into the category of ‘protease-modulating’ dressings or are sometimes

referred to as active dressings. According to the British National Formulary (BNF,

2009), currently there are six protease-modulating dressings available within the UK

wound care market: PromogranTM and Promogran PrismaTM (Johnson & Johnson),

Cadesorb (Smith & Nephew), Suprasorb® C (Activa Healthcare), Sorbion® S (H&R

Healthcare) and Tegaderm® Matrix with PHI technology (3MTM Healthcare Limited).

Additionally, the pro-ionic hydrogels marketed by First Water Limited have recently

become available to inhibit proteases activity (First Water Limited). Nevertheless,

scientific research continues to develop more protease-modulating wound dressings as

there has been a lot of interest to provide a variety of cost-effective protease-modulating

wound care products around the world. Table 8 summarises the protease-modulating

hydrogels that are currently marketed in the wound care market (primarily within UK/

Ireland) and being investigated by various scientists to control the imbalances of

proteases found in chronic wounds.

In 2002, the first enzyme responsive dressing to be marketed was PromogranTM (table 8,

entry 1) which is a sterile, 3 mm thick hexagonal sheet consisting of a freeze-dried

homogeneous matrix composed of bovine collagen (55%) and oxidised regenerated

cellulose (ORC, 45%). After being applied to the wound, when the freeze-dried matrix

comes in contact to the wound exudate, its initial mode of action is to absorb the wound


76

fluid (assisted by the highly absorbent ORC composition) and upon saturation the matrix

transforms into a soft biodegradable (bio-absorbable) conformable gel.

Table 8. Application of protease-modulating hydrogel dressings for chronic wounds

Entry Name Hydrogel Ingredients Type of response

Enzymes References

1 PromogranTM ORC/ collagen Inhibit enzyme activity

MMPs Elastase Plasmin

J&J Medical; Cullen et al. 2002a, 2002b; GMAWCP 2008; BNF 2009

2 S-SEBS polymer

S-SEBS

Na+, Ag+, doxycycline

Reduce enzyme activity

Neutrophil elastase, MMP-8

Vachon and Yager 2006

3 Promogran PrismaTM

ORC/collagen and silver

Inhibit MMP activity & bacterial infection


J&J Medical; BNF 2009

4 Sorbion® S Polyacrylate polymers in cellulose matrix

Modulates MMPs

MMPs Sorbion AG; H&R Healthcare; BNF 2009

5 TenderWet

Polyacrylate superabsorbers preswollen in Ringers’ solution

Reduce enzyme activity

MMPs Eming et al. 2008

6 Pro-ionic hydrogels/ matrix

Sulphonate groups, NaSPA/NaAMPS

Inhibit enzyme activity

MMP-2 MMP-9

FW Ltd, Munro and Boote 2009

7 Cadesorb Starch beads in PEG/PPG

Lowering pH to inhibit MMP activity


S&N Wound Management, GMAWCP 2008; BNF 2009

8 Tegaderm™ Matrix

Cellulose acetate matrix, impregnated with PHI in PEG


MMP 3M; BNF 2009; GMAWCP 2008

9 Bisphosphonate-functionalised hydrogels

Bisphosphonate (alendronate) tethered to pHEMA/PEG


MMPs Rayment et al. 2008

10 Modified gelatin microspheres (DHB-MS)

MMP inhibitor (DHBA) conjugated to gelatine MS and impregnated into collagen scaffold

Inhibit MMP activity and control bacterial infection

MMP-2 MMP-9

Adhirajan et al. 2009

Abbreviations: ORC (oxidised regenerated cellulose), MMP (matrix metalloproteinases), J&J (Johnson & Johnson), BNF (British National Formulary), S-SEBS (Sulfonated styrene– ethylene– butylenes–styrene), Na (sodium), Ag (silver), NaSPA (acrylic acid (3-sulphopropyl) ester sodium salt), NaAMPS (sodium salt of 2-acrylamido-2-methylpropanesulphonic acid), FW Ltd (First Water Limited), Ltd (Limited), PEG (polyethylene glycol), PPG (polypropylene glycol), pH (potential hydrogen), S&N (Smith and Nephew),PHI (polyhydrated ionogens), 3M (3MTM

Healthcare Limited), GMAWCP (The Global Market For Advanced Wound Care Products 2008), pHEMA (poly-hydroxyethylmethacrylate), DHB-MS (modified gelatin microspheres), DHBA (2,3-dihydroxybenzoic acid) and MS (microspheres).


77

Mechanistically, PromogranTM binds and inactivates MMP activity in addition to binding

endogenous growth factors and protecting them from being degraded by the

overabundant proteases (e.g. MMPs) found in wound exudates (Cullen et al. 2002b). The

function of bovine collagen is to behave as an alternative profuse substrate for MMPs.

In addition to its highly absorbent ability, ORC enhances the inhibition of these

proteases by demolishing the 3D structure of MMPs. This is accomplished by the

negative charges of ORC which bind and dislocate the positively charged divalent metal

ions e.g. Zn2+ within the catalytic site of the MMP structure. Removal of the divalent

metal ions inhibits the functional enzymatic activity of MMPs and as a result the MMPs

are incapable of binding to any substrate (Ovington 2007) within wounds. Given that

PromogranTM gel also binds growth factors (e.g. PDGF) in a non-covalent manner, as the

gel degrades over time the growth factors are discharged back into the wound in their

functional form (Johnson & Johnson Gateway). Furthermore, PromogranTM has the

ability of removing oxygen free radicals which are generated by lipid degradation of the

cell membrane (Johnson & Johnson Gateway).

The inhibition mechanism of PromogranTM for chronic wounds proteases was further

studied in comparison to wet gauze. Cullen et al. demonstrated the ability of

PromogranTM to significantly reduce the activity of neutrophil-derived elastase, plasmin

and MMPs activities in wound fluids collected from patients with diabetic foot ulcers

(DFUs) which was not observed with wet gauze (Cullen et al. 2002a). The highest rate

of reduction in protease activity was observed for MMP which reduced by 94% (just

within 30 minutes), plasmin reduced by 67% (within 1 hour) and the slowest rate was for

elastase which reduced by 86% (after 2 hours). Coinciding with these results, another

study showed a reduction of elastase, plasmin and gelatinase activity in wound exudates

collected from patients with chronic VLUs after treatment with OCR/collagen matrix

compared to control patients who were treated with a hydrocholloid dressing (Smeets et

al. 2008). While at the same time as reporting gelatinase activity is reduced, on the

contrary and unexpectedly this study reveals no reduction of MMP-2 activity in wound

exudates of chronic VLUs after treatment with OCR/collagen matrix (Smeets et al.

2008).

However, Vachon and Yager reported that the mode of action of their ion exchange S-

SEBS (sulfonated styrene – ethylene – butylenes – styrene) hydrogel dressing (table 8,


78

entry 2) was more efficient in binding and inactivating MMP-8 and elastase compared to

PromogranTM. The study established that the collagen matrix of PromogranTM was

susceptible to degradation by NE. The S-SEBS (sulfonated-triblock) polymer consisted

of a sulfonated-hydrocarbon backbone that was layered onto polyester fabric (behaves as

a support) and subsequently modified with various ion-exchange formulations (e.g. Na+,

Ag+ and doxycycline etc). The ion-exchange mechanism of the S-SEBS hydrogel

dressing enable the binding and inactivation of proteases and this was further facilitated

by releasing therapeutic antibiotics e.g. doxycycline into chronic wounds (Vachon and

Yager 2006). The release of doxycycline from the S-SEBS-doxycycline hydrogel

formulation was also found to control the bacterial growth of various pathogens (P.

aeruginosa, S. aureus, E.coli and Enterococcus facecalis) present in wounds for a limited

time of 48 hours (Vachon and Yager 2006).

In view of that, Johnson & Johnson developed a similar sterile, freeze-dried matrix to

that of PromogranTM known as Promogran PrismaTM (table 8, entry 3) consisting of

bovine collagen (55%), oxidised regenerated cellulose (ORC, 44%) and 1% silver-ORC.

Promogran PrismaTM differs from PromogranTM as it contains silver which is ionically

bound to ORC (25% w/w). The application of Promogran PrismaTM to a wound is the

same as Promogran. Even though Promogran PrismaTM has all the benefits as

PromogranTM it is more advanced as it has a dual approach where it not only can inhibit

protease activity but it also functions as an efficient antibacterial barrier as it kills bacteria

(such as P. aeruginosa, S. aureus, E. coli and Streptococcus pyogenes) in the dressing to

decrease the bacterial growth of these pathogens that are normally found within wounds.

By delivering silver to the wound it has the ability to facilitate healthy granulation tissue

formation and epithelialisation without bacterial infection (Johnson & Johnson Medical).

Collagen is increasingly considered as an ideal biomaterial for developing wound

dressings and gradually there has been a rise in developing collagen based dressings.

Elsewhere brief descriptions of the types of collagen dressings marketed for wound care

have been reviewed (see GMAWCP 2008) and will not be reviewed here. In spite of

that, the ideal properties of collagen dressings is that they can directly control elevated

proteases in chronic wounds since the abundant collagen within the matrix of these

dressings bio-mimics and substitutes as an important substrate for chronic wound

proteases as we have seen previously with Promogran. In a recent study, Schönfelder et


79

al. (2005) showed that Suprasorb® C was able to reduce the elastase activity as the

bovine collagen type I matrix of the Suprasorb® C (a collagen foam product marketed

by Activa Healthcare) behaved as a surrogate substrate for PMN elastase. In addition,

when the collagen matrix within collagen dressings breaks down, it’s broken down

fragments behave as chemotactic to recruit various cells to facilitate cell migration and

control bacterial infection to restore normal tissue repair during wound healing process.

Some authors have criticised collagen or collagen/ ORC dressings to have built-in risk

associated with the collagen composition of the dressing as it is made of animal origin

such as bovine (Eming et al. 2008) which amongst different species can be immunogenic

or problems can arise during conformational changes through processing (Leonard et al.

1998 as referenced in Uchegbu and Schätzlein 2006). Adding to this problem, we have

previously encountered that collagen matrix within wound dressings is susceptible to

degradation via wound proteases such as NE (Vachon and Yager 2006). For that reason,

there is a demand of advancing the wound management products consisting of

biocompatible synthetic origin (Eming et al. 2008) in conjunction with moist healing

properties and antimicrobial agents.

Hydroactive ionic wound dressings containing synthetic superabsorbers composed of

poylacrylate (table 8) are increasingly reported as a novel way of inhibiting the protease

activity normally elevated in chronic wounds. Polyacrylate superabsorbers are gel-

binding highly absorbent polymers compiled from polymerised acrylic acid with different

amounts of crosslinking and this enables polyacrylate dressings to absorb various

amounts of water compared to their own dry weight, hence the name polyacrylate

superabsorbers. The binding of water within these superabsorbing hydrogels is achieved

by electrostatic forces of the high density of ionic charges within polyacrylates (Eming et

al. 2008). Polyacrylates are highly used in diapers, feminine hygiene products and wound

dressings (Eming et al. 2008). Currently, Sorbion® S and TenderWet are two examples

of polyacrylate superabsorbing hydrogels marketed as wound dressings.

Sorbion® S (table 8, entry 4) marketed by Sorbion AG and distributed by H&R

Healthcare Limited (UK) as a sachet format (also known as Sorbion® Sachet S) consists

of polyacrylate superabsorbers in a cellulose matrix. Active control of the wound

environment is achieved by Hydration Response Technology (HRT) and this is achieved


80

by the propinquity of the polyacrylate superabsorbers with the cellulose fibres. The

striking mechanism of this dressing is that it is able to mop-up large amount of exudates

(~ 700 mL) from the wound surface/ bed via osmostic strength from the underlying

tissues. Once the fluids alongside bacteria are absorbed they become interlocked within

the super-absorbers and cellulose fibres (Sorbion AG: www.sorbion.com). Under the

influence of HRT, besides its absorbent property, according to the manufacturer

Sorbion® S has many advantages: (1) the debridement is very soft as the dressing has the

ability to remove slough and toxins thereby reducing odour within the wound; (2)

prevent infection by managing bioburden of pathogens, since the bacteria becomes

locked therefore risk of cross contamination is reduced; (3) it is cost effective as the

number of changes or visit from the nurse are reduced therefore saving on material and

duration of time spend in hospital; and (4) the dressing possess anti-inflammatory

properties. As a consequence the damage at the wound edge is prevented and healed,

high levels of exudates are removed and finally it has the ability to modulate MMPs

(Sorbion AG: www.sorbion.com). A recent case study reviewed the effectiveness of

using Sorbion® S as a primary dressing to treat a highly exuding DFU which only took

12 weeks to heal after its application and prior to that it would not heal (Chadwick 2008).

Even though the manufacturers fail to comment on the mechanism of how Sorbion® S

modulates MMPs; current investigations of polyacrylate superasbsorbers are shedding

light on determining or understanding the possible mechanistic binding or inhibition of

inflammatory proteases such as elastase and MMPs including the inhibition of ROS or

reactive nitrogen species (RNS).

Eming et al. (2008) demonstrated the first in vivo binding of polyacrylate superabsorber

particles: TenderWet (in Germany/ Hydroclean in France; manufactured by Hartmann)

to MMPs within CWFs from patients suffering from chronic VLUs (table 8, entry 5).

The study showed that MMP-2/-9 activity was inhibited by > 87% within CWFs.

Despite the fact that Eming et al. showed direct binding of MMPs with TenderWet

particles, the mechanism of inhibition was accomplished in an indirect manner as the

concentration of the divalent ions i.e. Ca2+ and Zn2+ within the buffer of an MMP assay

were found to decrease in the presence of TenderWet particles. This allowed the authors

to indirectly conclude that Ca2+ and Zn2+ ions within the catalytic active site of MMPs

competitively bind to polyacrylate superabsorber particles (TenderWet) and when bound


81

the catalytic Ca2+ and Zn2+ ions are no longer available for MMP activity and accordingly

inhibits the functional enzymatic activity of MMPs (Eming et al. 2008) and therefore are

incapable of binding to any substrate within wounds.

Recently, at the EMWA conference, Abel et al. (2009a) showed that polyacrylate

superabsorbers (e.g. Vilwasorb®, Zetuvit® and Sorbion® sachet) have a high affinity to

bind elastase and MMP-2 in vitro and demonstrated that protease inhibition was

significant after 24 hours incubation. Elastase was found to reduce by 90% (p < 0.001)

whereas MMP-2 reduced by ~ 85% (p < 0.01). Additionally, Abel et al. (2009b) used a

chemiluminescent method to show that polyacrylate superabsorbers i.e. Vliwasorb®

(manufacturer by Lobman & Rauscher) can inhibit free radical formation of oxygen

species (ROS i.e. superoxide from 55% to 90%) and nitrogen species (RNS i.e.

peroxynitrite anion from 70% to 85%).

According to First Water Limited (FW Ltd) even though polyacrylate absorbers are ionic

dressings they are poor at imitating natural body molecules because they utilise

carboxylate chemistry. Whereas the hydrophilic co-polymer formulated within their

newly invented biomimetic pro-ionic hydrogels (table 8, entry 6) is more superior as it

contains multiple synthetic pendant anionic sulphonate groups (e.g. –SO2– or –SO2–OH)

making these hydrogel dressings ideal at mimicking the sulphated glycosaminoglycans

(GAGs, e.g. heparin) that are naturally located on the surface of cell membranes and the

ECM (Munro and Boote 2009). In acute wound healing, GAGs are binding sites that aid

in the regulation of biochemical processes such as influencing proteases activity and

normalising inflammation, whereas in chronic wounds the GAGs are dysfunctional

(Munro and Boote 2009).

In their patent, Munro and Boote state that the pro-ionic hydrogels are composed of two

monomers and both monomers are made from olefinically unsaturated sulphonic acid

monomer/salt. Each monomer is considered to be different, in the sense that the first

monomer consist of acrylic acid ester suphonic acid monomer/salt e.g. acrylic acid (3-

sulphopropyl) ester sodium salt (NaSPA) and the second monomer consist of acrylamide

sulphonic acid monomer/salt e.g. sodium salt of 2-acrylamido-2methylpropanesulphonic

acid (NaAMPS). In this way, the composition of the pro-ionic hydrogel comprises a co-


82

polymer of NaSPA/NaAMPS with pendant sulphonyl groups counterbalanced by one/

more cations (Munro and Boote 2009).

Alongside managing moisture levels in wounds, the partially-hydrated superabsorbent

ionic polymer of the pro-ionic hydrogels/matrix has the dual functionality of working as

a hydrating and absorbing dressing. The hydration role of these wound dressings has

been design to uniquely mimic the natural water activity of the skin and the water levels

are controlled so that there is a balance with the amount of moisture delivered to the

wound. According to the manufacturer, the hydration levels of pro-ionic matrix is

reported as 0.63 which closely resembles the stratum corneum (i.e. 0.65) of the epidermal

layer of the skin (FW Ltd). Just like the polyacrylates superabsorbers (i.e. Sorbion® S

and TenderWet), pro-ionic hydrogels have the capacity to absorb large amounts of

fluids/exudate (i.e. 2.5 – 50 times more) compared to its own molecular weight and

securely interlock it within the matrix of the hydrophilic wound dressing. Additionally,

ongoing clinical work is demonstrating that the pro-ionic hydrogels/ matrix dressings

have the ability to inhibit the enzyme activity of MMP-2/-9 (FW Ltd) to control the

biochemical environment without the need of releasing active substances into the wound

(GMWCP, 2008).

Many scientists are interested in designing wound dressings that will alter the biochemical

wound environment by addressing the potential hydrogen (pH) concentration within

chronic wounds. It is envisaged that controlling protease activity can be achieved by

altering the pH of chronic wounds to acidic pH (Schultz et al. 2005b; Greener et al.

2005) see section 2.4.3. Cadesorb (table 8, entry 7) manufacturer by Smith & Nephew

(Healthcare, Hull) is a pH-modulating ointment that decreases proteases activity within

chronic wounds. The matrix of the hydrogel ointment/ dressing is composed of

carboxylated cross-linked starch beads within a carrier consisting of polyethylene glycol

and polypropylene glycol (PEG/PPG).

Cadesorb has to be secured by a secondary dressing so it firmly remains in contact with

the wound. After its application, the starch beads neutralise the wound environment by

simultaneously substituting the wound with H+ ions and exchanging them with basic ions

that are elevated in chronic wounds (as summarised in figure 15). The substitution of H+

ions decreases the local pH of the chronic wound from approximately pH 7 – 8 (i.e.

neutral – slightly alkaline) to approximately pH 4 – 5 (i.e. acidic). The reduction of the


83

pH decreases/ inhibits proteases activity and this facilitates matrix formation of the

healing process without affecting the wound tissue. Epithelialisation starts when

keratinocytes migrate across the wound and this promotes proliferation of new epithelial

cells/ tissue which fill up the wound (figure 15). According to the manufacturer

Cadesorb has been shown to control the elevated proteases activity of elastase, plasmin

and MMPs in chronic wounds.

Figure 15. Mechanism of Cadesorb. At pH 7 – 8 (i.e. neutral – slightly alkaline) the provisional matrix of chronic wounds is destroyed by surplus levels of proteases (1); with the application of Cadesorb to the chronic wound (2); the pH of the wound reduces to pH 4 – 5 (acidic) by the substitution of H+ ions and as a result protease activity decreases which facilitates matrix formation and epithelisation commences with the migration of keratinocytes across the wound (3), this promotes proliferation and new epithelial cells/ tissue fill the wound (4), and the pH profile of protease activity of a chronic wound treated with/ without Cadesorb (5). Figure adapted from Smith & Nephew Wound Management. The recent advance invention by 3M™ Healthcare Limited (termed as 3M in this thesis)

makes use of citric acid buffer to reduce the pH of chronic wound, referred to as

TegadermTM Matrix dressing with polyhydrated ionogens (PHI™) technology (table 8,

entry 8). This advanced wound care solution is made of cellulose acetate that is

impregnated with PHI ointment, water (20%) within the hydrophilic matrix of PEG and

buffered with citric acid. The mechanism of TegadermTM Matrix dressing with PHITM is

to regulate the micro-environment of a wound, by controlling the imbalance of MMP

activity in order to speed-up the smooth progress of re-epithelialisation to facilitate

wound closure of chronic wounds. The cellulose acetate is a non-biodegradable highly


84

versatile carrier which after the dressing is applied enables the access of wound exudate

into the dressing.

According to 3M, the PHITM technology is the striking feature of this dressing, as it is a

unique ointment that contains a combination of four metallic cations i.e. rubidium,

calcium, zinc and potassium that are present within the body (e.g. wound exudate and in

serum). The PHITM technology promotes natural moisture to modulate the production

of MMPs by down-regulating the protease activity (e.g. MMP-2) within fibroblast cells.

In the presence of wound exudate and under the influence of body heat and moisture,

the PEG moiety enables the accurate activation for the steady liberation of the metal ions

into the wound bed followed by their penetration into fibroblast cells (3M). The first ion

to penetrate the fibroblasts cells is rubidium and its function is to make a channel

opening within the cell membrane to allow the remaining ions of the PHITM technology

to penetrate the cells. The production of MMPs is inhibited by both calcium and zinc

ions. The cell membranes of fibroblasts are depolarized by potassium ion which further

facilitates the inhibition of MMP production at the transcriptional level by switching off

protein synthesis of MMPs (3M). Additionally, potassium supports tissue regeneration.

Citric acid buffers the wound environment by decreasing the local pH of chronic wounds

from neutral/ alkaline to slightly acidic (i.e. pH 5). This reduction of the pH not only

fosters the transport of metal ions over the cell membrane it also increases both the

synthesis of neutral complexes along with cell proliferation to accelerate the normal

wound healing cascade of deadlocked chronic wounds (3M).

TegadermTM Matrix dressing with PHITM is contraindicated for wounds that cannot be

treated with any other wound care product after the duration of 4 – 6 weeks. It can be

used to treat non-infected chronic wounds such as diabetic/ venous ulcers and stages II-

IV pressure ulcers (3M). The manufacturers have carried out some case studies to

determine the efficacy of this dressing; according to 3M TagadermTM Matrix can heal

chronic wounds within the range of 4 – 10 weeks depending on the wound.

It is worth nothing that Epi-Max (in USA), DerMax and MelMax (in EU and Middle

East) are all precursor wound dressings of TagadermTM (in UK) and are all composed of

an acetate carrier impregnated with PHIs ointment (Greystone Pharmaceuticals, Inc.).

Originally, the PHITM technology was extracted from the Red Oak Bark tree but it is now


85

formulated and synthesised by Greystone Pharmaceuticals, Inc. (sometimes referred to as

PHI formulation, PHI-5 or metal ionogen (MI) technology). It was patented by their

subsidiary group Dermagenics Inc. (Patent Dermagenics Europe B.V., as cited in van

Rossum et al. 2007). For wound care products, in 2007 Greystone Pharmaceuticals put

in force a worldwide licence agreement with 3M for use of Greystone’s patented PHI

technology (Greystone Pharmaceuticals, Inc.); additionally it gave permission for 3M to

distribute TagadermTM within the USA.

Furthermore, various in vitro studies have demonstrated that DerMax has the ability to

inhibit MMP activity such as MMP-2 (van den Berg et al. 2003; Karim et al. 2006)

alongside inhibiting the production of ROS (van den Berg et al. 2003; Pirayesh et al.

2007) and inhibit complement activation (van den Berg et al. 2003). Likewise, at the

molecular level, DNA microarrays studies have shown that the gene expression of

diabetic fibroblasts cells is increased and resembles normal fibroblasts after treatment

with PHITM technology since MMP activity (MMP-9) is found to decrease with increasing

concentration of PHITM technology (Schultz 2008 as cited in 3M PHITM technology

technical brochure). Additionally, Pirayesh et al. carried out an in vivo multi-centre pilot

study on a total of 20 patients with DFUs and demonstrated the effectiveness of the

PHITM technology using DerMax. A total wound closure of the DFUs was observed

within 80% of the patients (16 patients) which was maintained within 6 to 38 weeks with

a total mean time of 18 weeks (Pirayest et al. 2007).

Increasingly a choice of studies have reported the use of synthetic inhibitors as a

beneficial approach of inhibiting MMP activity and a way of promoting forward the

healing of chronic wounds, since the main drawback of previous dressings specifically

the collagen/ ORC dressings is the total inactivation of the proteases within the wound

bed given that the MMPs are required by other processes such as activating growth

factors and regulating the immune system during various phases of the wound healing

process other than just ECM degradation. To overcome this problem, consequently

Rayment et al. (2008) proposed the functionalisation of pHEMA/PEG hydrogels with

the MMP-inhibitor, alendronate (figure 16) in the form of a sodium salt (figures 17) to

develop bisphosphonate-functionalised hydrogels (table 8, entry 9, figure 16) as a novel

way of neutralising the wound environment by inhibiting the activity of MMPs in CWFs

whilst keeping the protease activity active next to the upper cellular layer of a wound bed.


86

P

PNH

2

OH

OH

O

O

OH

OH

OH

P

P

R2 R1

OH

O

O

OH

OH

OH

AlendronateBisphosphonate (BP)

P-C-P bond has a

strong affinity for

divalent calcium ions

Calcium ions of MMP are

chelated more effectively when

R1 group of BP contains an -OH

The number of nitrogen

groups within the R2 group

concludes the potency of BPs

Figure 16. Chemical structure of the MMP-inhibitor, bisphosphonate. The general structure of BP demonstrates the functional groups that display a strong affinity for the divalent cations e.g. calcium ions within the MMP structure (left) and the structure of the germinal BP, alendronate (right). Figure adapted from Santini et al. 2003 and Heymann et al. 2004.

Alendronate is a small, aliphatic synthetic amino/nitrogen-containing bisphosphonate

(BP), distinctively a germinal BP as it bears two C – P bonds on the same carbon atom

i.e. P–C–P (Fleisch 2002; Rayment et al. 2008) see figure 16 (highlighted in red). This

bond alongside the hydroxyl (OH) group at the R1 side chain of alendronate has a strong

affinity for binding divalent ions (e.g. Ca2+ or Zn2+). Seeing as the aliphatic R2 group of

alendronate contains a single amino group (–NH2) it is said to be highly potent (Santini et

al. 2003; Heyman et al. 2004; Rayment et al. 2008).

The functionalised-bisphosphonate-pHEMA/PEG hydrogels were synthesised via a

Schotten-Baumann reaction which is a nucleophillic acylation reaction that occurs in the

presence of dilute alkali as summarised by figure 17 (Rayment et al. 2008). Since they

efficiently uncovered the inhibition of MMPs in CWF by the unsaturated alendronate

analogue, alendronate-methacrylate was then tethered to pHEMA/PEG hydrogels via

gamma induced polymerisation, generating alendronate-methacrylate-pHEMA/PEG

hydrogels (termed as functionalised-bisphosphonate-pHEMA/PEG hydrogels in this

thesis). The study reported that these hydrogels were able to render the catalytic activity

of MMP inactive as the degradation of collagen type I in CWF was found to decrease

significantly compared to an untreated control (p < 0.01).

It was proposed that once the MMPs are absorbed from the CWFs into the hydrogel

dressing then the mechanism of MMP inactivation is a result of divalent cation (e.g. Zn2+

or Ca2+) chelation (Rayment et al. 2008). In the literature this is achieved by the P–C–P

bond which has a strong affinity for calcium ions and the chelation is further enhanced

by the hydroxyl R1 group of alendronate which has a strong affinity to chelate Ca2+ ions


87

(or Zn2+) of MMPs (figure 16) via a tridentate binding/ conformation (Santini et al. 2003;

Heyman et al. 2004).

Cl

O P

PNH

2

OH

OH

O

O

OH

ONa . H2O

OH

O

NH

P

P

OH

OH

O

O

OH

ONa . H2O

OH

O

NH

P

P

OH

OH

O

O

OH

ONa . H2O

OH

O

OOH

O

O

OH

ONa

OH

OH

O

O

O

O

O

NH

P

P

OH

OH

O

OOH

O

O

OH

OH

+

3 3NaOH

ice salt bath

Stir vigorously

20 h

Methacryloyl

Chloride

Alendronate

sodium saltAlendronate-methacrylate

+ HCl

+

Polymerisation

(gamma-irradiation)

Alendronate-methacrylate-pHEMA/PEG hydrogel

Alendronate-methacrylate

HEMA monomer

(A)

(B)

+ PEG

3

Figure 17. Bisphosphonate-functionalised hydrogels synthesised via Schotten-Baumann reaction. (A) In a single step reaction the lone pair of the amine group of alendronate sodium salt attacks the carbonyl carbon of the vinyl-containing acid chloride, methacryloyl chloride generating a tetrahedral intermediate (not shown). The swinging back of the alkoxide electrons immediately generates an amide bond forming alendronate-methacrylate and HCl. (B) Co-polymerisation (gamma-induced) of alendronate-methacrylate with hydroxyethylmethacrylate (HEMA) monomer in the presence of PEG generates alendronate-methacrylate-pHEMA/PEG hydrogels. Figure adapted from Rayment et al. 2008.

Previous in vitro studies have shown that nitrogen-containing BPs have the ability to

impede cell growth of normal human epidermal keratinocytes (Reszka et al. 2001) and

the P–C–P containing BPs have the ability to inhibit cell function and possibly promote

apoptosis (Fleisch 2002). In view of that Rayment and colleagues also demonstrated that

the functionalised-bisphosphonate-pHEMA/PEG hydrogels were biocompatible when

exposed to a 3D ex vivo human skin equivalent model (Rayment et al. 2008) making them

promising hydrogels for wound care therapy.

Recently, Adhirajan and colleagues reported a similar approach of tethering MMP

siderophore inhibitors to gelatin (Type B from bovine skin) hydrogel microspheres (MS).

The gelatin MS were functionalised by covalently conjugating the COOH group of the

MMP inhibitor, 2,3-dihydroxybenzoic acid (DHBA, a synthetic catecholate type metal


88

chelator) to the NH2 groups of gelatin to develop modified gelatin MS (DHB-MS, 60–

120 µm, table 8, entry 10) with the intention of inhibiting the activity of MMP-2 and

MMP-9 within diabetic tissue/wounds (Adhirajan et al. 2009). Complete inhibition of

MMP-2 and MMP-9 activity within tissue lysate by DBH-MS was observed after 6 hours.

A wound dressing with dual ability of both inhibiting MMP activity combined with

controlling pathogen infection was developed in which Adhirajan et al. (2009) loaded

DHB-MS with deoxycycline and by fusion, a reconstituted collagen scaffold (DHB-MS

collagen scaffold) was constructed. The release of deoxycycline from the DHB-MS

collagen scaffold was slow compared to DBH-MS; but antibacterial activity was

continuously delivered by both DHB-MS and DHB-MS collagen scaffold to control the

infection of P. aeruginosa, S. aureus, K. pneumoniae and E. coli (Adhirajan et al. 2009).

2.7 SUMMARY

After the high demands from various clinicians/ scientists, the wound care market has

entered into a new era of developing advanced wound care products to control the

biochemical environment of elevated proteases found in chronic wounds. This is

because such abundant proteases (e.g. elastase, plasmin, thrombin and MMPs) are found

to be detrimental to ECM proteins or factors causing a prolonged inflammatory/

proliferation phases resulting in the development of deadlocked chronic wounds.

Functionalising synthetic hydrogels with a stimulus that is only responsive to one (or

more) of the elevated proteases enables the generation of highly exclusive/specific

bioresponsive wound dressings to manipulate the biochemical wound bed micro-

environment to facilitate and advocate wound healing of chronic wounds.

The moist and transparent ability of hydrogels make them ideal wound dressings. The

swelling and collapsing states exhibited by hydrogels make them an exceptional material

to interlock detrimental proteases within the hydrogel matrix upon removal of the

responsive stimulus. The use of hydrogel particles as opposed to sheets is an ideal

approach as some chronic wounds are deep open wounds, therefore the configuration

and dimension of these particles allows them to accommodate and fit all types of chronic

wounds without the necessity of cutting the hydrogel dressing to size. In contrast to the

protease-modulating wound dressings described within this chapter, this thesis presents


89

an alternative way of selectively ‘mopping-up’ excess elastase from sample fluids (which

mimic CWFs) into hydrogel particles in the attempt of providing a potential way of

down-regulating excess elastase in deadlocked chronic wounds.

CHAPTER 3: Functionalisation of PEGA Particles

90

CCHHAAPPTTEERR 33

FFuunnccttiioonnaall iissaattiioonn ooff PPEEGGAA PPaarrttiicclleess


91

3.1 INTRODUCTION

Within the last decade, there has been an increased interest in fabricating stimuli-

responsive materials to engineer smart polymers for biomedical applications which have

been reviewed extensively within this scientific field (Ulijn 2006; Ulijn et al. 2007a,

McDonald et al. 2009; Carpi and Smela 2009; Mohammed and Murphy 2009; Roy et al.

2010). In reference to this thesis, this chapter will only summarise the developments of

the enzyme-responsive approach previously demonstrated within the Ulijn group

(Thornton et al. 2005) as this research provided the fundamental inspirations for the

work described within this thesis.

An enzyme-responsive hydrogel is a polymer that is firstly engineered, tuned or

functionalised so that it responds to a target protease. This is achieved by incorporating

or modifying the polymer with peptides that are recognised by the enzymes within the

body. Such peptides are commonly known as substrates or enzyme cleavable peptides or

linkers (ECPs or ECL, respectively); in this thesis they will be referred to as ECPs. As

previously mentioned in section 2.6.3 for a hydrogel to be ‘responsive’ it is defined as a

polymer that will transform its dimension or conformation as a result of physiochemical

or biochemical stimuli (such as pH, temperature, ionic strength etc.). In this fashion, the

enzyme-responsive hydrogel is further modified by an ‘actuator’ which dictates and

manages the electrostatic interactions of the polymer in response to its external

environment and this causes the polymer to macroscopically transform its structure or

swelling property (Ulijn 2006; McDonald et al. 2009).

In 2005, Ulijn and co-workers functionalised PEGA (hydrogel) particles with positively

charged ECPs i.e. Fmoc-Arg(+)-Phe-Gly or Fmoc-Arg(+)-Gly-Phe in each case the Arg

residue acted as the positively charged actuator that caused the PEGA particles to swell

which in turn increased the molecular accessibility of the particles as a result of

electrostatic repulsion between the positive charges of the ECPs. Additionally, they

revealed that when these positively charged PEGA particles were treated with a protease,

such as thermolysin (functions as the external stimulus) the cleaving of the ECPs by

thermolysin resulted in the removal of the terminal positive charge (Arg(+)) from PEGA

particles. As a result, this caused PEGA particles to structurally collapse due to the


92

reduction in electrostatic repulsions and accordingly the molecular accessibility of the

PEGA particles; and in this manner thermolysin became entrapped within the PEGA

particles (Thornton et al. 2005). The inspiration of this thesis arose after Ulijn and co-

workers envisaged that these macroscopic transitions of the hydrogel particles could

potentially be used to selectively remove unwanted macromolecules from biological

systems (Thornton et al. 2005; Ulijn et al. 2007a) such as chronic wounds in which this

thesis investigates.

As previously encountered in chapter 2: abundant levels of NE was the culprit of

propagating positive feedback mechanisms causing the elevation of other proteases (e.g.

MMPs) with an overall effect of developing debilitating chronic wounds (see section 2.5).

Ideally, reducing the levels of elastase activity in chronic wounds would reduce the

vicious cascade reactions of elastase, thus restrain the synthesis of other proteases and

overall decrease the proteolytic activity observed in chronic wounds. By utilising the

macroscopic swelling/ collapsing property of hydrogels via the enzyme-responsive

approach developed by our group (as summarised above) an elastase-responsive hydrogel

wound dressing could potentially be designed so that it could exclusively and selectively

mop-up excess elastase and then entrap it within the hydrogel matrix.

For the basis of this elastase-responsive wound dressing, the hydrophilic PEGA resin

(figure 1) in the form of particles was chosen as a suitable hydrogel matrix. As

encountered previously, PEGA is a three dimensional hydrogel that is synthesised by the

co-polymerisation of polyacrylamide and PEG (Meldal 1992). The advantages of using a

hydrogel wound dressing to treat chronic wounds has previously been covered in chapter

2 (see section 2.6 for details). PEGA particles were chosen for several reasons

compared to other polymers. Firstly, Meldal and colleagues showed that PEGA resins

are much better compared to the current polymer supports for peptide synthesis (Medal

1992; Renil et al. 1998) since PEGA particles are compatible (i.e. stable) and permeable

and to both chemical (organic) and biological reagents meaning that ECPs can easily be

immobilised to the matrix of PEGA via SPPS utilising standard Fmoc chemistry.

Secondly, there has been increasing reports showing the compatibility and permeability

of PEGA particles with biological molecules such as enzymes for enzymatic reactions,

especially the complete accessibility of enzymes throughout the core of PEGA particles


93

(Meldal et al. 1994; Kress et al. 2002; Ulijn et al. 2002; Thornton et al. 2005). Generally,

it is estimated that PEGA1900 has a molecular weight cut-off 35 kDa (Kress et al. 2002;

Ulijn et al. 2002) however enzymes of 50 kDa – 80 kDa have also been reported to enter

the matrix of PEGA1900 (Auzanneau et al. 1995; Meldal et al. 1994; Kress et al. 2002) to

enable enzyme catalysis inside PEGA particles (Meldal et al. 1994; Thornton et al. 2005).

Other advantages of PEGA particles include: they are transparent, under the correct

stimuli they are able to swell and collapse (Thornton et al. 2005, 2007, 2008; McDonald

2009), they are bio-inert and with PEG being the major component of these PEGA

particles they are highly flexible and biocompatible. Additionally, PEG retards non-

specific protein adsorption to the surface of the particles and this feature is interesting

because after functionalising the PEGA particles with ECPs, the specific sensing-

actuation process intended to mimic the elastase substrates involved during the wound

healing can exclusively enhance or solely direct the biocatalysis of elastase compared to

other chronic wound proteins.

Commercially, enzymes (such as NE) are very expensive and essentially the purification

of enzymes to gain high yields can take a long time as many steps are required to attain

good purity (Komives and Chen 2004). For proof-of-concept of removing excess

elastase from sample fluids that are mimicking CWFs, porcine pancreatic elastase (PPE,

EC 3.4.21.36) was chosen as the model protease to NE because of its similaries to NE.

Firstly, PPE is a serine protease (Barrett et al. 2004) which contains the catalytic triad

His–Asp–Ser within its active site and accordingly shares the general catalytic mechanism

to other serine proteases such as NE as illustrated by figure 8. Secondly, the active site

region of PPE has been reported as being similar to NE (Barrett et al. 2004) and PPE

shows 40% homology to NE (Travis 1998). In comparison to NE, it is the most studied

serine protease for determining elastase activity and alongside NE the broad substrate

specificity of PPE includes its preference to cleave elastin which as mention in chapter 2

is considered as the main substrate for elastase(s). PPE prefers to cleave peptide bonds

at the carboxyl-terminal side of amino acid residues with small hydrophobic, non-polar

side chains such as Ala, Gly (Stryer 1995; Barrett et al. 2004) including Val and Leu

(Travis 1998). In comparison to NE, catalytically PPE has preference to hydrolyse

peptides with Ala~X bonds more readily (Purich and Allison 2002; Travis 1998; Barrett

et al. 2004) than Val~X and Leu~X bonds (Travis 1998).


94

Similarly, thermolysin was chosen as a control protease because it is a bacterial elastase

(Morihara and Tsuzuki 1967) that is isolated from Bacillus thermoproteolyticus and belongs

to the metalloendopeptidase family of enzymes (Barrett et al. 2004; Purich and Allison

2002). It contains a functional Zn2+ ion and four calcium ions bound to the polypeptide

chain (316-residue) with a 3D structure at 1.6Å resolution (Holmes and Matthews 1982).

Compared to PPE, the activity of thermolysin depends on the Zn2+ ion and it is reported

as being extremely stable as well as being four to eight times more active than PPE

(Morihara and Tsuzuki 1967) depending on the substrate. The specificity of thermolysin

has preference to cleave at the N-terminal side of hydrophobic amino acid residues in the

P1′ position e.g. Ala, Gly, Phe, Leu, Ile or Val (Morihara et al. 1968; Barrett et al. 2004;

Purich and Allison 2002) in which the order of preference is Leu > Ala > Phe > Gly

(Barrett et al 2004). Interestingly, thermolysin has a molecular weight of 35,000 kDa

(Miki et al. 1996; Barrett et al. 2004) which is the suitable molecular weight for enzyme

catalysis on PEGA particles and various studies have demonstrated thermolysin as being

robust in completely penetrating into the core PEGA particles (Ulijn et al. 2002; Basso et

al. 2003; Thornton et al. 2005, 2007, 2008; McDonald et al. 2009).

3.2 OBJECTIVES

The research objectives of this chapter include firstly characterising unmodified PEGA

particles by FTIR. Secondly, design short charged enzyme cleavable peptides (ECPs) to

functionalise PEGA particles in response to the elastase and subsequently monitor the

loading of the immobilised ECPs to PEGA particles. Thirdly, investigate the enzyme

hydrolysis of the ECPs coupled to PEGA particles by proteases on the basis of charge

under the influence of pH in the range of chronic wounds. Finally, to study the swelling

behaviour of these particles as a function of ionic strength and pH in order to

understand the swelling hence molecular accessibility of functionalised PEGA particles at

low to high ionic strength.

3.3 MATERIALS & METHODS

3.3.1 Materials

PEGA(800 and 1900)–NH2 particles in methanol were supplied by Polymer Laboratories Ltd

(UK). Fluorenylmethoxycarbonyl (Fmoc) protected amino acids and peptides (Fmoc-


95

Ala-OH.H2O, Fmoc-Ala-Ala-OH, Fmoc-Arg(pbf)-OH, Fmoc-Glu(OtBu)-OH, Fmoc-

Gly-OH, Fmoc-Gly-Gly-OH, Fmoc-Phe-OH, Fmoc-Phe-Phe-OH) were all from

Bachem (UK). Elastase (pancreatic from porcine pancreas, PPE, purity 8 units/mg of

protein, EC 3.4.21.36) and Thermolysin (from Bacillus thermoproteolyticus, purity 40

units/mg of protein, EC 3.4.24.27), trifluoroacetic acid (TFA), silver nitrate (AgNO3)

were from Fluka Analytical (UK). Aldrich (UK) supplied acetonitrile (ACN),

dimetylformanide (DMF), methanol (MeOH), 1-hydroxybenzotriazole (HOBt), N,N'-

diisopropylcarbodimide (DIC), potassium cynanide (KCN), pyridine, phenol and ethanol

(EtOH) and acetic acid. Piperidine, 3-[(3-Cholamidopropyl)dimethylammonio]-1-

propanesulfonate (CHAPS), potassium mono-/di-phosphate solutions (1 M) in deionised

water, glutardialdehyde were purchased from Sigma (UK). HiPerSolv HPLC grade ACN

was from VMR international (UK). Precasted pH 3 – 10 isoelectric gels (15-well, 15µl,

8.6 x 6.8 cm), isoelectric focusing stain/markers, lameali sample buffer, IEF loading

buffer, Mini-Protean® 3 Cell and PowerPacTM Universal for electrophoresis were all

supplied by Biorad (Hertfordshire, UK). Sodium thiosulphate, sodium acetate, sodium

carbonate (Na2CO3) anhydrous, glycine, diaminoethanetetra – acetic acid disodium salt

(EDTA Na2+), glycerol solution were from Fisher Scientific. Formaldehyde (37% w/v)

was from Sigma-Aldrich; and trichloroacetic acid was from BDH. All solvents used were

of the highest purity without further purification. Isolute double fritted filtration

columns were purchased from Kinesis. Stuart Scientific supplied the blood rotator and

roller mixer. HPLC 680 series was purchased from Dionex and the C18 nucleosil

column was purchased from Fisher.

3.3.2 Methods

3.3.2.1 Fourier-transform infrared spectroscopy (FT-IR)

Unmodified PEGA particles (0.1g) were dried in a vacuum oven overnight at ambient

temperature. A small amount of dried PEGA particles were transferred onto the

diamond ATR-platform of the Nicolet 5700 FT-IR spectrophotometer and scanned

using a diamond tip (used for polymer analysis). The diamond tip was tightly pressed on

top of the unmodified PEGA particles to reduce the thickness as well as ensuring that no

air gaps were present. The FTIR spectra of unmodified PEGA particles was scanned

between the wavelength of 4000 – 500 cm-1 region of the electromagnetic radiation over

128 scans at an interval of 1.9285 cm–1.


96

3.3.2.2 Fmoc solid phase peptide synthesis

The pioneered solid phase peptide synthesis (SPPS) method by R. Bruce Merrifield

(1963) was used to functionalise unmodified PEGA particles with neutral and charged

ECPs (figure 19 and table 13) using stepwise Fmoc chemistry (Carpino and Han 1972)

which involved covalently attaching the C-terminus of an activated amino acid to PEGA

particles by sequential coupling followed by the de-protection of protecting groups and

washing steps until the desire peptide sequence was coupled. A fresh batch of

unmodified PEGA particles was functionalised for each experiment.

The SPPS mechanism is illustrated by figure 18. The initial step involved activating an

Fmoc-amino acid (10 equivalents in DMF) with the coupling reagents DIC (5 equivalents

in DMF) and HOBt (10 equivalents in DMF) for 15 minutes on a roller. Unmodified

PEGA particles (~ 0.1 – 2.0 g wet) were washed using wash 1 (entry 1, table 9). The C-

terminus of the activated Fmoc-amino acid was coupled to amino group of the washed

unmodified PEGA particles on a blood rotator for a minimum of 1 hour – maximum

overnight. The modified PEGA particles were washed with wash 2 (entry 2, table 9) and

the successfulness of the coupling reaction was monitored using the Kaiser test (section

3.4.2.3); and the variation in the coupling time also depended on the results achieved

from the Kaiser test. If needed the coupling reaction of the activated Fmoc-amino acid

was repeated to efficiently ensure the coupling of all possible amine groups present

within PEGA particles with Fmoc-amino acid was complete. Before commencing to the

next step (after the Kaiser test) modified PEGA particles were washed thoroughly again

with wash 1 (entry 1, table 9).

Table 9. Washing steps during SPPS

Wash Wash 10x each with Reasons of prewashing step:

1 MeOH, DMF;MeOH (50:50), DMF unmodified PEGA particles or before the coupling step of an amino acid

2 DMF, DMF:MeOH (50:50), MeOH, EtOH Before the ninhydrin test

3 DMF, DMF:MeOH (50:50), MeOH, dH2O Before removing protecting side chain groups (pdf, OtBu)

4 DMF, DMF:MeOH (50:50), MeOH End of SPPS and preparing the particles for storage


97

The mechanism of functionalising PEGA particles with Fmoc-ECPs (di/ tripeptide):

O NH

O

R1

O

O

N = C = N CH3

CH3

CH3

CH3

H+

O NH

O

R1

O

O N CH3

CH3

CH3

NH2

O

NH2+O

NH

O

R1O

CH3

NH

CH3

N CH3

CH3

NH

CH3

O

NNH

O

R1O

H

N

NHCH3

CH3

NH

CH3

O CH3

O

NNH

O

R1O

N

CH2

H O

NNH

O

R1O

NH

O

NNH2

R1

CO2

O

NNH

O

R2O

N

R1

O

O

NNH

O

R3O

N

R2

O

N

R1

O

O

NNH

O

R2O

N

R1

O

O

NNH

O

R3O

N

R2

O

N

R1

O

+ HOBt/ DMF(15 minutes)

STEP 1: Activation of Fmoc-amino acid

STEP 2: Coupling activated Fmoc-amino

acid to PEGA particles

(ninhydrin test)

+

STEP 3: Deprotection of Fmoc group+ DMF (2 hrs)

H+

H+

+

+

(1)

(2)

(5) (6)

(7)

(9)

(4)

(8)

(ninhydrin test)

(3)

STEP 4: Deprotect protecting

side chains

(10) (11)

Fmoc-ionic ECP-PEGA particles (charged)

+ 95% TFA/ Water(2 hours)

(12) (13)

Fmoc-ECP-PEGA particles (uncharged)

(+/-) (+/-)

(repeat steps 1 & 2 if Fmoc-amino acid

has not fully coupled to PEGA particles)

(repeat steps 1, 2 & 3 until

desire length of Fmoc-ECP

has coupled to PEGA particles)

Figure 18. Mechanism for the functionalisation of unmodified PEGA particles with enzyme cleavable peptides (ECPs) using Fmoc SPPS. STEP 1: Fmoc-protected amino acid (2) reacts with the activating reagent DIC (1) to form the active intermediate ester, O-acylisourea (3) in the presence of HOBt which prevents racemisation and reduces the formation of the unreactive N-acylurea. STEP 2: O-acylisourea collapses by intramolecular acyl transfer when it reacts directly with the amine groups of the PEGA particles (4) to form an amide bond between Fmoc-amino acid and PEGA particles (5) alongside the by-product, diisopropylurea (6). Completion of the coupling reaction is monitored using the Kaiser test (section 3.4.2.4). STEP 3: The Fmoc group is de-protected with piperidine (7) and several intermediates including the adduct dibenzofulvene (8) and de-protected PEGA particles (9) are formed. Again completion of the de-protection step is monitored using the ninhydrin test. Steps 1 – 3 are repeated until the desired length of ECP is coupled to PEGA particles producing uncharged Fmoc-ECP-PEGA particles (10, 11), where the length of ECP is a dipeptide (10) or a tripeptide (11). STEP 4: Fmoc-ionic-ECP-PEGA particles (12, 13) are produced after treating (10) and (11) with 95% TFA:5%H2O. Figure tailored from Montalbetti and Falque 2005; Newcomb et al. 1998; Jone 2002; Carey and Sundberg 2001; and Sabationo et al. 2004.


98

O

NNH

O

R2O

N

R1

O

O

NNH

O

R3O

N

R2

O

N

R1

O

O

NNH

O

R2O

N

R1

O

O

NNH

O

R3O

N

R2

O

N

R1

O

Gly

Phe

CH3

HCH3

Arg+

N N

N+

Glu

O

O

Gly

PheCH3

H

PheCH3

Arg+

N N

N+

Glu

O

O

(+/-)

(+/-)

(a)

(c)

(b)

(d)

R1, R2 Ala R1

R2

Ala

R1, R2 Ala

R3

R1, R2 Ala

R2

Figure 19. Chemical structures of uncharged (neutral) and charged Fmoc-ECPs coupled to PEGA particles. Amino acids with uncharged R-groups: alanine (Ala), glycine (Gly), phenylalanine (Phe); and charged R-groups: arginine (Arg(+)) and glutamic acid (Glu(–)). Neutral ECPs (a, c) generating Fmoc-ECP-PEGA particles; charged ECPs (b, d) generating Fmoc-ionic ECP-PEGA particles. The length of ECPs were designed as a dipeptide (a, b) or a tripeptide (c, d).

Prior to coupling the next amino acid, the Nα – protecting Fmoc groups were de-

protected using 20% piperidine dissolved in DMF on blood rotator for 2 hours at room

temperature (Note: at this stage the filtration columns were wrapped in foil). For the

mechanism of Fmoc deprotection via piperidine refer to figure 18: step 3. The de-

protected Fmoc groups were collected for UV/VIS analysis to calculate Fmoc loading

(as described in section 3.4.2.4) to PEGA particles. The modified PEGA particles were

washed again using wash 2 (entry 2, table 9) and the Kaiser test was repeated again to

ensure all Fmoc groups were totally removed. The modified PEGA particles were

washed again using wash 1 (entry 1, table 9). The sequential coupling of activated Fmoc-

amino acids, de-protection of Fmoc-group, Kaiser test and washing steps were all

repeated until the desired ECP was achieved, producing functionalised PEGA particles.

In order to achieved charged ECPs i.e. Fmoc-ionic ECP-PEGA particles, the protecting

side chain groups tert-butyl ester (OtBu) and pentamethyldihydrobenzofuran-5-sulfonyl

(Pbf) for glutamic acid (i.e. Fmoc-Glu(OtBu)-OH) and for arginine (i.e. Fmoc-Arg(pbf)-

OH) were removed by treating the functionalised PEGA particles with 95% TFA:5%

dH2O for 2 hours on blood rotator and the washing step prior to removing the


99

protecting side chain groups was wash 3 (entry 3, table 9). At the end of modification, all

functionalised PEGA particles were thoroughly washed using wash 4 (entry 4, table 9)

and if not used immediately the functionalised PEGA particles were vacuum dried using

a filtration tank to remove all excess MeOH and then stored in fridge.

3.3.2.3 Kaiser test

Ninhydrin test referred to as the Kaiser test (Kaiser et al. 1970) in this thesis was used as

a simple rapid colormetric test for detecting the presence of free amino groups in order

to observe whether each of the coupling/ de-protecting steps during SPPS were

complete. Two Kaiser stock solutions (A and B) were prepared. Kaiser solution A: 20 µl

of aqueous KCN solution (13 mg KCN in 20 ml dH2O) was diluted with 980 µl pyridine

and then further diluted to 100 µl with phenol/ethanol solution (8.0 g phenol dissolved

in 2 ml ethanol). Kaiser solution B: 1.0 g ninhydrin was dissolved in 20 ml ethanol.

Both stock solutions were stored in foil wrapped containers at room temperature. To

carry out the Kaiser test, two drops from each of the Kaiser solution (A and B) were

added to a 5 mg of modified PEGA particles and the colour was observed after heating

the PEGA particles at a temperature > 100oC for 5 minutes. Unmodified PEGA

particles were always used as a standard reference alongside modified PEGA particles

since the unmodified PEGA particles possess free amine groups and will always give a

positive result (as summarised in table 10).

Table 10. Colour observation of PEGA particles and surrounding solution after Kaiser test.

Entry Kaiser Test Solution PEGA particles Concluding observation

1 Positive Clear Blue Amino groups (NH2) present

2 Negative Yellow Clear No amino group present, amide bond produced (RNHC=O)

3.3.2.4 Fmoc loading

Unmodified PEGA particles (0.1 gram) were left to dry overnight in a vacuum oven at

room temperature. The weight of the dried PEGA particles was measured and the

percentage weight loss was calculated using mass difference of wet and dry weight of

PEGA particles. The percentage weight loss for both PEGA800 and PEGA1900 was

calculated as 9.16 % and 6.00 % respectively.


100

A stock solution of 5 mg/ml was prepared by dissolving Fmoc-Ala-OH in 20%

piperidine: DMF (in foil wrapped container at room temperature) to deprotect the Fmoc

group from Fmoc-Ala-OH. This solution was then diluted 1/100 with DMF and from this

diluted solution four individual standard solutions were prepared with a concentration

ranging from 0.01 – 0.04 mg/ml in DMF. Fmoc loading of PEGA particles was achieved

by coupling Fmoc-amino acids (such as Fmoc-Ala-OH) to unmodified PEGA particles

(0.5 grams) using Fmoc SPPS (as described above). As stated above, the removal of

Fmoc group was achieved by treating the modified PEGA particles with 20%

piperidine:DMF (2 ml) on a blood rotator for 2 hours. To collate all possible Fmoc

groups, modified PEGA particles were further washed with DMF (8 ml) and this

solution was further diluted 1/100 with DMF. The concentration of the Fmoc groups for

both standards and samples was measured using UV/Vis spectrophotometer (at 301 nm)

using a 1 cm path-length quartz cuvette. For the standards a linear curve was produced

(figure 20) which was used to calculate the Fmoc loading of the functionalised PEGA

particles as summarised by equation 1.

y = 7.7594x

R2 = 0.9998

0.0

0.3

0.5

0.8

1.0

1.3

1.5

0.00 0.05 0.10 0.15 0.20

Fmoc (mmoles/L)

Absorbance (at 301 nm)

Figure 20. Standard curve for the absorbance (301nm) of Fmoc group.

Equation 1. Fmoc loading calculation:

mmoles = Absorbance (301nm)

Gradient (7.7594)x volume collected (L) x Dilution factor

Mass of PEGA (g) = Mass of PEGA (g) x (% Weight loss ÷ 100) dry wet

Mass of PEGA (g) dry

mmoles Loading (mmoles/g) =


101

3.3.2.5 Isoelectric focusing

The pI value of elastase (1 mg/ml was dissolved in various solutions lameli buffer, water

and CHAPS) was determined using 1-D gel isoelectric focusing on 15 well isoelectric gels

(pH 3 – 10) using a Biorad-Miniprotean cell. Five microlitres of both the IEF stain and

elastase were loaded on to the gel. Voltage steps were applied for: 1 hour at 100 V, 1

hour at 200 V and 30 minutes at 500 V. After separation, the gel was stained using the

silver nitrate approach (all solutions were heated to the desired temperature using a

hotplate and all washing steps were manually shaken) as follows:

The gel was first fixed by washing the gel with fixing solution 1 (20 % w/v

trichloroacetic acid) at room temperature for 15 minutes; followed by fixing solution 2

(20 % v/v ethanol, 8 % v/v acetic acid) at 50oC for 10 minutes and then finally with

fixing solution 3 (20% v/v ethanol, 8% v/v acetic acid, 0.4% v/v glutardialdehyde) at

50oC for 10 minutes. After that, the gel was washed with the incubation solution (0.1 %

w/v sodium thiosulphate, 0.4M sodium acetate with the pH adjusted to 6.5 using acetic

acid, 0.4 % v/v glutardialdehyde) at 50oC for 10 minutes. Then washed with the rinsing

solution (20 % v/v ethanol, 8 % v/v acetic acid) at 50oC for 10 minutes; and finally the

gel was washed three times with deionised water at 50oC for 10 minutes.

Next, the gel was washed and then incubated with the silvering solution (0.1 % AgNO3,

25 µl of 37 % w/v formaldehyde per 100 ml) at room temperature for 20 minutes. Next

the gel was washed using two developing steps: developing step 1 (2.5 % w/v Na2CO3,

32.5 µl of 37 % w/v formaldehyde) at room temperature for 1 minute; followed by

developing step 2 (2.5 % w/v Na2CO3, 32.5 µl of 37 % w/v formaldehyde) at room

temperature until elastase bands appeared and were of the desired intensity. The gel was

then de-silvered (with 2 % (w/v) glycine, 0.5 % w/v EDTA Na2+ at room temperature

for 10 minutes. (Note: the waste from the silvering step on-wards was poured into a

suitable container for silver nitrate waste). Finally, the gel was prepared for drying by

washing it in 2 % (v/v) glycerol solution at room temperature for 10 minutes. The gel

was then preserved by transferring the gel on to a plastic cassette and then left to air dry

completely at room temperature for 2 days. The gel was subsequently wrapped with

cling film and stored at 4ºC.


102

3.3.2.6 Cleaving functionalised PEGA particles with proteases

After SPPS, functionalised PEGA particles were thoroughly washed with MeOH, dH2O

and potassium phosphate buffer at the appropriate pH and ionic strength (as indicated).

Functionalised PEGA particles (50 mg) were then cleaved with elastase and thermolysin

(1 ml at 0.1 or 1.0 mg/ml in 0.001M at pH 6.0 – 9.0) overnight at 34 – 37 oC. The

supernatants of the cleaved products were collected and the reaction was stopped by

washing the PEGA particles with ACN:H2O (80:20, 9 ml) containing 0.1% TFA. This

washing also ensured that all the cleaved products were collected from the interior of the

functionalised PEGA particles. The procedure was carried out in duplicates (n = 2 or 4,

as indicated).

For enzymatic analysis and in order to quantify the percentage cleaved product yield after

enzyme cleaving, 20 µl of the cleaved product was injected and analysed by reverse-phase

HPLC (rpHPLC), a technique used to achieve separation of the collected cleaved

product. A linear curve was produced for the standard solution of Fmoc-Ala-OH in the

range of 0.00 – 0.40 mmoles (figure 21) by dissolving Fmoc-Ala.H2O.OH in ACN:H2O

(80:20); and the percentage cleaved product by a given protease was calculated according

to equation 2.

y = 122.07x

R2 = 0.9928

0

11

22

33

44

55

0.00 0.10 0.20 0.30 0.40

Fmoc (mmoles)

Area of Fmoc peak at 301 nm

(mAU*min)

Figure 21. Standard curve for the area of Fmoc group (301nm) via HPLC.

The Dionex HPLC system consisted of P680 pump with ASI-100 Automated sample

injector. Chromatographic analyses were carried out using a Nucleosil C18 reverse-phase

column (5 µm particle size, 100 mm x 4.6 mm I.D.) connected to a UVD170U detector.


103

The mobile phase consisted of ACN:H2O (0.1% TFA) and 0.1% TFA:H2O and was

pumped at a flow rate of 1 ml/min with a solvent gradient of 40% (ACN:dH2O:TFA)

gradually rising to 90% (ACN:dH2O:TFA) over the course of 25 minutes. The cleaved

product was quantified by detecting the UV absorbance at λ = 254 nm (capped/

uncapped amino acids and peptides) and at λ = 301 nm (Fmoc group). HPLC

chromatographs were analysed using Chromeleon 6.60 software and the retention time

of cleaved product was compared with known standards.

Equation 2. Percentage cleave product using Fmoc group obtained:

Fmoc cleaved (mmoles) = Peak Area (301nm)

Gradient (122.07)x Dilution factor

Mass of PEGA (g) = Mass of PEGA (g) x (% Weight loss ÷ 100) dry wet

Mass of PEGA (g) dry

Fmoc cleaved (mmoles) Fmoc cleaved (mmoles/g) =

% Cleaved product = Loading (mmoles/g)

x 100 Fmoc cleaved (mmoles/g)

3.3.2.7 Swelling

Potassium phosphate buffer at appropriate pH (ranging from 7.0 – 9.0) and ionic

strength (0.001M – 0.2 M) were prepared by serial dilution of the 1 M potassium mono-

and di-phosphate solutions in deionised water. Each buffer solution was adjusted to its

desired pH with HCl and NaOH using a pH meter. After SPPS, functionalised PEGA

particles (ionic and neutral) were thoroughly washed with dH2O to remove all traces of

MeOH and were then further washed with the appropriate buffer (at the required pH

and ionic strength). A small amount of functionalised PEGA particles (~ 50 mg) were

swollen in 3 mls of potassium phosphate buffer at the appropriate pH and ionic strength

for 5 minutes. With the aid of the SPOTCam software, photographic images were taken

using a camera coupled to a light microscope (at x10 magnification) of the swollen

functionalised PEGA particles and the diameters (mm) of 100 swollen particles were

measured using the SPOTCam software.


104

3.3.2.8 Statistics

Statistical analysis was carried out with the SPSS for Windows statistical package (SPSS

Inc., version 16.0). The parametric ANOVA test (one-way and three-way, full factorial)

was used to evaluate the statistical differences between: (1) the specificity of proteases in

relation to hydrolysing ECPs on the basis of charge under the influence of pH; and (2)

the diameter measurements of PEGA particles depending on both the pH and ionic

strength. For pair-wise comparisons, post-hoc tests (Tukey and Duncan) alongside a 2-

tailed paired t-test was conducted to determine which ECP or pH affected specificity of

each protease; or which diameters differed significantly from one another. Statistically a

significant difference was observed at the 95% confidence level in which values with p <

0.05 were considered significant and values with p > 0.05 were not significant.

3.4 RESULTS & DISCUSSION

3.4.1 FT-IR of PEGA particles

Since PEGA particles were commercially purchased (Polymer labs, UK) it was essential

to determine molecular structure of unmodified PEGA particles using Fourier-transform

infra-red (FTIR) spectroscopy. FTIR is a non-invasive technique that enables the

functional groups of molecules to be determined by a single beam of infra-red (IR)

through the sample (Faust 1992) in which the IR is absorbed within the electromagnetic

radiation region with a frequency ranging from 4000 – 400 cm-1 (Manning 2005). The

atoms of a molecular structure are in constant motion relative to each other, although the

atoms are linked by chemical bonds their positions are not fixed so during FTIR they are

subjected to several vibrations in which the absorbed IR radiation is converted into

energy of molecular vibrations as the bonds move from the lowest vibrational state to

the next highest due to the bending and stretching of bonds (Nakanishi 1964).

Unmodified PEGA particles were initially dried in a vacuum oven at ambient

temperature and then analysed by FTIR. Figure 22 displays the FTIR spectra of both

PEGA800 and PEGA1900 and the assigned functional groups (entries 1 – 15) are

summarised in table 11. The FTIR spectra of both PEGA800 and PEGA 1900 were found

to be the same and the intensity of some vibrations were weak especially within the


105

fingerprint region (< 1500 cm-1) and the functional region in the range of 3500 – 3100

cm-1 due to the overlap of various functional groups absorbing within the same IR region

(Furniss et al. 1989).

NH

O n

NHO

NHO

*

*

NH2

O

n

n

CH3

CH3

OO C

H2

CH

2

O CH3

xl

n

NH2

CH2

CH2 O

CH3

xl

3, 10

1, 2, 14

3, 6, 7, 10

15

9, 12

1, 2, 14, 5, 10

11, 12

3, 6, 7, 10

15

12, 13, 15

1, 4,

5, 11

10, 13

10, 13

3, 10, 13, 15

10

3, 6, 8

11

11 3, 6, 83, 6, 83, 6, 8

1, 4,

5, 11

1, 4,

5, 11

10

1, 4,

5, 11

99

9

9

11, 12

Figure 22. FTIR spectrum of unmodified PEGA(800 and 1900) particles (top) and the molecular structure of PEGA. Details of the assigned numbers are given in table 11.

The broad stretch ranging from 3500 – 3100 cm-1 was collectively the result of aliphatic

N–H stretches for both amines and amides (Furniss et al. 1989; Rubinson and Rubinson

2000; Simons 1978; Coates 2000). Within this range, the aliphatic vibration at 3344.2 cm-

1 (entry 1: figure 22, table 11) originated from two types of NH2 stretches: (1) a primary


106

amine i.e. >CH–NH2; and (2) a primary amide i.e. – C=O–NH2 (Coates 2000; Rubinson

and Rubinson 2000).

Table 11. Assignments of FTIR frequencies of unmodified PEGA(800 and 1900) particles.

Entry Wavenumber (cm-1) Functional Assignments

1 3344.2 N–H (aliphatic) stretch: 1o NH2 for amine (i.e. > CH–NH2) and amide (i.e. –C=O–NH2) including a 2o amide (i.e. –C=O-NH–R).

2 3204.8 N–H (aliphatic): 1o amine stretch; and 1o amide bend

3 2865.1 C–H (aliphatic) sym. stretch: CH2 and CH3

4 1662.8 C=O stretch (amide I band: 1o and 2o amides); scissoring NH2 (1o amide)

5 1545.7 N–H bend: amide II band (2o amides) and 1o amine (>CH–NH2)

6 1450.7 C–H (aliphatic) bend: scissoring of CH2 (i.e. CH2–C–O) and CH3 (asym.) deformation of C–CH3

7 1411.1 C–H (aliphatic) deformation adjacent to C–O (i.e. CH2–C–O)

8 1372.2 CH3 (sym.) deformation of C–CH3

9 1347.8 C–H (aliphatic) bend; wagging C–H amide bend; and an amine C–N

10 1298.6 CH2 (wagging and twisting); C–O stretch; C–C skeletal vibration; C–N amines

11 1248.8 N–H bend (amide III band); C–O (asym.) stretch of C–O–C

12 1089.9 C–O stretch (asym. and sym.); amine C–N stretch; C–C skeletal vibration

13 946.0 C–C stretch

14 844.3 N–H (oop) arising from NH2 twisting and wagging deformation

15 ~ 750 – 700 C–C backbone; (CH2)n rocking (where n < 4)

Abbreviations: 1o (primary); 2o (secondary); sym. (symmetrical); asym. (asymmetrical); bend a.k.a. deformation; amide I band (C=O stretch + C–N); amide II band (N-H in-plane bend + C–N stretch); amide III band (N–H bend + C–N/ C-C stretch + C=O in-plane bend); and oop (out-of-plane). This table was tailored together from: Kuptosov and Zhizhin 1998; Nakanishi 1964; Furniss et al. 1989; Smith 1999; Bellamy 1975; Faust 1992; Rubinson and Rubinson 2000; Coates 2000; Manning 2005; Ganim, et al. 2008.

Additionally, the vibration at 3204.8 cm-1 (entry 2: figure 22, table 11) were collectively

due to an amine N–H stretch and a primary amide N–H bend (Nakanishi 1964; Furniss

et al. 1989; Kuptosov and Zhizhin 1998). The N–H bend vibration of the primary amide

at 3204.8 cm-1 is sometimes referred to an ‘amide II’ or ‘amide A’ band (Manning 2005;

Kuptosov and Zhizhin 1998; Ganim et al. 2008) as indicated in table 12. In terms of

PEGA, the NH2 stretch of the primary amine corresponds to the free amine group that

is used to couple Fmoc-amino acids/ECPs via SPPS (as highlighted in green in figure

22); and the NH2 of primary amides arises from the polyacrylamide backbone (as

highlighted in red in figure 22).


107

An NH2 stretch is normally observed as a doublet consisting of two absorptions

indicating the asymmetrical and symmetrical stretches for both primary amines and

amides (Furniss et al. 1989; Simons 1978; Smith 1999). From figure 22 the asymmetrical

and symmetrical N–H stretches for primary amine were observed at 3344.2 cm-1 and

3204.8 cm-1, respectively (Nakanishi 1964; Furniss et al. 1989; Kuptosov and Zhizhin

1998; Coates 2000; Smith 1999). The asymmetrical N–H stretch for primary amide was

also observed at 3344.2 cm-1 (Kuptosov and Zhizhin 1998; Furniss et al. 1989) whereas

the symmetrical N-H stretch could not be distinguished within the range of 3500 – 3100

cm-1 because the N–H vibrations were weakly absorbed within this region due to the

overlap of other N–H (free or associate) groups such as an N–H (aliphatic) stretch of a

secondary amide i.e. –C=O–NH–R also absorbing at 3344.2 cm-1 (Furniss et al. 1989;

Nakanishi 1964; Smith 1999) including the N–H vibrations as previously mentioned.

Furthermore, N–H bend vibrations were observed: at 1545.7 cm-1 (entry 5: figure 22,

table 11) due to a secondary amide i.e. ‘amide II band’ and a primary amine i.e. >CH–

NH2 (Rubinson and Rubinson 2000); at 1248.8 cm-1 (entry 11: figure 22, table 11) was the

result of an ‘amide III band’ (Kuptosov and Zhizhin 1998; Nakanishi 1964). In terms of

amides, the composite conformations of the ‘amide II and III bands’ are discussed below

(see table 12). Finally, an out-of-plane N–H deformation arising from the twisting and

wagging of an NH2 was observed at 844.3 cm-1 (entry 14: figure 22, table 11; Furniss et al.

1989).

A combination of C–H stretches and bends were absorbed at several frequencies (entries

3, 6-10 and 15: figure 22, table 11) resulting from various C–H functional groups. A

medium C–H (symmetrical) stretch due to both CH2 and CH3 vibrations were

collectively observed at 2865.1 cm-1 (entry 3: figure 22, table 11); and since the absorption

was below 3000 cm-1 this confirmed that the molecular structure of PEGA was aliphatic

and not aromatic (Faust 1992; Rubinson and Rubinson 2000; Furniss et al. 1989;

Nakanishi 1964; Coates 2000). Within the fingerprint region, C–H bend vibrations

(weak) of CH2 and CH3 were further observed at frequencies ranging from 1475 cm-1–

1290 cm-1 (Rubinson and Rubinson 2000; Coates 2000) and at ~ 750 – 720 cm-1

(Nakanishi 1964) and these are now discussed. At 1450.7cm-1 (entry 6: figure 22, table 11)

a C–H deformation of a scissoring CH2 bend (Furniss et al. 1989); a C–H bend adjacent

to an C–O i.e. CH2–CO group at 1411.1 cm-1 (Furniss et al. 1989; Nakanishi 1964); a

wagging and twisting CH2 deformation at 1298.6 cm-1 (entry 10: figure 22, table 11); and


108

a rocking (CH2)n vibration was observed at ~750 – 720 cm-1 (entry 15: figure 22, table 11)

where n < 4 (Nakanishi 1964). Furthermore, CH3 has two deformations of a methyl-

carbon (i.e. C–CH3): the asymmetric vibration was observed at 1450.1 cm-1 (Furniss et al.

1989; Rubinson and Rubinson 2000) and the symmetric vibration at 1372.2 cm-1 (Bellamy

1975; Furniss et al. 1989; Rubinson and Rubinson 2000). Finally, a combination of an

aliphatic C–H bend alongside an amine C–N bend and a wagging C–H bend of an amide

were all observed at 1347.8 cm-1 (entry 9: figure 22, table 11).

Table 12. Types of conformational amide bands for FTIR frequencies.

Entry Type Wavenumber (cm-1) Functional groups

1 Amide A ~ 3300 N–H stretch

2 Amide B ~ 3080

3 Amide I 1700 – 1600 C=O stretch coupled with a C–N

4 Amide II ~ 3200; 1550 – 1450 N–H (in plane) bend coupled with an C–N stretch

5 Amide III 1350 – 1200; 1290 – 1230 N–H bend coupled with a C–N/ C–C stretch and a C=O bend

Note: the C=O and C–N groups are sometimes written as: C÷O and C÷N respectively. Source: Kuptosov and Zhizhin 1998; Manning 2005; Ganim et al. 2008; Nakanishi 1964.

The molecular structure of PEGA contains both primary (i.e. –C=O–NH2) and

secondary (i.e. –C=O–NHR) amides. Amides are characterised by the functional groups:

C=O, N–H, C–N, and C–C. When the vibrations of these functional groups combine

they tend to absorb within the same IR frequency and their combinations are commonly

characterised into various conformational bands: amide A, amide B, amide I, amide II

and amide III; in which table 12 summaries the distinction between these bands.

The ‘amide I band’ consisting predominantly of a C=O (carbonyl) vibration was strongly

absorbed at 1662.8 cm-1 (entry 4: figure 22, table 11) for both a primary and secondary

amides (Kuptosov and Zhizhin 1998; Nakanishi 1964; Furniss et al. 1989; Rubinson and

Rubinson 2000; Coates 2000; Manning 2005). A scissoring NH2 bend of an amide tends

to overlap and fall at the same frequency or as a shoulder upon the C=O stretch (Smith

1999). As mentioned previously, the N-H bend of secondary amide was absorbed at

1545.7 cm-1 (entry 5: figure 22, table 11) and it is normally expected to absorb at a lower

frequency than the a C=O group of an amide (Furniss et al. 1989) and is usually referred

to an ‘amide II band’ which is a complex vibration consisting of a scissoring N–H (in-

plane) bend strongly coupled with a C–N stretch (Bellamy 1975; Furniss et al. 1989;


109

Kuptosov and Zhizhin 1998; Ganim et al. 2008; Nakanishi 1964). Similarly, an ‘amide

III band’ was also observed at 1248.8 cm-1 (entry 11: figure 22, table 11) and this is

another complex vibration in which the N–H bend is coupled with a C=O bend along

with a C–N/ C–C stretch (Kuptosov and Zhizhin 1998; Manning 2005; Nakanishi 1964).

The amide I, II and III bands appear from the polyacrylamide backbone and the PEG

cross-links within the PEGA structure.

Due to the PEG crosslinks, a strong C–O (rocking) stretch of a symmetrical C–O–C was

observed at 1089.9 cm–1 (entry 12: figure 22, table 11; Nakanishi 1964; Kuptosov and

Zhizhin 1998; Coates 2000) whereas the asymmetrical C–O–C was observed at 1248.8

cm–1 (Furniss et al. 1989; Nakanishi 1964). As expected, the intensity of the C–O stretch

at 1089.9 cm–1 for PEGA1900 was slightly more than PEGA800 this is because PEGA1900

contains more PEG groups (note: the superscript refers to molecular weight of PEG).

Additionally, a C–O stretch was also observed at 1298.6 cm-1 (Rubinson and Rubinson

2000) and it was found to overlap with other functional groups as summarised in table 11

which have previously been discussed elsewhere within this section. Furthermore, C–N

(aliphatic) stretches of primary amine (R–NH2) were observed at 1298.6 cm-1 and 1089.9

cm-1 (Nakanishi 1964; Coates 2000); and finally, the C–C (skeletal/ backbone) vibrations

were observed at: 1089.9 cm-1 (Faust 1992), 946.0 cm-1 (Kuptosov and Zhizhin 1998) and

~750 – 720 cm-1 (Faust 1992).

3.4.2 Homogenous loading of solid-phase peptide synthesis

The combinational method, SPPS first described by Merrifield in 1963 (Merrifield 1963)

based on Fmoc chemistry (Carpino and Han 1972) was used to couple peptides (referred

to as ECPs in this thesis) to PEGA particles (Renil and Meldal 1995). Fmoc SPPS

involved the sequential coupling (sometimes referred as immobilisation) of Fmoc-amino

acids to PEGA particles followed by the de-protection of Fmoc group from the coupled

Fmoc-amino acid, these steps were repeated until the desired length of the ECP was

coupled to PEGA particles. The mechanism of Fmoc SPPS has previously been

illustrated by figure 18; and a list of all the ECPs studied within this thesis are given in

table 13 and the chemical structures of the ECPs have been illustrated previously in

figure 19.


110

O

O

O

O

NNH2

R1

O

O

OH

OH- H2O

NH

O

O

OHN

R1

O

H+

NN

R1

O

O

O

HN

N

R1

O

O

O

:

NN

R1

O

O

O

H

HO

H

NH2

O

O

H

NH2

OH

O

H+

N

R1

O

O

N

O

OO

O

N

O

OO

O

: :

O

O

OH

OH

HO

H

H

NH

O

O

N

R1

OOH

H+

H+

- H2O

(1) (2) (3)

(6)

Anhydrous

form

Hydrated form

(stable)

+

(5a)

+

(5c) stable anion

Hydrolyse

imine

Imine hydrolysis

continues

+

(7)

(8)

(4)

..

..

-

(5b)

Figure 23. Mechanism of ninhydrin with free amino groups on PEGA particles via imine formation. Ninhydrin has two forms: an anhydrous form (1) also known as 1,2,3-triketone and since the middle ketone is electron deficient ninhydrin is used as the hydrate form (2) which is also known as 1,2,3-indanetrione hydrate or triketohydrindene hydrate. The hydrate form of ninhydrin (2) reacts with the free amino groups within PEGA particles (3) to form the intermediate product (4) followed by a ketimine (5a) which stabilises by resonance to form a stabilised imine anion (5b) followed by a new imine (5c). This is then attacked by a water molecule to facilitate the hydrolysis of the imine producing a co-intermediate product (6) and the amine (7). The latter is stabilised by resonance and subsequently the condensation of the free amino group of the amine (7) with another ninhydrin molecule (2) forms the Ruhemann’s purple-blue dye (8). Adapted from Bailey 1990; Sabationo et al. 2004; Friedman 2004).

During each step of the SPPS, the Kaiser test (Kaiser et al. 1970) was used as a rapid

straightforward colorimetric test to monitor the successful completion for both the

coupling and de-protection steps. The mechanism of the Kaiser test (figure 23) involved

the reaction of ninhydrin (hydrate form) with the free amino groups (NH2) of either

amino acids or amines via imine formation to form a purple-blue adduct commonly

referred to as the Ruhemann's purple dye (Bailey 1990; Sabationo et al. 2004; Friedman

2004). The development of the Ruhemann's purple dye normally observed as an intense

blue colour gave a positive Kaiser test indicating both the successful de-protection of the

Fmoc group and an incomplete coupling step; while the coupling of amino acids to


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completion gave a negative test as a yellow colour was observed (Bailey 1990; Sabationo

et al. 2004; Mergler and Durieux 2005).

Figure 24. Colorimetric observations of the Kaiser test. A positive test is observed when Ruhemann’s purple-blue adduct forms, the PEGA particles are stained blue whereas the solution remains clear which indicates the presence of free amino groups (a, c); and a negative test is observed when the blue adduct is not formed, the PEGA particles remain clear and the solution remains yellow and this indicates the formation of an amide bond (b). Unmodified PEGA particles are used as a standard (a) in reference to modified PEGA particles (b, c).

Figure 24 demonstrates the results observed from the Kaiser test in which unmodified

PEGA particles were always used as a reference standard as the unmodified PEGA

particles contains free NH2 groups therefore the Ruhemann’s blue colour was always

observed (figure 24a). Coupling of an amino acid to completion involved forming an

amide bond and this resulted in a negative test in which a yellow colour was observed

(figure 24b); whereas the de-protection of an Fmoc group gave an incomplete coupling

step and this resulted in obtaining a positive test due to the development of the

Ruhemann’s blue colour (figure 24c). It is worth nothing that upon tilting of the vials

containing the PEGA particles soaked within the Kaiser solutions as expected for the

positive test the particles were found to stain blue colour and the surrounding solution

remained clear; whereas in the presence of an amide bond (after successfully coupling an

amino acid to PEGA particles) the particles remained clear and the colour of the

surrounding solution remained yellow.

It was essential to maintain the same loading of amine groups between each of the

coupling/ de-protecting steps during SPPS and in order to achieve homogenous loading

of the amine groups throughout SPPS double coupling of Fmoc-amino acids to PEGA

particles was carried out and in each case the Kaiser test constantly relieved a yellow

colour indicating complete coupling of the amino acid. To quantify the loading of amine

(a) (b) (c)


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functional groups of PEGA particles per gram, the de-protected Fmoc groups were

collected during SPPS and analysed by UV–Visible spectroscopy to determine the

concentration of Fmoc loading per gram of PEGA particles. As a maximum of three

amino acids were coupled to PEGA particles to form an ECP consisting of an Fmoc-

tripeptide sequence, this meant that the de-protected Fmoc groups of the first two amino

acids coupled to PEGA particles were only quantified.

The manufacturer (Polymer labs) reported PEGA800 and PEGA1900 to have a loading of

0.4 mmol/g and 0.2 mmol/g, respectively. The average loading was calculated in the

range of 0.47 – 0.33 mmol/g for of PEGA800 particles; and 0.15 – 0.18 mmol/g for

PEGA1900 particles. Other researchers have also reported variations of loading between

different batches (Ulijn et al. 2003a) and consequently the loading calculation was

repeated for every fresh batch of unmodified PEGA particles functionalised with any

given ECPs to obtain an accurate loading for that particular batch of functionalised

PEGA particles. Additionally, the loading measurements are expected to vary in contrast

to manufacturer’s loading because it is possible that the loading measurements are done

under separate conditions since there are various methods available to calculate the Fmoc

loading on a polymeric support (Sabationo et al. 2004; Cin et al. 2002).

3.4.3 Isoelectric point of elastases

A protease exhibits a net charge of zero when the pH equals the pI value (Stryer et al.

2002) however the charge of the protease can vary when the pH of the surrounding

environment is above and below the pI value. In the literature, confusion arose to the

exact isoelectric point (pI) of elastase i.e. PPE as it was reported to have a pI > 8 – 11

(Travis 1988; Barrett et al. 2004; Worthington Limited). To overcome this issue, several

attempts were carried out to determine the pI value of PPE using isoelectric focusing

(IEF), an experimental technique that separates proteins according to their net isoelectric

charge. Initially PPE was dissolved in sample buffer of lameali buffer and after

separation it was observed that the separation was poor for the standards. Although, the

sample buffer of lameali buffer is appropriate for SDS-page, it is not appropriate for iso-

electric focusing. Since PPE could dissolved in deionised water, the separation was

repeated and after silver nitrate staining the separation was found to be poor for both

standards and PPE. Next, attempts were carried out to dissolve PPE in CHAPS to


113

improve the separation; and again after silver nitrate staining the separation was found to

be poor for both standards and PPE. As IEF was not successful and time consuming it

was abandoned.

.

(a) Primary sequence for 3est (PPE, EC 3.4.21.36)

(b) pI value of 3est (PPE, EC 3.4.21.36)

-40

-30

-20

-10

0

10

20

30

0 2 4 6 8 10 12 14 16

pH

Charge

Figure 25. Total net charge of elastase specifically PPE (EC 3.4.21.36) within the pH range of 1 – 14. The primary sequence (3est) of PPE (a) was entered into pI calculator and was used to plot the total net charge of PPE over pH 1 – 14 (b). The overall net charge i.e. pI value was predicted as 8.31; meaning that PPE bears a positive charge below pH 8.31 and a negative charge above pH 8.31 (b). For source refer to reference list under the headings: 3est and pI calculator (chapter 7).

Successfully, the pI value of PPE was determined using a pI calculator in which figure 25

depicts the total net charge of the primary sequence, 3est for elastase i.e. PPE (EC

3.4.1.36) over the pH range of 1 – 14. From figure 25, the pI of PPE was predicted as

8.31; this meant that PPE bears a positive charge below pH 8.31 and a negative charge

above pH 8.31 as summarised by figure 25b. Thermolysin was used as a control enzyme

in this chapter and in the literature the isoelectric point (pI) of thermolysin was found to

be 4.97 (Miki et al. 1996). Thermolysin bears a positive charge below pH 4.97 and a

negative charge above pH 4.97. In this thesis, the pH of sample fluids were examined in

the range of pH 6.0 – 9.0 meaning that PPE was studied both in its cationic/ anionic

forms, whereas thermolysin was only studied in its anionic form. For the remaining

VVGGTEAQRNSWPSQISLQYRSGSSWAHTCGGTLIRQNWVMTAAHCVDRELTFRVVVGEHNLNQNNGTEQYVGVQKIVVHPYWNTDDVAAGYDIALLRLAQSVTLNSYVQLGVLPRAGTILANNSPCYITGWGLTRTNGQLAQTLQQAYLPTVDYAICSSSSYWGSTVKNSMVCAGGDGVRS


114

parts of this thesis, PPE will be referred to as ‘elastase’ and only when necessary it will be

indicated as PPE

3.4.4 Designing of the elastase responsive ECPs coupled to PEGA

(a)

(b)

Figure 26. Schematic illustration of designing elastase responsive ECPs that accommodate the environment of chronic wounds both above and below pH 8.31. (a) Elastase has a pI value of 8.31 consequently within the pH range of chronic wounds (pH 5.45 – 8.65) it is as a positive protease below pH 8.31 and a negative protease above pH 8.31. (b) In this manner, depending on the pH of chronic wounds PEGA particles were functionalised with charged ECPs that were of opposite charge to elastase. The ECPs consist of two sections: a charge modified residue (CMR: positive, negative or neutral) and an enzyme recognition sequence (ERS: P1-P1′ scissile bond). Since the CMR controls the molecular accessibility of elastase into PEGA particles, a negative CMR is suitable when elastase is positive (pH < 8.31); and a positive CMR is suitable when elastase is negative (pH > 8.31) in order to remove elastase from sample fluids mimicking chronic wound fluids because opposite charges are required for attraction and entrapment of elastase into PEGA particles. Other abbreviations: pI (iso-electric point); positively charged elastase/ CMR (red); negatively charged elastase/ CMR (blue); P1-P1′ (scissile bond), Fmoc (fluorenylmethoxycarbonyl) and side chains of amino acids (R1-3).

Enzyme cleavable peptides (ECPs) were coupled to PEGA particles via SPPS and

Fmoc/DIC chemistry (see section 3.4.2.2). Fmoc-amino acids have previously been

reported to have anti-inflammatory properties (Burch et al. 1991) making them

pH > 8.31

CMR ERS

FmocNH

NHNH

NH

O

O

OR2

R1R3

P1 P1’

P1 P1’

(Charged Modified

Residue)

(Enzyme Recognition

Sequence)

PEGA

particles

pH < 8.31

pI elastase

pH range chronic wounds

3 4 5 6 8 9 7 10

pH

+ve –ve


115

potentially interesting for designing ECPs in the context of generating a hydrogel wound

dressing.

The ECPs consisted of two sections: a charged modified residue (CMR) and an enzyme

recognition sequence (ERS) as schematically illustrated by figure 26. The CMR behaves

as the actuator and its function was to control the attraction and hence the accessibility

of the target protease into the interior of the PEGA particles depending on the ionic

strength and pH of sample fluids. The CMR attracts a target protease of opposite charge

(depending on its pI value) to its own charge. In terms of ionic strength, both the

positive and negative CMRs (achieved by Fmoc-Arg(+) and Fmoc-Glu(-) respectively)

increase the swelling hence the molecular accessibility into PEGA particles compared to

the neutral CMR (i.e. Fmoc-Gly) as discussed below (i.e. selective hydrolysis of ECPs by

proteases including swelling analysis). The ERS behaves as the cleavable substrate

containing the P1-P1′ scissile bond (figure 26b) that is specifically recognised and

selectively cleaved by the target protease.

Table 13. ECPs coupled to PEGA particles.

ECP Entry CMR ERS

Length of ECP

Overall charge

Abbreviations

1 n/a Ala – Ala Ala – Ala

2 n/a Gly – Gly Di-peptide Uncharged Gly – Gly

3 n/a Phe – Phe Phe – Phe

4 Arg(+) Positive RAA

5 Glu(–) Ala – Ala Negative EAA

6 Gly Tri-peptide Neutral GAA

7 Arg(+) Positive RFF

8 Glu(–) Phe – Phe Negative EFF

9 Gly Neutral GFF

10 Arg(+) Positive RA

11 Glu(–) Ala Di-peptide Negative EA

12 Gly Neutral GA

Other abbreviations: ECP (enzyme cleavable peptides); CMR (charge modified residue); ERS (enzyme-recognition sequence); n/a (not applicable). Entries 1-3 and 10 – 12 (Fmoc-dipeptide-PEGA particles); and entries 4 – 9 (Fmoc-tripeptide-PEGA particles).


116

As encountered in chapter 2, chronic wounds are said to have a pH within the range of

5.45 – 8.65 (Dissemond et al. 2003) and previously we encountered that elastase

(specifically PPE) has a pI value of 8.31; in this manner, ionic ECPs of both positive and

negative charges can be considered to accommodate a hydrogel dressing that would be

suitable for the entire pH range of chronic wounds in response to both the cationic/

anionic forms of elastase which is schematically summarised by figure 26. A list of all the

ECPs used to functionalise PEGA particles within this thesis are tabulated in table 13.

From table 13: the uncharged Fmoc-dipeptide-PEGA particles (entries 1-3) were utilised

to determine the ERS in response to elastase; the Fmoc-tripeptide-PEGA particles with

the Fmoc-X-Ala-Ala-PEGA configuration (entries 4 – 6) and the Fmoc-dipeptide-PEGA

particles with the Fmoc-X-Ala-PEGA configuration (entries 10 – 12) were used to

determine the bond specificities of thermolysin and elastase by the selective preference

of cleaving ECPs under the influence of charge depending on pH. The Fmoc-tripeptide-

PEGA particles with the configurations: Fmoc-X-Ala-Ala-PEGA (entries 4 – 6) and

Fmoc-X-Phe-Phe-PEGA (entries 7 – 9) were exploited to study the swelling behaviour

of these particles under the influence of pH and ionic strength. The X residue within the

following configurations: Fmoc-X-Ala-Ala-PEGA, Fmoc-X-Phe-Phe-PEGA and Fmoc-

X-Ala-PEGA (entries 4 – 12) is the CMR. As it can be seen from table 13, the ionic

CMR consisted of two different amino acids: an arginine residue (Arg(+)) or a glutamic

acid residue (Glu(–)) to respectively maintain an overall positive (table 13: entries 4, 7 and

10) or negative charge (table 13: entries 5, 8 and 11) of the ECP coupled to PEGA

particles. In contrast, the neutral PEGA particles contained an uncharged CMR which

was maintained by a glycine residue (Gly; table 13: entries 6, 9 and 12). For simplicity, all

the functionalised PEGA particles (including their ECPs) studied within this thesis are

abbreviated according to the ECPs coupled to PEGA particles as given in table 13 (i.e.

the last column on the right hand side).

3.4.4.1 Selective design of the ERS responsive to elastase

In the literature, the specificity of elastase has been reported to cleave substrates

containing small, neutral and hydrophobic amino acid residues such as alanine and

glycine in the P1 position (Stryer 1995; Purich and Allison 2002) with the preference of

cleaving the di-alanine bond (Ala-Ala) more readily and does not recognise or cleave


117

bulky hydrophobic amino acid residues (Stryer 1995; Copeland 2000; Purich and Allison

2002; Barrett et al. 2004). In order to design the ERS for elastase and to test the bond

specificity of the ECP in response to elastase, initial studies involved coupling three

simple non-polar dipeptides to PEGA800 particles: Ala-Ala and Gly-Gly (test ECPs) and

Phe-Phe (control ECPs).

Figure 27 shows that elastase (1 mg/ml in water) significantly and selectivity cleaved the

Ala-Ala bond of Fmoc-Ala-Ala-PEGA800 particles (89.91%) more efficiently compared to

the Gly-Gly bond (1.97%) of Fmoc-Gly-Gly-PEGA800 particles (p > 0.05); and as

expected elastase did not recognise nor cleave the bulky aromatic Phe-Phe bond within

Fmoc-Phe-Phe-PEGA800 particles. The neutral hydrophobic Ala-Ala bond was selected

as the ERS (P1-P1′) for elastase.

0

20

40

60

80

100

Ala-Ala Gly-Gly Phe-Phe

Cleaved product (%

)

Figure 27. Selective hydrolysis of neutral ECPs (Fmoc-dipeptides) coupled to PEGA800 particles by elastase. Three configuration of hydrophobic Fmoc-dipeptide ECPs: Fmoc-Ala-Ala and Fmoc-Gly-Gly (test samples) and Fmoc-Phe-Phe (control sample) were coupled to PEGA800 particles. These were then cleaved with elastase (PPE, 1 mg/ml in dH2O, pH 7.2) overnight at ambient temperature on a blood rotator. The cleaved products were collected and analysed by rpHPLC. Elastase selectively hydrolysed the Fmoc-Ala-Ala-PEGA particles more efficiently in contrast to Fmoc-Gly-Gly-PEGA and Fmoc-Phe-Phe-PEGA particles (p < 0.05). Data represents the mean + SE of two measurements (n=2).

3.4.4.2 Selective hydrolysis of Fmoc-tripeptide-PEGA particles by proteases

To accommodate the pH of chronic wounds (pH 5.45 – 8.65; Dissemond et al. 2003) the

pH range in this section was studied between pH 6.0 – 9.0; and figure 28 schematically

compares the charges of both elastase and thermolysin within this pH range as previously

we encountered that proteases vary in charge depending on their pI value hence pH of


118

the surrounding media. As it can be seen elastase was studied both in its cationic

(positive) and anionic (negative) forms, whereas thermolysin was studied only in its

anionic form.

Figure 28. Schematic illustration comparing the charges of thermolysin and elasatse within the pH range of 6.0 – 9.0 (indicated by purple). Within this pH range elastase (pI 8.31) exists both in its cationic form (pH < 8.31; +ve protease indicated by blue) and anionic form (pH > 8.31; –ve protease indicated by red); whereas the acidic thermolysin (pI 4.97) always existed in its anionic form (pH > 4.97; –ve protease indicated by red).

In terms of charge (i.e. protease and ECPs) and under the influence of pH, it was

hypothesised that both proteases (thermolysin and elastase) should access PEGA

particles and thus readily cleave ECPs of opposite charge to that of the protease.

Therefore, in this section the terminal end of the ERS (Ala-Ala bond) was then flanked

by a single CMR to generate functionalised Fmoc-tripeptide-PEGA particles with the

configuration Fmoc-X-Ala-Ala-PEGA (as summarised in table 13: entries 4 - 6).

O NN

N

O R3

O R2

ON

R1

OO N

NOH

O R3

O R2

ON

R1

ONH2+

Protease

HPLC

Fmoc-tripeptide-PEGA particles Cleaved product

pH 6.0 - 9.0

0.001M

Figure 29. A chemical reaction displaying the selective hydrolysis of Fmoc-tripeptide-PEGA particles by a protease. The Ala-Ala bond of Fmoc-tripeptide-PEGA particle (RAA: positive; EAA: negative; and GAA: neutral) was selectively cleaved by thermolysin (control protease) and elatase (test protease) at 0.001M and within the pH range of 6.0 – 9.0. The total cleaved products achieved enzymatic treatment were collected and analysed by rpHPLC.

As a starting point, the ionic strength of potassium phosphate buffer was selected at low

ionic strength i.e. 0.001 M and after synthesising the Fmoc-tripeptide-PEGA particles

(entries 4 – 6, table 13) they were thoroughly washed and then swollen in 0.001 M

potassium phosphate buffer at different pH values ranging from pH 6.0 – 9.0. Next, all

the swollen Fmoc-tripeptide-PEGA particles (i.e. RAA, EAA, GAA) were incubated

CH3

Ala

N N

N+

Arg

-O

O

Glu

H

Gly

R1, R2: R3:

pI elastase

pH range chronic wounds

3 4 5 6 8 9 7 10

pH

+ve –ve

+ve –ve

pI thermolysin


119

with thermolysin and elastase (0.1 mg/ml) in potassium phosphate buffer (0.001M) at the

appropriate pH (as indicated). The total cleaved product(s) for each functionalised

PEGA particles were collected and analysed by rpHPLC (as schematically summarised by

figure 29).

(a) (b)

0

20

40

60

80

100

RAA EAA GAA

PEGA800

Cleaved product (%)

0

20

40

60

80

100

RAA EAA GAA

PEGA1900

Cleaved product (%

)

(c) (d)

0

20

40

60

80

100

RAA EAA GAA

PEGA800

Cleaved product (%

)

0

20

40

60

80

100

RAA EAA GAA

PEGA1900

Cleaved product (%

)

pH 6.0 pH 6.5 pH 7.0 pH 7.5 pH 8.0 pH 8.5 pH 9.0

Figure 30. Selective hydrolysis of Fmoc-X-Ala-Ala-PEGA(800,1900) particles by thermolysin and elastase under the influence of pH. PEGA800 (a, c) and PEGA1900 (b, d) particles were functionalised with positive (RAA), negative (EAA) and neutral (GAA) ECPs to generate Fmoc-X-Ala-Ala-PEGA particles (where X (CMR): Arg(+); Glu(-); Gly). These particles were treated with 0.1 mg/ml of both thermolysin (a, b) and elastase (c, d) within the pH range of 6.0 – 9.0 in 0.001M potassium phosphate buffer, overnight on a blood rotator at 34oC. The cleaved products were collected and analysed by rpHPLC. Under the influence of pH, thermolysin bears a negative charge (pH > 4.97) and had a high preference to cleave RAA more compared to EAA and GAA for both PEGA800 (a) and PEGA1900 (b); whereas elastase selectively cleaved EAA more compared to RAA and GAA for both PEGA800 (c) and PEGA1900 (d). Interestingly, for the negative ECP (EAA): elastase activity increased when elastase was positively charged between pH 6.0 – 8.0; on the contrary when elastase became negative (pH > 8.31) a decrease in elastase activity was observed for EAA of both PEGA800 and PEGA1900 particles (c-d, respectively) and an increase in elastase activity was observed for RAA (PEGA1900 particles, d). Statistical analysis was conducted using ANOVA and pair-wise comparisons were conducted by post hoc: Tukey and Duncan test: a significant difference observed at the 95% confidence level (p > 0.05). Data represents the mean + SE of four measurements (n=4).

0

5

10

15

20

RAA EAA GAACleaved product (%

)


120

Figure 30 demonstrates that both thermolysin and elastase were able to access into

Fmoc-tripeptide-PEGA particles (i.e. Fmoc-X-Ala-Ala-PEGA) and cleave all ECPs

(RAA, EAA and GAA) for both PEGA800 and PEGA1900 particles. Overall, it was found

that thermolysin cleaved all ECPs to a high degree (i.e. PEGA800 particles: 41.00 –

89.52%; and PEGA1900 particles: 38.65 – 98.53 %) compared to elastase (PEGA800

particles: 1.86 – 60.68 %; and PEGA1900 particles: 2.87 – 52.28 %) which is expected as

previously Morihara and co-workers reported thermolysin to be more active than elastase

(Morihara and Tsuzuki 1967).

Initially, the selective hydrolysis of Fmoc-tripeptide-PEGA particles with thermolysin

will be considered followed by elastase. Since thermolysin was studied in its anionic

form (pH > 4.97) the expected observations were observed for both PEGA800 and

PEGA1900 particles (figure 30a and figure 30b, respectively) in which the specificity of

thermolysin was found to selectively hydrolyse the positive ECP (RAA) significantly

more for both PEGA800 (figure 33a) and PEGA1900 (figure 33b) compared to both the

negative (EAA) and neutral (GAA) ECPs for both PEGA800 and PEGA1900 (p < 0.05).

Additionally, there was no significance difference in the specificity of thermolysin

between the negative (EAA) and neutral (GAA) ECPs for both PEGA800 (figure 30a) and

PEGA1900 (figure 30b) particles (p > 0.05) this is because the Ala residue of both EAA

and GAA is located in the P1–P1′ position of Fmoc-X-Ala-Ala-PEGA particles and it is

well-known that the specificity of thermolysin has preference to cleave hydrophobic

residues in the P1 or P1′ positions such as Ala (Barrett et al. 2004).

Next the specificity of thermolysin to hydrolyse each ECP was compared under the

influence of pH. Thermolysin activity was found to be independent of pH (range 6.0 –

9.0) for both the positive ECP (RAA: PEGA800 and PEGA1900 particles, p > 0.05) and

negative ECP (EAA: PEGA1900 particles, p > 0.05). For the remaining ECPs (EAA:

PEGA800 particles; and GAA: PEGA800 and PEGA1900 particles) it can be said that

thermolysin activity was partially independent on pH. This is because after pair-wise

comparisons using the ANOVA test, the post hoc (Tukey) test revealed that for the

negative ECP (EAA: PEGA800 particles) thermolysin activity was not significantly

different between: (a) pH 6.0 and pH 7.0 (p > 0.05); and (b) 6.5, 7.5, 8.5 and pH 9.0 (p >

0.05); but was significantly high at pH 8.0 compared to remaining pH values (p < 0.05).

In the same way, for the neutral ECP (GAA) thermolysin activity was not significantly


121

different between (a) pH 6.0 – 8.0 (p > 0.05) and (b) pH 8.5 – 9.0 (p > 0.05) for

PEGA800 particles (figure 30a); whereas thermolysin activity for PEGA1900 particles was

not significantly different between (a) pH 6.0 – 8.5 (p > 0.05) and (b) pH 6.5 – 9.0 (p >

0.05; figure 30b). This is due to the Ala residues of the ERS because it has previously

been reported that the specificity of thermolysin is independent of pH when an

hydrophobic Ala residue is located either in the P1 or P2 positions of a given substrate

(Murakami et al. 2001) such as Fmoc-X-Ala-Ala which is the configuration of all ECPs

(RAA, EAA, GAA) within this section.

Finally, the yields of the cleaved products were not significantly different between

PEGA800 and PEGA1900 particles for each ECP (RAA, EAA and GAA) which shows that

the accessibility of thermolysin into PEGA800 and PEGA1900 particles was significantly

similar for each type of Fmoc-tripeptide-PEGA particles i.e. RAA, EAA, GAA (p >

0.05).

Next, the effect of elastase was studied for both PEGA800 and PEGA1900 particles (figure

30c and figure 30d, respectively) and elastase had a high specificity to selectively

hydrolyse the negatively charged ECP (EAA: PEGA800 7.65 – 60.68 % and PEGA1900

7.39 – 52.50 %) significantly more compared to both the positive ECP (RAA: PEGA800

2.36 – 5.36 % and PEGA1900 2.87 – 14.27 %) and the neutral ECP (GAA: PEGA800 1.86

– 11.46 % and PEGA1900 2.77 – 6.07 %) over the pH range of 6.0 – 9.0 (p < 0.05).

Previously, another study reported that human elastase (i.e. NE) had a high affinity for

negatively charged substrates because the surface of NE contains Arg residues on one

side of its surface (Watorek et al. 1991). As previously mentioned, PPE has 40%

homology to NE and correspondingly the native primary sequence of PPE has been

reported to have 22 nominal basic sites constituting of 12 Arg, 6 His, 3 Lys residues

including the N-terminus (Hogan and McLuckey 2003). Interestingly, under the

influence of pH this means that Glu(-) residue of the negative ECP for EAA particles

would ion pair with the amine of the Arg, His and Lys side chains on the surface of

elastase i.e. PPE in its cationic form (pH < 8.31). Likewise, the native primary sequence

of PPE contains 12 negative sites comprising of 8 Asp and 3 Glu residues including the

C-terminus (Hogan and McLuckey 2003) therefore the Arg(+) residue of the positive ECP

for RAA particles would ion pair with the carboxyl groups of both the Asp and Glu side

chains on the surface of elastase i.e. PPE in its anionic form (pH < 8.31).


122

Consequently, when comparing the pH activity of elastase to hydrolyse each ECP, the

specificity of elastase was found to be dependant on pH for the charged ECPs (i.e. EAA:

PEGA800 and PEGA1900 particles; and RAA: PEGA1900 particles only). Here, as expected

the specificity of elastase in its cationic form (pH < 8.31) selectively cleaved the negative

ECP (EAA) significantly more for both PEGA800 (i.e. 7.65 – 60.69 %, figure 30c) and

PEGA1900 (i.e. 45.84 – 52.28%, figure 30d) compared to the positive ECP (RAA) for

both PEGA800 (i.e. 7.65 – 60.69 %, figure 30c) and PEGA1900 (i.e. 45.84% – 52.28%,

figure 30d) particles within the pH range of 6.0 – 7.5 (p < 0.05). Quite the opposite,

when elastase was in its anionic form (pH > 8.31), elastase activity was found to decrease

from pH 8.5 – 9.0 for negative ECP (EAA: PEGA800 i.e. 23.95% – 20.74 %, figure 30c

and PEGA1900 i.e. 9.44 – 7.39, figure 30d) whereas for the positive ECP (RAA) it was

found to increase from pH 8.5 – 9.0 for PEGA1900 particles (i.e. 11.28% – 14.27%). In

contrast, elastase activity was significantly independent of pH for the neutral ECP (GAA)

for both PEGA800 and PEGA1900 particles; including the positive ECP (RAA) only for

PEGA800 particles (p > 0.05; figure 30c-d). This shows that the molecular accessibility of

elastase into positive PEGA particles (RAA) was more suitable for PEGA1900 particles

compared to PEGA800 particles.

3.4.4.3 Selective hydrolysis of the CMR from Fmoc-tripeptide-PEGA particles by proteases

When the HPLC separation of the total cleaved products for Fmoc-tripeptide-PEGA

particles as studied in section 3.4.4.3 were carefully looked at, it was found that both

thermolysin and elastase had the ability to cleave at the carboxyl end of the CMR for all

Fmoc-tripeptide ECPs i.e. Fmoc-X~Ala-Ala (Note: ~ indicates the cleaving site; R~AA,

E~AA, G~AA) and the data is summarised by figure 31. Figure 31, shows that

thermolysin had the ability to selectively cleave the Glu(-) and Gly CMRs more compared

to the Arg(+) CMR for both PEGA800 and PEGA1900 particles. In contrast, elastase was

found to selectively cleave the Glu(-) CMR more compared to both the Arg(+) and Gly

CMRs.

Just like the previous section (3.4.43), the specificity of thermolysin to cleave the CMR

was found to be independent of pH. Over the pH range of 6.0 – 9.0, thermolysin was

found to cleave the: Arg(+) residue by 0.39 – 0.69 % (PEGA800) and 18.03 – 25.82 %

(PEGA1900); Glu(-) residue by 21.70 – 54.84 % (PEGA800) and 37.96 – 58.36 %


123

(PEGA1900); and Gly residue by 40.23 – 74.09 % (PEGA800) and 38.34 – 48.80 %

(PEGA1900). When the cleaved product of the CMR for each ECP (figure 31) was

compared with the total cleaved product of each ECP (figure 30) for both PEGA800 and

PEGA1900: the selective cleaving of the Arg(+) and Glu(–) residues from Fmoc-X~Ala-Ala

by thermolysin were significantly lower compared to the total cleaved product for RAA

and EAA (p < 0.05). In contrast, the selective cleaving of the Gly residue from Fmoc-

X~Ala-Ala by thermolysin was not significantly different to the total cleaved product for

GAA (p > 0.05).

(a) (b)

0

20

40

60

80

100

RAA EAA GAA

PEGA800

Cleaved product (%)

0

20

40

60

80

100

RAA EAA GAA

PEGA1900

Cleaved product (%

)

(c) (d)

0

20

40

60

80

100

RAA EAA GAA

PEGA800

Cleaved product (%

)

0

20

40

60

80

100

RAA EAA GAA

PEGA1900

Cleaved product (%

)

pH 6.0 pH 6.5 pH 7.0 pH 7.5 pH 8.0 pH 8.5 pH 9.0

Figure 31. Selective hydrolysis of the CMR from Fmoc-X-Ala-Ala-PEGA(800,1900) particles by thermolysin and elastase. PEGA800 (a, c) and PEGA1900 (b, d) particles were functionalised with Fmoc-X-Ala-Ala-PEGA particles (where X (CMR): Arg(+); Glu(-); Gly). Positive (RAA), negative (EAA) and neutral (GAA) particles were treated with 0.1 mg/ml of both thermolysin (a, b) and elastase (c, d) within the pH range of 6.0 – 9.0 in 0.001M potassium phosphate buffer, overnight on a blood rotator at 34oC. The separation of the CMR by rpHPLC shows that the specificity of both thermolysin and elastase had preference to cleave the Fmoc-tripeptide ECP at the carboxyl end of the CMR i.e. Fmoc-X~Ala-Ala. Data represents the mean + SE of four measurements (n=4).

0.0

0.2

0.4

0.6

0.8

1.0

RAA

Cleaved product (%

)

0.00

0.05

0.10

0.15

0.20

RAA

Cleaved product (%)

0.0

0.5

1.0

1.5

GAA

Cleaved product (%

)

0

2

4

6

8

10

RAA

Cleaved product (%)

0.0

0.5

1.0

1.5

2.0

2.5

GAA

Cleaved product (%)


124

In contrast, over the pH range of 6.0 – 9.0 elastase was found to selectively cleave the

Glu(–) residue (PEGA800: 4.81 – 45.65 %; PEGA1900: 0.20 – 45.37 %) more compared to

both the Arg(+) (PEGA800: 0.03 – 0.15 %; PEGA1900: 0.30 – 8.06 %) and Gly (PEGA800:

0.10 – 1.45 %; PEGA1900: 0.49 – 2.19 %) residues. When the cleaved product of the

CMR for each ECP (figure 31) was compared with the total cleaved product of each

ECP (figure 30) for both PEGA800 and PEGA1900: the selective cleaving of all three

CMRs (Arg(+), Glu(–) and Gly) from Fmoc-X~Ala-Ala by elastase were significantly lower

compared to the total cleaved products for RAA, EAA and GAA (p < 0.05).

Interestingly, just as we saw in figure 30, the pH activity for the specificity of elastase to

cleave the Glu(–) residue from EAA for both PEGA800 and PEGA1900 was found to be

dependant on pH. As expected, the pH profile for the selective hydrolysis of the Glu(–)

residue from EAA by the cationic form of elastase was found to increase (pH < 8.31)

and then elastase activity was found to decrease when elastase was in its anionic form

(pH 8.5 – 9.0) for both PEGA800 and PEGA1900.

3.4.4.4 Selective hydrolysis of Fmoc-dipeptide-PEGA particles by proteases The ECP bond specificities were further characterised for both thermolysin and elastase

in this section since both thermolysin and elastase had the ability to cleave at the carboxyl

end of the CMR for all Fmoc-tripeptide ECPs i.e. Fmoc-X~Ala-Ala (figure 31, in the

previous section 3.4.4.3). In view of that, it was envisaged that for commercial purposes

designing a simple wound dressing with a small ECP could potentially be beneficial as it

would be cheaper to synthesis.

Consequently, in this section the CMRs were flanked at the terminal end of a single Ala

residue coupled to PEGA particles (i.e. Ala-PEGA) to generate Fmoc-dipeptide-PEGA

particles with the configuration of Fmoc-X-Ala-PEGA (where X was again equivalent to

the CMR: Arg(+), Glu(–), or Gly; as summarised by figure 32) and both the ECPs and

PEGA particles in this section are abbreviated as: RA, EA and GA (as indicated in table

13: entries 10-12). Subsequently, the selective hydrolysis of the dipeptide ECPs for both

PEGA800 and PEGA1900 particles by both thermolysin and elastase was investigated

within the pH range of 7.5 – 9.0 (at 0.001M) and the collected cleaved products were

again separated and purified by rpHPLC (figure 32).


125

O NN

N

O R2

O R1

O

O NOH

O R2

O

NH2

NR1

O

+Protease

HPLC

Fmoc-dipeptide-PEGA particles Cleaved product

pH 7.5 - 9.0

0.001M

Figure 32. The chemical reaction of cleaving of Fmoc-dipeptide-PEGA particles (RA, EA, GA) by proteases. The X~Ala bond of the Fmoc-dipeptide-PEGA particle (RA: positive; EA: negative; and GA: neutral) was selectively cleaved by a protease such as thermolysin (control protease) or elatase (test protease) at 0.001M and within the pH range of 7.5 – 9.0. The total cleaved products achieved enzymatic treatment were collected and analysed by rpHPLC.

From figure 33, it can be seen that thermolysin and elastase were compatible in accessing

Fmoc-X~Ala-PEGA particles and had the ability to cleave all ECPs (RA, EA and GA)

for both PEGA800 and PEGA1900 particles. Again, thermolysin was found to cleave all

ECPs (RA, EA and GA) to a high degree (i.e. PEGA800 particles: 59.33 – 95.94 %; and

PEGA1900 particles: 65.89 – 97.50 %) compared to elastase (PEGA800 particles: 0.41 –

1.53 %; and PEGA1900 particles: 0.31 – 11.81 %) which again confirmed that thermolysin

was more selective than elastase (Morihara and Tsuzuki 1967).

For PEGA800 particles as expected thermolsyin selectively cleaved the positive ECPs

(RA: average 79.23 %) significantly more compared to the negative ECP (EA: average

62.89 %) (p < 0.05, figure 33a). Additionally, it can be seen that GA (neutral ECP) was

cleaved to a high degree (average: 85.16 %) compared to both RA and EA (figure 33a);

this is expected because as mentioned above the specificity of thermolysin has preference

to cleave hydrophobic residues such as Gly or Ala in both the P1 and P1′ positions

(Barrett et al. 2004). Although, on average thermolysin cleaved GA more than RA (i.e.

PEGA800 particles), statistically it was found that the specificity of thermolysin was

significantly similar for GA (76.58 – 95.94 %) and RA (72.93 – 89.64 %) because the

average percentage of the cleaved products for both GA (85.16 %) and RA (79.23 %)

were not significantly different (p > 0.05); whereas for EA the cleaved products were

significantly lower (i.e. 59.33 – 66.59 %) than both RA and GA (p < 0.05). For

PEGA1900 particles (figure 33b) no obvious trend was observed for thermolysin activity

between pH 7.5 – 9.0. Here, thermolysin hydrolysed all ECPs to a high degree: RA

(77.09 %), EA (88.40 %) and GA (86.84 %). When comparing the specificity of

thermolysin for all ECPs, it was found to be significantly different between RA and EA

CH3

Ala

N N

N+

Arg

-O

OR1 R2

Glu

H

Gly


126

(p < 0.05) including RA and GA (p < 0.05); whereas for EA and GA it was significantly

similar (p > 0.05).

(a) (b)

0

20

40

60

80

100

RA EA GAPEGA800

Cleaved product (%

)

0

20

40

60

80

100

RA EA GAPEGA1900

Cleaved product (%

)

(c) (d)

0.0

0.5

1.0

1.5

2.0

RA EA GA

PEGA800

Cleaved product (%

)

0.0

3.0

6.0

9.0

12.0

15.0

RA EA GA

PEGA1900

Cleaved product (%

)

pH 7.5 pH 8.0 pH 8.5 pH 9.0

Figure 33. Selective hydrolysis of Fmoc-X-Ala-PEGA(800,1900) particles by thermolysin and elastase at different pH values. PEGA800 (a, c) and PEGA1900 (b, d) particles were functionalised with Fmoc-X-Ala-PEGA particles (where X (CMR): Arg(+); Glu(-); Gly). Positive (RA), negative (EA) and neutral (GA) particles were treated with 0.1 mg/ml of both thermolysin (a, b) and elastase (c, d) within the pH range of 7.5 – 9.0 in 0.001M potassium phosphate buffer, overnight on a blood rotator at 34oC. The cleaved products were collected and analysed by HPLC. Data represents the mean + SE of four measurements (n=4).

Next, the pH activity of thermolysin to hydrolyse each ECP was compared and the

specificity of thermolysin was found to be independent of pH for majority of the ECPs

i.e. RA, EA, GA (i.e. PEGA800 particles, p > 0.05; figure 33a) including EA (PEGA1900

particles, p > 0.05; figure 33b). This is because in these particles, the CMR residues

(referred as X in Fmoc-X-Ala-PEGA particles) is located in the P1 position of the ECP

and in the Murakami et al. study it was also reported that when an Arg, Ala and Gly

0.0

0.2

0.4

0.6

0.8

1.0

RA EA GACleaved product (%

)


127

residues (alongside others) are at the P1 and P2 positions then the specificity of

thermolysin is independent of pH in the order of L-Arg > L-Ala> L-Gly (Murakami et

al. 2001). However, the specificity of thermolysin was found to be dependant on pH for

RA and GA (PEGA1900 particles: figure 33b). After pair-wise comparisons, both post-

hoc tests (Tukey and Duncan) revealed that for RA: thermolysin activity was significantly

different only for pH 7.5 and pH 9.0, and was not significantly different between (a) pH

7.5 – 8.5; and (b) pH 8.0 – 9.0 (p > 0.05). In contrast, for GA thermolysin activity was

significantly different only between pH 7.5 and pH 8.5, and was not significantly

different between: (a) pH: 7.5, 8.0 and 9.0 (p > 0.05); and (b) pH 8.0 – 9.0 (p > 0.05). In

view of that, there is no conclusive trend as to why thermolysin activity depended on pH

for both RA and GA since majority of the pH values show that the specificity of

thermolysin was independent of pH for PEGA1900 particles (figure 33b).

The discussion will now focus on the selective hydrolysis of all the ECPs for Fmoc-X-

Ala-PEGA particles by elastase (figure 33c-d). Although the yields of the cleaved

product collected after treatment with elastase were low in comparison to thermolysin;

interestingly on the basis of charge and under the influence of pH the expected

observations were observed for PEGA800 particles (figure 33c). On average, elastase

cleaved the charged ECPs more (RA: 1.04% and EA: 1.00 %) compared to the neutral

ECPs (GA: 0.77%) between pH 7.5 – 9.0 (figure 33c) but the specificity of elastase for

cleaving these ECPs was significantly similar between pH 7.5 – 9.0 (p > 0.05). However,

for the charge ECPs (RA and EA) it was found that the specificity of elastase activity

significantly depended on pH (p < 0.05). As expected, when elastase was in its cationic

form (pI 8.31, pH < 8.31) that is at pH 7.5 – 8.0, it selectively cleaved the negative ECP

(EA: 1.35 % at pH 7.5; and 0.97 % at pH 8.0) more compared to the positive ECP (RA:

0.41 % – 0.80 % at pH 7.5 – 8.0). On the other hand, the opposite effect was observed

under the conditions when elastase was in its anionic form (pI 8.31, pH > 8.31) it

selectively cleaved the positive ECP (RA: 1.44 % – 1.53 % at pH 8.5 – 9.0) more

compared to the negative ECP (EA: 0.94 – 0.84 % at pH 8.5 – 9.0). From figure 33c it

can be seen that elastase also cleaved the neutral ECP (i.e. GA of PEGA800 particles)

which is not surprising because it is well-known that the specificity of elastase has

preference to cleave small hydrophobic residues such as Gly and Ala residues in the P1

and P1′ positions (Stryer 1995; Purich and Allison 2002).


128

For PEGA1900 particles (figure 33d), elastase selectively cleaved all ECPs more at pH 7.5;

and at this pH the negative ECP (EA: 11.80 %) was significantly cleaved more compared

to both positive (RA: 0.99 %) and neutral (GA: 1.90 %) ECPs which is expected because

at pH 7.5 elastase exists in its cationic form. Overall, for PEGA1900 particles the

selective specificity of elastase for cleaving Fmoc-X~Ala was independent of the charge

of ECP at high pH (p > 0.05), whereas at low pH (i.e. pH 7.5) the specificity of elastase

depended on the charge of ECP.

When comparing the specificity of both thermolysin and elastase for hydrolysing the

X~Ala bond (where X is the CMR) for both the Fmoc-dipeptide ECP (Fmoc-X~Ala

(figure 33) and the Fmoc-tripeptide ECP (Fmoc-X~Ala-Ala (figure 31), overall it was

found that:

• thermolysin had a high preference to hydrolyse the X~Ala bond of the Fmoc-

dipeptide ECP (for all ECPs: RA, EA, GA) for both PEGA800 and PEGA1900

particles; whereas

• elastase had a high preference to cleave the X~Ala bond of the Fmoc-tripeptide

ECP for all ECPs (RAA, EAA and GAA) meaning that the specificity of elastase

for X~Ala bond decreased for the Fmoc-dipeptide sequence i.e. Fmoc-X~Ala

for both PEGA800 and PEGA1900 particles.

This section (3.4.4.4) demonstrated that on the basis of charge and under the influence

of pH, the specificity of both thermolysin and elastase for preference of hydrolysing the

Fmoc-dipeptide ECP (Fmoc-X~Ala, where X was Arg(+), Glu(-) and Gly) was selectively

controlled by PEGA800 particles. For the remaining parts of this thesis, the

functionalised PEGA1900 particles with the configuration of Fmoc-X-Ala-Ala-PEGA

(where X was Arg(+), Glu(-) and Gly) were selected for further investigation having

established that elastase selectively cleaved the Fmoc-tripeptide ECP (i.e. Fmoc-X-Ala-

Ala; figure 30 in section 3.4.4.2) more compared to the Fmoc-dipeptide ECP (i.e. Fmoc-

X-Ala, figure 33 in section 3.4.4.4). This observation confirmed that the di-alanine bond

(Ala-Ala) was required to attain a good response for elastase activity, and this was

expected since the specificity of elastase (PPE) had preference to cleave at the carboxyl-

terminal side of small hydrophobic amino acid residues such as Ala (Stryer 1995; Barrett

et al. 2004). Additionally, high yields of the collected cleaved products for Fmoc-X-Ala-


129

Ala-PEGA in relation to elastase were obtained with PEGA1900 particles compared to

PEGA800 particles.

3.4.5 Swelling analysis

In this section potassium phosphate buffer was chosen as the swelling environment to

maintain both the internal and external pH and ionic strength of functionalised PEGA1900

particles in order to examine the electrostatic swelling behaviour of these PEGA

particles.

3.4.5.1 Swelling behaviour of un-cleaved Fmoc-tripeptide-PEGA particles

The average diameter of 100 particles was measured to study the swelling behaviour of

Fmoc-X-Ala-Ala-PEGA1900 and Fmoc-X-Phe-Phe-PEGA1900 as a function of different

pH and ionic strength. As previously mentioned, the CMR (again indicated as X within

this section) determines the overall charge of the ECP coupled to PEGA particles

producing either ionic (positive and negative) or neutral PEGA particles; as a reminder

the positive and negative charges were achieved by Fmoc-Arg(+) and Fmoc-Glu(–)

respectively; and the neutral charge was maintained by Fmoc-Gly (see section 3.4.4). As

the CMR is the actuator within these functionalised PEGA particles, in this section it will

become evident that the CMR plays an important role in modifying the swelling of the

hydrogel PEGA particles as the CMR can be susceptible to ionisation under aqueous

conditions.

The pH was chosen in the range of 7.0 – 9.0 to accommodate firstly the pH observed in

chronic wounds, and secondly within this pH range elastase (pI 8.31) can exist as both

cationic and anionic protease (see section 3.4.3). While, the ionic strength of the buffer

ranged from 0.001 – 0.2 M, in order to compare the swelling ability of functionalised

PEGA1900 particles at both low and high ionic strengths. The swelling for Fmoc-X-Phe-

Phe-PEGA1900 particles was studied within this chapter because in chapter 4 these

particles were chosen as a control for studying the effect of elastase cleaving at high ionic

strength.


130

Statistical analysis was performed in order to make comparisons between the swelling of

all types (charged and neutral) of functionalised PEGA1900 particles using ANOVA test

which included post-hoc tests (Tukey and Duncan) to determine which diameters differ

significantly from one another followed by a paired t-test. The statistical analysis was

performed at the 95% confidence level, and the effect of swelling was significant when

p< 0.05; whereas when p> 0.05 it was not significant.

The degree of swelling for Fmoc-X-Ala-Ala-PEGA1900 and Fmoc-X-Phe-Phe-PEGA1900

are displayed by figures 34 and 35, respectively. Within this section the Fmoc-X-Ala-Ala-

PEGA1900 and Fmoc-X-Phe-Phe-PEGA1900 particles are abbreviated again according to

table 13 (entries 4 – 9). From figure 34, the effect of swelling for Fmoc-X-Ala-Ala-

PEGA1900 particles was significantly higher for the charged particles (RAA and EAA; p <

0.05) compared to the neutral PEGA particles i.e. on average the diameter of the positive

particles (RAA) increased by 11.00 % and the negative particles (EAA) increased by

8.41% compared to the neutral particles (GAA). The same trend was observed for the

Fmoc-X-Phe-Phe-PEGA1900 particles (figure 35); in which the average diameters of both

the positive particles (RFF) and negative particles (EFF) significantly increased by 6.60 %

and 4.39 % respectively, compared to the neutral particles (GFF) (p < 0.05).

In terms of pH, the effect of swelling for the charged particles varied according to the

pKa of the ionisable side chain of the CMR, in this case the Arg residue for the positive

particles (RAA and RFF) and the Glu residue for the negative particles (EAA and EFF).

The Arg residue for the positive particles were protonated (i.e. C=NH2+) because the

pH of the surrounding buffer (i.e. pH 7 – 9) was below pKa 12.48 of the Arg side chain

causing the particles to swell as a result of electrostatic repulsion between positively

charged groups. Similarly, the Glu residues for the negative particles were carboxlayted

(COO–) because the pH of the surrounding buffer (i.e. pH 7 – 9) was above pKa 4.25 of

the Glu side chain causing the negative particles swell due to electrostatic interactions

between the anions groups. Overall, the effect of swelling for each of the charged

particles (i.e. RAA, EAA, RFF and EFF) did not depend on the pH because the average

diameter for each particle was not significantly different between pH 7 – 9 (p > 0.05;

figures 34 and 35).


131

(a) pH 7.0

0.245

0.270

0.295

0.320

0.345

0.370

0.001 M 0.01 M 0.1 M 0.2 M

Diameter (mm)

(b) pH 8.0

0.245

0.270

0.295

0.320

0.345

0.370

0.001 M 0.01 M 0.1 M 0.2 M

Diameter (m

m)

(c) pH 9.0

0.245

0.270

0.295

0.320

0.345

0.370

0.001 M 0.01 M 0.1 M 0.2 M

Diameter (mm)

Fmoc-Arg(+)-Ala-Ala Fmoc-Glu(-)-Ala-Ala Fmoc-Gly-Ala-Ala

Figure 34. Swelling behaviour of functionalised Fmoc-X- Ala-Ala-PEGA1900 particles as a function of pH (a: 7.0, b: 8.0 and c: 9.0) and ionic strength (ranging from 0.001M – 0.2M) in potassium phosphate buffer. The effect of swelling for the positive (Fmoc-Arg(+)-Ala-Ala), negative (Fmoc-Glu(–)-Ala-Ala) and neutral (Fmoc-Gly-Ala-Ala) PEGA1900 particles were statistically compared using ANOVA (including post-hoc tests) followed by a paired t-test at the 95% confidence level. For detail descriptions of these comparisons see the text within this section. Data represents the mean + SE of 100 measurements (n=100).


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(a) pH 7.0

0.245

0.270

0.295

0.320

0.345

0.370

0.001 M 0.01 M 0.1 M 0.2 M

Diameter (mm)

(b) pH 8.0

0.245

0.270

0.295

0.320

0.345

0.370

0.001 M 0.01 M 0.1 M 0.2 M

Diameter (mm)

(c) pH 9.0

0.245

0.270

0.295

0.320

0.345

0.370

0.001 M 0.01 M 0.1 M 0.2 M

Diameter (mm)

Fmoc-Arg(+)-Phe-Phe Fmoc-Glu(-)-Phe-Phe Fmoc-Gly-Phe-Phe

Figure 35. Swelling behaviour of functionalised Fmoc-X-Phe-Phe-PEGA1900 particles as a function of pH (a: 7.0, b: 8.0 and c: 9.0) and ionic strength (ranging from 0.001M – 0.2M) in potassium phosphate buffer. The effect of swelling for the positive (Fmoc-Arg(+)-Phe-Phe), negative (Fmoc-Glu(–)-Phe-Phe) and neutral (Fmoc-Gly-Phe-Phe) PEGA1900 particles were statistically compared using ANOVA (including post-hoc tests) followed by a paired t-test at the 95% confidence level. For detail descriptions of these comparisons see the text within this section. Data represents the mean + SE of 100 measurements (n=100).


133

On the contrary, the swelling of the charged particles (RAA, EAA, RFF, EFF)

significantly depended on ionic strength (p < 0.05; figures 34 and 35). Principally, the

electrostatic interactions for both the positive (RAA and RFF) and negative (EAA and

EFF) particles decreased with increasing ionic strength (0.001 – 0.2 M) because the

counter ions of the buffer screened the electrostatic interactions of the CMR for each

ECP, causing the swelling to significantly decrease with increasing ionic strength (0.001 –

0.2 M, p< 0.05); whereas at low ionic strength the electrostatic interactions of the CMR

for all ECPs increased causing the swelling to significantly increase (p < 0.05).

For neutral particles (GAA and GFF: figures 34 and 35 respectively) as there are no

ionisable groups, the effect of swelling remained the same for all tested pH values (pH 7,

8 and 9, p> 0.05); and similarly no measureable effect of swelling was observed

depending on the ionic strength within the range of 0.001 – 0.1 M (p > 0.05) and these

observations confirm with the trends observed by Basso and co-workers for PEGA

particles possessing an overall neutral charge (Basso et al. 2004). The swelling was

significantly lower at 0.2 M (p < 0.05) which was unexpected.

Next, the swelling of Fmoc-X-Ala-Ala-PEGA1900 particles were compared with each

other. The swelling for RAA was significantly different when compared with EAA (p <

0.05) and GAA (p < 0.05) for all ionic strength and pH values. Additionally, the swelling

of EAA and GAA was significantly different at low ionic strength (0.001 – 0.01 M, at all

tested pH values; p < 0.05) whereas at high ionic strength and pH (i.e. 0.1 – 0.2 M and

pH 8 – 9) the effect of swelling for EAA was similar to GAA (p > 0.05). For the Fmoc-

X-Phe-Phe-PEGA1900 particles, the swelling for RFF was significantly similar to EFF at

low ionic strength (0.001 – 0.1 M, at pH 7 – 9; p > 0.05) whereas at high ionic strength

(0.2 M, at pH 7 – 9) it was significantly different (p < 0.05) in which the swelling of the

positive particles (RFF) was more than negative particles (EFF). As expected, the

swelling of RFF against GFF was found to be significantly different at both low ionic

strength (0.001 – 0.01 M, at pH 7 – 9; p < 0.05) and high ionic strength (0.2 M, pH 7 – 9:

p < 0.05) in each case: RFF swelled more than GFF; whereas at 0.1 M (pH 7 – 9) the

swelling for RFF was unexpectedly similar to GFF (p > 0.05). Similarly, as expected the

swelling of EFF against GFF was significantly different at low ionic strength (0.001 –

0.01 M, at pH 7 – 9; p < 0.05) i.e. EFF swelled more than GFF; whereas at high ionic

strength (0.1 – 0.2 M, at pH 7 – 9) the swelling of EFF and GFF was significantly similar


134

(p < 0.05). Overall, it was expected at high ionic strength that the charge particles (RAA,

EAA, RFF, EFF) swelled the same as neutral particles (GAA and GFF) because as

previously mention with increasing ionic strength the counter ions of the buffer increase

which then shield the ionic charges of ECPs and accordingly swelling decreases for

charge particles (RAA, EAA, RFF, EFF), whereas at low ionic strength the counter ions

of the buffer are low in this manner there are more repulsions between the electrostatic

interactions and consequently swelling increases.

Finally, the swelling of Fmoc-X-Ala-Ala-PEGA1900 particles (figure 34) were compared

with Fmoc-X-Phe-Phe-PEGA1900 particles (figure 35). The swelling for Fmoc-X-Phe-

Phe-PEGA1900 particles was significantly lower than Fmoc-X-Ala-Ala-PEGA1900 particles

(p < 0.05) this is because the ECP of the Fmoc-X-Phe-Phe-PEGA1900 particles contain

Phe residues which are more hydrophobic than the Ala residues. The decrease in

swelling by the Fmoc-X-Phe-Phe-PEGA1900 particles is due to ‘hydrophobic effect’

wherein the Phe residues tend to cluster/ pack themselves together in aqueous

environments and therefore prevent water/ hydrophilic molecules to pass between them.

For the reason that, Fmoc-X-Phe-Phe-PEGA1900 particles will let less water in compared

to the Fmoc-X-Ala-Ala-PEGA1900 particles this is the possible reason why the Fmoc-X-

Ala-Ala-PEGA1900 particles are found to swell more compared to the Fmoc-X-Phe-Phe-

PEGA1900 particles. The effect of swelling between the negative particles (EAA vs EFF)

was significantly different at all the tested pH and ionic strength (p < 0.05) which shows

that the swelling of EAA particles to be significantly higher than EFF (p < 0.05; figure

34 and 35). Whereas for the positive particles (RAA vs RFF) the effect of swelling was

not significantly different at low ionic strength (0.001 M) for all the tested pH values (p >

0.05; figure 34 and 35), but was significantly different within the range of 0.01 – 0.2 M

for all pH values (p < 0.05; figure 34 and 35). As expected for the neutral particles

(GAA vs GFF) because there are no ionisable groups the effect of swelling was not

significantly different at low ionic strength: 0.001 – 0.01 M at pH 7, 8 and 9 (p > 0.05);

whereas the swelling was significantly lower for GFF compared to GAA at high ionic

strength i.e. 0.1 – 0.2 M at pH 7 , 8 and 9 (p < 0.05; figure 34 and 35) which is again

possibly due to the counter ions of the buffer shielding the Phe residue at high ionic

strength causing the swelling to decrease.


135

3.4.5.2 Swelling behaviour of cleaved Fmoc-tripeptide-PEGA particles

After Fmoc-tripeptide-PEGA1900 particles (i.e. Fmoc-X-Ala-Ala-PEGA1900 and Fmoc-X-

Phe-Phe-PEGA1900) were treated with either elastase or thermolysin, it was envisaged that

the following cleaved products were produced: (+)Ala-PEGA1900 and (+)Phe-PEGA1900

(figure 36c) because the ERS was selectively cleaved by the protease between the Ala-Ala

or the Phe-Phe bonds of the Fmoc-tripeptide-PEGA1900 particles (chemically indicated

by figure 36a). Subsequently, the diameter of 100 particles for each cleaved product was

measured and the effect of swelling was again studied as a function of pH (7 – 9) and

ionic strength (0.001 – 0.2 M) as demonstrated by figure 37a for (+)Ala-PEGA1900 and

figure 36a for (+)Phe-PEGA1900.

Figure 36. Fmoc-tripeptide-PEGA1900 particles (a) cleaved by protease to generate cleaved products (b-c). Fmoc-tripeptide-PEGA1900 particles (a: Fmoc-R3-Ala-Ala-PEGA1900 and Fmoc-R3-Ala-Ala-PEGA1900); cleaved product 1 (b: Fmoc-R3-R2-COO–) and cleaved product 2 (c: (+)R1-PEGA1900); R-groups (R3: Arg(+)/ Glu(–); R1 and R2: Ala/ Phe). Diameters of Fmoc-tripeptide-PEGA1900 particles (a) and cleaved product 2 (c) were compared to determine the difference in swelling after enzyme treatment.

The swelling behaviour of both (+)Ala-PEGA1900 and (+)Phe-PEGA1900 particles were

similar. The effect of swelling significantly depended on ionic strength because the

swelling was found to decrease with increasing ionic strength (0.001M – 0.2M, p < 0.05)

this is because the electrostatic interactions decreased for both (+)Ala-PEGA1900 and (+)Phe-PEGA1900 particles because the counter ions of the buffer screened the

O

NNH

O

R3O

N

R2

O

N

R1

O

H3N N

R1

O

O

NNH

O

R3O

O

R2

O

Phe

CH3

Arg+

N N

N+

Glu

O

O

Gly H

(a)

(c)(b)

+

Protease

+

Ala R3:R1, R2:


136

electrostatic interactions of the α-NH3+ for both the Ala and Phe residues causing the

swelling to decrease. Nevertheless, within the range of 0.001 – 0.1M the effect of

swelling was not significantly different for the (+)Ala-PEGA1900 particles at pH 8 (p >

0.05) and (+)Phe-PEGA1900 particles at pH 9 (p > 0.05) which is possibly because the pH

of the surrounding buffer solution was closer to the pKa of the terminal α-NH3+ group

for both the Ala and Phe residues as discussed below.

(a) (b)

0.290

0.305

0.320

0.335

0.350

0.365

0.001 M 0.01 M 0.1 M 0.2 M

Diameter (mm)

-8

-4

0

4

8

12

16

20

0.001 M 0.01 M 0.1 M 0.2 M

Swelling (%)

(c) (d)

-8

-4

0

4

8

12

16

20

0.001 M 0.01 M 0.1 M 0.2 M

Swelling (%)

-8

-4

0

4

8

12

16

20

0.001 M 0.01 M 0.1 M 0.2 M

Swelling (%)

pH 7 pH 8 pH 9

Figure 37. Swelling behaviour of cleaved product (+)Ala-PEGA1900 and the percentage decrease/increase in swelling when compared to the un-cleaved Fmoc-X-Ala-Ala-PEGA1900 particles (n=100). Cleaved product: (+)Ala-PEGA1900 (a); cleaved Fmoc-X-Ala-Ala-PEGA1900 particles: RAA (b), EAA (c), and GAA (d). In graphs b-d the percentage increase in swelling is displayed by positive values; whereas the percentage decrease in swelling is displayed by negative values. The effect of swelling was studied as a function of pH (7, 8, 9) and ionic strength (0.001M – 0.2M) in potassium phosphate buffer. Statistical analysis was carried out using ANOVA (including post-hoc tests) followed by a paired t-test at the 95% confidence level. For detail descriptions of these comparisons see the text within this section.


137

In terms of pH, the effect of swelling for (+)Ala-PEGA1900 was significantly higher at pH

7 – 8 compared to pH 9 (p < 0.05) this is because the pH of the surrounding buffer at

pH 9 for the (+)Ala-PEGA1900 particles was closer to pKa 9.96 of the terminal α-NH3+ of

the Ala residue causing the swelling to significantly decrease (p < 0.05). The same trend

was observed for the (+)Phe-PEGA1900 particles because the swelling was significantly

higher at pH 7 compared to pH 8 – 9 (p < 0.05) because in this instance the pH of the

surrounding buffer at pH 8 – 9 was closer to pKa 9.13 of the terminal α-NH3+ of the Phe

residue causing the swelling to significantly decrease (p < 0.05).

(a) (b)

0.290

0.305

0.320

0.335

0.350

0.365

0.001 M 0.01 M 0.1 M 0.2 M

Diameter (mm)

-8

-4

0

4

8

12

16

20

0.001 M 0.01 M 0.1 M 0.2 M

Swelling (%)

(c) (d)

-8

-4

0

4

8

12

16

20

0.001 M 0.01 M 0.1 M 0.2 M

Swelling (%)

-8

-4

0

4

8

12

16

20

0.001 M 0.01 M 0.1 M 0.2 M

Swelling (%)

pH 7 pH 8 pH 9

Figure 38. Swelling behaviour of cleaved product (+)Phe-PEGA1900 and the percentage decrease/increase in swelling of the cleaved Fmoc-X-Phe-Phe-PEGA1900 particles (n=100). Cleaved product: (+)Phe-PEGA1900 (a); cleaved Fmoc-X-Phe-Phe-PEGA1900 particles: RFF (b), EFF (c), and GFF (d). In graphs b-d the percentage increase in swelling is displayed by positive values; whereas the percentage decrease in swelling is displayed by negative values. The effect of swelling was studied as a function of pH (7, 8, 9) and ionic strength (0.001M – 0.2M) in potassium phosphate buffer. Statistical analysis was carried out using ANOVA (including post-hoc tests) followed by a paired t-test at the 95% confidence level. For detail descriptions of these comparisons see the text within this section.


138

Next, the swelling behaviour of the cleaved products were each compared with the

swelling of the un-cleaved Fmoc-tripeptide-PEGA particles as given in the previous

section i.e. (+)Ala-PEGA1900 against Fmoc-X-Ala-Ala-PEGA and (+)Phe-PEGA1900 against

Fmoc-X-Phe-Phe-PEGA. The difference in swelling between the two configurations

(cleaved and un-cleaved particles) was calculated and the percentage increase/decrease in

swelling for the positive (RAA, RFF), negative (EAA, EFF) and neutral (GAA, GFF)

particles were plotted separately for both the Ala containing particles (figure 37b-d) and

the Phe containing particles (figure 38b-d). In both figures, the percentage increase in

swelling is displayed by positive values; whereas the percentage decrease in swelling is

displayed by negative values. From figures 37b-d and figures 38b-d it can be seen that

majority of the particles showed an increase in swelling between the diameters of the un-

cleaved versus cleaved particles for both Ala and Phe containing particles (figures 37 and

38, respectively). In both configurations, the increase in swelling was observed more for

the neutral particles (figure 39c: GAA and figure 40c: GFF) compared to the charged

particles (figures 37b-c: RAA, EAA; and figures 38b-c: RFF, EFF). The average increase

in swelling for the neutral particles was 12.19 % for GAA (figure 37d) and 13.61 % for

GFF (figure 38d), whereas for the positive particles it was 3.81 % for RAA (figure 37b)

and 9.89 % for RFF (figure 38b); and for the negative particles it was 8.66 % for EAA

(figure 37c) and 10.64 % for EFF (figure 38c).

On the contrary, a decrease in swelling was only observed for the charged particles at low

ionic strengths. For the Ala containing particles, the swelling of the positive particles

(RAA, figure 37b) decreased at pH 9 by 2.33 % (0.001 M) and 3.77 % (0.01 M); whereas

the swelling of the negative particles (EAA, figure 37c) decreased by 4.09 – 6.67 %

between pH 7 – 9 (at 0.001 M). For the Phe containing particles, the swelling of the

positive particles (RFF, figure 38b) decreased at 0.001M by 3.51 % at pH 8, and 4.31 %

at pH 9; whereas the negative particles (EFF, figure 38c) only decreased by 0.90 % at pH

8 (0.001 M). For the charged particles of both Ala and Phe containing particles (RAA,

EAA, RFF, EFF) the difference in swelling was found to gradually increase with

increasing ionic strength; whereas no measurable effect was observed between 0.001 –

0.2 M for the neutral particles of both Ala and Phe containing particles (GAA: figure

37d; and GFF: figure 38d). Finally, the difference in swelling at each ionic strength for

majority of the particles was found to decrease with increasing pH (7 – 9), which as

mention previously is caused by the pH of the surrounding buffer at pH 9 being closer to


139

the pKa of the terminal α-NH3+ of both the Ala/ Phe residue(s); whereas at pH 7 the pH

of the surrounding buffer is further away from the pKa of α-NH3+ causing the particles

to swell more.

3.5 CONCLUSION

Using SPPS, PEGA particles were successfully functionalised with short ECPs (Fmoc-

dipeptides and Fmoc-tripeptides) that selectively responded to selected proteases such as

thermolysin and elastase. Comparative studies demonstrated that in sample fluids

mimicking the environment of chronic wounds (i.e. pH 6.0 – 9.0; with a protease

concentration > 0.1 mg/ml) elastase selectively hydrolysed Fmoc-tripeptide ECPs

containing the Ala-Ala bond, whereas thermolysin had a high preference to hydrolyse

both the Fmoc-tripeptide and Fmoc-dipeptide ECPs (the latter ECP did not contain the

Ala-Ala bond). Additionally, the results illustrate that under the influence of pH and

charge, the enzyme-response mechanism of the hydrogel PEGA particles was specifically

tuned so that each protease in its cationic or anionic form could simultaneously and

selectively penetrate functionalised PEGA particles and hence selectively hydrolyse ECPs

of opposite charge to that of the protease. Thermolysin in its anionic form readily

cleaved a positively charged ECP compared to a negatively charged ECP; whereas

elastase in its cationic form had preference to cleave a negatively charged ECP and in its

anionic form the opposite effect was observed. The swelling of PEGA particles

increased with the incorporation of ionic charges (i.e. positive and negative) compared to

PEGA particles possessing a neutral charge. For the ionic particles, removal of the

charge residue (i.e. CMR: positive or negative) by proteases showed a reduction in

particle swelling at low ionic strength compared to neutral particles; whereas at high ionic

strength the swelling was found to increase for all particles (ionic and neutral). Overall,

this chapter exemplifies that on the basis of charge selective cleaving of ECPs coupled to

PEGA800 and PEGA1900 particles can be exploited to selectively remove excess proteases

such as elastase from sample fluids mimicking the environment of chronic wounds. In

doing so, PEGA particles containing ionic charges alongside the Ala-Ala bond may have

an application as new responsive dressings to treat chronic wounds by selectively

removing and entrapping elastase into PEGA particles as investigated further in the next

chapters.

CHAPTER 4: Selective Entrapment of Elastase into PEGA Particles

140

CCHHAAPPTTEERR 44

SSeelleeccttiivvee EEnnttrraappmmeenntt ooff EEllaassttaassee iinnttoo PPEEGGAA PPaarrttiicclleess


141

4.1 INTRODUCTION

This chapter focuses on understanding the accessibility and diffusion of elastase followed

by its entrapment into PEGA particles by exploiting the use of different fluorescence

techniques. The term fluorescence is very broad as it has many applications and various

techniques can be used to study it. Fluorescence techniques are used in chemistry,

physics and biology and are highly sensitive for the detection and quantification of

molecular interactions and chemical reactions over time. They are highly valuable for

studying cellular structure and function, and bio-molecular interactions of molecules in

biological systems and have gradually become important for the detection and

quantification of: nucleic acids and proteins in gel electrophoresis, microarrays, material

and polymer science and fluorescence spectroscopy.

Fluorescence occurs only in poly-aromatic hydrocarbons and/ or heterocyclic molecules

which are normally referred to as ‘fluorophores’ or fluorescent dyes/ markers as they

absorb light. These molecules are further classed as ‘chromophores’ as part of the

molecule is responsible for colour. A chromoprotein is a protein that absorbs light in the

visible or near-UV range but one that does not necessarily emit fluorescence. To-date

many fluorophores are available which have been synthesised and modified to specifically

interact with cellular/ chemical structures to make them detectable in many different

colours across the spectrum (UV/VIS/IR). The Jablonski energy diagram is a useful

diagram that summarises the kinetic spectra of light absorption and fluorescence (figure

39a). The phenomenon of fluorescence dates back to 1952 when Sir George Gabriel

Stokes reported the shift of fluorescence to a lower energy compared to the absorption

spectrum (Stokes 1852), which is famously known as the ‘Stokes shift’. The process of

fluorescence via the ‘Stokes shift’ of a given fluorophore is summarised in figure 39b-c.

For PEGA research, a choice of fluorophores have been utilised to examine the

accessibility and permeability of enzymes into PEGA particles. Previously, 1-

dimethylaminonapthalene-5-sulfochloride (dansyl chloride) has been used to understand

the diffusion of enzymes into PEGA particles over the course of time (Ulijn et al.

2003b). This study showed that after thermolysin cleaved the ECP, Fmoc-Phe-Phe

coupled to PEGA particles free amino group (NH2) became available and after that the


142

thermolysin reaction was stopped, the free amino groups on the cleaved PEGA particles

were labelled with dansyl chloride.

(a) (b)

(c) (d)

Figure 39. Jablonski diagram and Stokes shift of fluorescence. (a) Jablonski diagram: A fluorophore molecule in its ground state, S0 absorbs sufficient light from an external source at one wavelength causing the molecule to raise an electron to an excited state that is at an higher-energy state (S1) resulting in luminescence (Campell, 1988) via the process of excitation. A fluorophore can attain multiple excited states (S1, S2) depending on the wavelength or the energy of the external light source. The excited lifetime is the length of time the fluorophore exist in excited states; it lasts for a very short time between 10-15–10-9 seconds. At high-energy level the electrons of the fluorophore are unstable therefore the fluorophore adopts a semi-stable state (strong dotted lines). The excess of energy is then converted to the emission of fluorescence or the light returns the fluorophore back to the ground state, S0 (a-d). The emitted fluorescence energy contains less energy and is at a longer wavelength whereas the absorbed light energy is at a shorter wavelength, this is known as the ‘Stokes Shift’ (b-c) where some of the energy is lost through molecular vibrations during the transient excited lifetime as dissipated heat to the surrounding solvent molecules as they collide with the excited fluorophore (c). The absorption of fluorescence is a cyclic process (d) unless the fluorophore is irreversibly destroyed by continuous light excitation leading to photobleaching. Figure and text tailored from Haugland 2005, Invitrogen technical notes, Kealey and Haines 2002; and Campell 1998.

So

S1

S2

Ab

sorp

tion

(k

ex : 10-15 s)

Flu

orescen

ce (k

em: 10

-9 to -7 s)

Internal conversion (10-12 to -6 s)

750

Stokes shift

Wavelength of light (nm)

400 450 500 550 600 650 700

Emission Excitation

Freq

uen

cy

Heat

Energy levels

Ex

citatio

n

Em

ission

Stokes shift Absorption

Ground State

Emission

Excitation


143

Then, after washing the cleaved PEGA particles, the spatial distribution of the

fluorescence for the dansyl fluorophore was detected using two-photon microscopy

(TPM) to show where thermolysin had cleaved the ECP from the Fmoc-Phe-Phe-PEGA

particles. The complete diffusion of the thermolysin from the outer surface throughout

the central core of PEGA particles was achieved within 45 minutes (Ulijn et al. 2003b).

Since unmodified PEGA particles contain free NH2 groups (which as mentioned

previously are used during SPPS to couple amino acids) and these can be labelled with

dansyl chloride to examine whether an amino acid, hence ECP has completely been

coupled to PEGA particles. Though, this approach is more expensive compared to

Kaiser test given that dansyl coupling requires advanced microscopy for analysis unlike

the Kaiser test (for details of the Kaiser test: see chapter 3).

Fluorescein-isothiocyanate (FITC) labelled dextrans (referred as FITC-labelled dextrans

in this thesis) of known molecular weights have been used to understand the accessibility

and diffusion of enzymes into unmodified or functionalised PEGA particles that have

been treated with or without a protease. Thornton et al. (2005) examined the diffusion

of FITC-labelled dextrans with molecular weights in the range of 4 – 77 kDa into the

core of the PEGA800 particles in water. The study showed that FITC-labelled dextrans <

20 kDa were able to access unmodified PEGA800 particles, whereas FITC-labelled

dextrans of high molecular weight (40 kDa and 77 kDa) were unable to access

unmodified PEGA particles. However, the penetration of these FITC-labelled dextrans

was enhanced when the swelling of the PEGA particles was increased by the

introduction of positive charges in which the ECP examined were Arg(+)-Phe-Gly and

Arg(+)-Gly-Phe. Additionally, they demonstrated that after both trypsin and thermolysin

selectively cleaved the Arg(+)-Gly-Phe ECP, the accessibility of the 77 kDa FITC-labelled

dextran into PEGA particles was reduced since the Arg(+) residue from the ECP was

removed causing the diameter of the PEGA800 particle to reduce. In contrast, the study

showed that when the unspecific protease i.e. chymotrypsin did not cleave Arg(+)-Gly-

Phe ECP the molecular accessibility of these particles was not reduced. As a result the

FITC-labelled dextran was able to access and diffuse into PEGA800 particles, and its

accessibility was comparable to that observed by the control PEGA800 particles which

were not cleaved with proteases (Thornton et al. 2005).


144

Both dansyl chloride and FITC-labelled dextrans require the enzyme reaction to be

stopped and then the accessibility and diffusion of the target protease into PEGA

particles can be examined. However, real-time monitoring would be more advantageous

to directly visualise the accessibility, penetration and entrapment of a target protease into

functionalised PEGA particles. In cell biology, FITC is the simplest and cheapest way of

labelling a protein of interest. In this way, a FITC-labelled protein can then be used as a

tracer to examine the permeability or the location of the protein inside cells to

understand the cellular functions of the target protein (Berg et al. 2007). This chapter

takes advantage of this by labelling elastase with FITC to directly visualise the

accessibility and diffusion followed by entrapment of elastase into functionalised PEGA

particles in real-time via TPM.

TPM involves two-photon excitation which means that two photons (low energy) are

simultaneously absorbed to excite a fluorophore’s electrons in a single quantum event as

summarised in figure 40. The emission of fluorescence is of higher energy compared to

the excitatory photons (Rocheleau and Piston 2003; Ulijn et al. 2003b; Haugland 2005,

Invitrogen technical notes). However, the two photons are approximately twice the

wavelength of a one-photon excitation because the fluorescence emission depends on

the square of the excitation intensity (Rocheleau and Piston 2003; figure 40).

Figure 40. Jablonski diagram comparing the absorption of two-photon against one-photon. Left: a single-photon (one purple arrow) absorbed, generates fluorescence of an excited state. Right: simultaneous absorption of two-photons (two purple arrows) can produce identical fluorescence of an identical exited state. Abbreviations: So (ground state), S1 (excitated stated), purple arrow (absorbed photon), green arrow (fluorescence of an excited state). Figure adapted from Rocheleau and Piston 2003. Coinciding with TPM, confocal microscopy enables cross-sectional images to be

obtained, however for the hydrogel PEGA particles TPM is recommended (Ulijn et al.

So

S1

One-photon Two-photon


145

2003b). In view of the fact that, the fluorescence can be quenched at the centre of

PEGA particles with confocal microscopy, whereas with TPM the spatial distribution of

the fluorophore, hence fluorescence is equally distributed across the entire surface and

interior of PEGA particle (Kress 2002; Ulijn et al. 2003b). This is because in TPM the

higher wavelength (lower photon energy) excitation produces fluorescence at the focal

point and does not produce background fluorescence because the excitation is confined

to the vicinity of the focus with no excitation outside the focal point. Therefore, the

fluorescence is purely generated from the sample that is in focus (Ulijn et al. 2003b;

Bosma et al. 2003). In this way, more of the excitation light can profoundly penetrate

into the sample (e.g. PEGA particles) to the plane of focus producing high quality 3D

cross-sectional images.

For the detection of protease activity, synthetic fluorescence substrates are more

commonly used as the assays are practical and highly sensitive. A fluorescence substrate

contains a fluorophore at one end with an ECP in the middle and a quencher at the other

end. In this way, enzyme activity is directly determined by an increase in fluorescence after

the protease cleaves a peptide bond of the ECP releasing the fluorophore which is then

excited by an external source to emit fluorescence (as described above). The commercial

fluorescence substrate, methylsucciyl-alanyl-alanyl-proline-valine-alanyl-coumarin (MeO-

Suc-Ala-Ala-Pro-Val-Ala-AMC) is graded as the second most fluorescence substrate used

to assess elastase activity of PPE compared to MeO-Suc-Ala-Ala-Pro-Val-Ala-pNA

(Castillo et al. 1979). On the contrary, in this thesis, MeO-Suc-Ala-Ala-Pro-Val-Ala-AMC

was selected since it is highly sensitive because its fluorophore i.e. 7-amino-4-

methylcoumarin (AMC) is effortlessly detected compared to the peptidyl-4-nitroanilide

(pNA) fluorophore of the MeO-Suc-Ala-Ala-Pro-Val-Ala-pNA fluorescence substrate

(Castillo et al. 1979). For the remaining chapters, MeO-Suc-Ala-Ala-Pro-Val-Ala-AMC

was used to demonstrate the reduction of elastase activity in sample fluids after they had

been treated with functionalised PEGA1900 particles.

For a chronic wound dressing, the capacity of the dressing to absorb wound exudates

containing proteases into the dressing is important depending on both the pH and ionic

strength of the wound environment. Consequently, in this chapter the pH of the buffer

solution was selected within the range of 7 – 9 in order to examine the accessibility,

diffusion and entrapment of elastase both in its cationic and anionic forms. In chapter 3,


146

the selective hydrolysis of elastase was studied at low ionic strength, but for medical

relevance, the accessibility and diffusion of elastase is required to be studied at high ionic

strength in which 0.1 M was selected. This ionic strength was chosen since previously the

swelling of GAA particles was found to decrease un-expectantly at 0.2 M compared to

0.001 M – 0.1 M (see section 3.4.5). Consequently, 0.1 M was chosen as it is closer to

physiological ionic strength of 0.15 M.

4.2 OBJECTIVES

Having established in chapter 3 that Fmoc-X-Ala-Ala-PEGA1900 particles were more

appropriate in targeting elastase, accordingly the focus of this chapter was to understand

the selective hydrolysis of all ECPs (RAA, EAA, GAA) by sample fluids mimicking the

elastase environment (0.01 – 0.1 mg/ml) observed in chronic wounds under the

influence of pH 7 – 9 with an ionic strength of 0.1 M. In conjunction, a variety of

fluorescence techniques were used to understand and demonstrate the accessibility and

diffusion followed by entrapment of elastase (cationic and anionic forms) into

functionalised PEGA particles (RAA, EAA, GAA) depending on the pH and at high

ionic strength (as indicated).


4.3.1 Materials

PEGA(800 and 1900) – NH2 particles in methanol were supplied by Polymer Laboratories Ltd

(UK). Fmoc protected amino acids (Fmoc-Ala-OH.H2O, Fmoc-Arg(pbf)-OH, Fmoc-

Glu(OtBu)-OH, Fmoc-Gly-OH, Fmoc-Phe-OH) and the elastase substrate, MeO-Suc-

Ala-Ala-Pro-Val-AMC were all supplied by Bachem (UK). Elastase (pancreatic from

porcine pancreas, PPE, 3.9 mg/protein) was purchased from Worthington Biochemical

Corporation (USA). Aldrich (UK) supplied acetonitrile (ACN), dimetylformanide

(DMF), hydroxybenzotriazole (HOBt), di-isopropylcarbodimide (DIC). Piperidine and

potassium mono- and di-phosphate solution (1.0 M, 1L), FITC and FITC-labelled

dextrans (MW 20 kDa) were all purchased from Sigma (UK). HiPerSolv HPLC grade

ACN was from VMR international (UK). Dansyl chloride was purchased from Fluka

Analytical (UK). Precasted 4-20% Tris-glycine gels, molecular weight markers, loading


147

buffer and Tris-glycine SDS (sodium dodecyl sulphate) buffer, Coomassie brilliant Blue

R-250, Mini-Protean® 3 Cell and PowerPacTM Univeral for electrophoresis were all

supplied by Biorad (Hertfordshire, UK). All solvents used were of the highest purity

without further purification. Isolute double fritted filtration columns were purchased

from Kinesis. Stuart Scientific supplied the blood rotator and roller mixer. HPLC 680

series was purchased from Dionex and the C18 nucleosil column was purchased from

Fisher.

4.3.2 Methods

4.3.2.1 Fmoc SPPS and Kaiser test

Functionalised PEGA particles with the configuration Fmoc-X-Ala-Ala-PEGA (RAA,

EAA, GAA) and Fmoc-X-Phe-Phe-PEGA (RFF, EFF and GFF) were prepared using

the protocol as previously described in chapter 3, section 3.2.2.2. The successful

coupling of amino acids and de-protecting of the Fmoc groups during SPPS was

monitored using the Kaiser test (see section 3.3.2.3).

4.3.2.2 Potassium phosphate buffer

Stocks of 0.001 M and 0.1 M potassium phosphate buffer within the pH range of 7.0 –

9.0 were prepared by serial dilution of the 1 M potassium mono- and di-phosphate

solutions in deionised water. Each buffer solution was adjusted to its desired pH with

HCl and NaOH using a pH meter.

4.3.2.3 Enzyme hydrolysis using HPLC

After SPPS, functionalised PEGA particles were thoroughly washed with MeOH, dH2O

and potassium phosphate buffer at appropriate pH (7, 8, 9) and ionic strength (0.001M

and 0.1M). Swollen functionalised PEGA1900 particles (50 mg) were then cleaved with

elastase (PPE, 2 ml at 0.01 or 0.1 mg/ml in 0.001M or 0.1M, pH 7 – 9) for various time

points at 34 – 37oC. The supernatants of the cleaved products were collected and the

reaction was stopped by washing the PEGA particles with ACN:H2O (80:20, 9 ml)

containing 0.1% TFA. This washing ensured that all the cleaved products were collected

from of the interior of the functionalised PEGA particles. The procedure was carried


148

out in duplicates. The supernatants of the cleaved products were analysed by HPLC and

the percentage cleaved products were calculated as previously described in chapter 3

(section 3.3.2.6).

PEGA particles were then stained or incubated with various fluorophores (as described

below). In order to maintain the swelling of PEGA particles depending on pH and ionic

strength, prior to TPM the PEGA particles were washed with the appropriate buffer at

the appropriate pH and ionic strength (as indicated).

4.3.2.4 Dansyl chloride staining

After elastase hydrolysis, PEGA particles (un-cleaved/ cleaved) were thoroughly washed

with 0.1 M potassium phosphate buffer (at appropriate pH 7 – 9), followed by DMF:

MeOH (50:50) and finally with DMF. The prewashed PEGA particles were stained with

the fluorophore, dansyl chloride which reacts with free amine/ amino groups (figure 41)

that are present on both unmodified and cleaved PEGA particles (including amino

acids).

Figure 41. Chemical reaction of dansyl chloride with cleaved PEGA particles. The free amine group of cleaved PEGA particles (or of unmodified PEGA particles) react with dansyl chloride to generate dansylated-PEGA particles (dansyl-PEGA particles) that fluoresce blue when viewed by TPM.

Dansyl chloride (18 mg) was dissolved in 2 ml DMF and 20 µl DIPEA and this solution

was added to the foil wrapped filtration columns containing PEGA particles (un-

cleaved/ cleaved: 50 mg) and left to stain between 3 – 12 hours on a blood rotator at

room temperature. After staining, PEGA particles were thoroughly washed with DMF

to remove all excess dansyl chloride that did not react with the free amine group of

PEGA particles. The PEGA particles were further washed with MeOH: DMF (50:50),

N

S

Cl

O O

CH3

CH3

N

S

NH

O O

CH3

CH3

O

R1

N

O

NNH2

R1

+

Dansyl choride

PEGA particles

(with free amine group)

Dansylated-PEGA particles

DIPEA + DMF

(3 hours - overnight)


149

MeOH, dH2O. Finally washed and swollen with potassium phosphate buffer (0.1 M and

pH 7 – 9) for 15 minutes in order to control the swelling of PEGA particles at 0.1 M and

at the desired pH 7 – 9. The fluorescence of the dansylated-PEGA particles was

examined using TPM at λexc 340 nm (excitation) and λem 490 nm (emission). Cross-

sectional equatorial images of PEGA particles were taken, which were quantified

according to section 4.3.2.8 (as described below).

4.3.2.5 SDS-Page

The molecular weight of the commercially bought porcine pancreatic elastase (3.9 mg per

protein at 1 mg/ml) was determined by SDS-polyacrylamide gel electrophoresis (SDS-

Page). Elastase was dissolved in dH2O (1 mg/ml) and then diluted (1:1, v/v) with Biorad

loading buffer (consisted of 0.125 M Tris-Cl, 4% SDS, 20% glycerol, 10% dithiothreitol,

0.2% bromphenol blue) and heated for 10 minutes at 100oC in a water bath. Molecular

weight marker (5 µl, lane 1) and elastase (10 µl, lane 2) were loaded into the wells of the

gel. Elastase was electrophoretically separated on a pre-casted 4 – 20% SDS-Page gel

using 1X Tris-glycine SDS buffer at step voltage changes: at an initial constant voltage of

100 mV for 15 minutes to align the proteins and then at a final constant voltage of 200

mV for 45 minutes to separate into individual bands. Elastase bands were visualised by

staining the gel with Coomassie brilliant Blue R-250 and the gel was destained with 40%

methanol and 10% acetic acid for up to an hour at room temperature. The gel was then

transferred to a plastic cassette and subsequently wrapped with cling film and then

imaged using a flatbed scanner.

4.3.2.6 Accessibility of FITC-dextran

After SPPS and enzyme hydrolysis, un-cleaved/ cleaved functionalised PEGA particles

(50 mg) were thoroughly washed with potassium phosphate buffer (at pH 8 and at

0.001M or 0.1 M ionic strength). FITC-labelled dextran with a molecular weight of 20

kDa was dissolved in potassium phosphate buffer (at pH 8 with an ionic strength of

0.001M or 0.1 M) at a concentration of 1 mg/ml. To protect the FITC from light, the

vials were foil wrapped. Pre-washed functionalised PEGA particles (un-cleaved/

cleaved, 5 mg) were added to a glass slide and 1 drop of the FITC-labelled dextran was

added to the particles on the glass slide. The permeability of the 20 kDa FITC-labelled


150

dextran into the core of PEGA particles was monitored by taking cross-sectional

equatorial images of PEGA particles using TPM at λexc 494 nm and λem 520 nm over the

course of 10 - 30 minutes. The fluorescence of the cross-sectional equatorial images was

quantified using the analysis software as described in section 4.3.2.8 below.

4.3.2.7 Accessibility of FITC-elastase

O

O

OOH OH

S=C=N

R-NH2

O

O

OOH OH

R-N-C=N

SH

H

+

Elastase

FITC FITC-elastase

Figure 42. Chemical reaction of staining elastase with the base fluorescein molecule, FITC. The free amine groups within the elastase structure were coupled and cross-linked with FITC to generate a fluorescence-labelled elastase, referred as FITC-elastase.

The chemical reaction of cross-linking FITC to elastase is given in figure 42. A 24-fold

molar ratio is optimal for the conjugation of FITC (MW: 389 Da) with proteins. In a foil

wrapped vial, FITC dye (1 mg) was dissolved in DMF (100 µl) and the FITC dye was

completely dissolved by drawing the solution up-and-down using a micropipette.

Elastase (1 mg/ml) was dissolved in PBS buffer (pH 7.4, 0.15 M) in another foil wrapped

vial.

The appropriate amount of FITC (36.05 µl) was transferred to the elastase containing

vial. FITC dye and elastase were thoroughly mixed by drawing them up-and-down using

a micropipette and then incubated at room temperature for 1 hour on a blood rotator.

After labelling, FITC-elastase was then dialysed to remove any excess FITC dye that did

not crosslink to elastase. A dialysis tube (pore size 12 kDa) was cut to size and a clip was

placed at the bottom of the tube. The FITC-elastase solution was added into the dialysis

tube and the top of the dialysis tube with closed by another clip. The dialysis tube was

then dialysed in a beaker containing PBS (pH 7.4, 0.15 M) and to protect the FITC from

the surrounding light the beaker was covered with foil. The beaker was placed on a

stirrer-plate and continuously stirred in the range of 1 – 3 hours (maximum: overnight) at

room temperature. After that FITC-elastase was transferred to a clean foil wrapped vial

and stored at 4oC.


151

After SPPS, functionalised PEGA particles were thoroughly washed with dH2O and then

with PBS buffer (0.15M, pH 7.4). Functionalised PEGA particles (5 – 10 mg) were

added to glass slide, then two drops of FITC-elastase was added and after that the direct

penetration of fluorescently labelled elastase into PEGA particles was viewed by taking

cross-sectional equatorial images of functionalised PEGA particles using TPM at λexc 494

nm and λem 520 nm over the course of 15 minutes. The fluorescence of the cross-

sectional equatorial images was quantified using Leica or Image J analysis softwares as

described in section 4.3.2.8.

4.3.2.8 Quantification of TPM micrographs

The cross-sectional equatorial images of PEGA particles obtained from TPM were

quantified using two different analysis softwares. The average fluorescence was

quantified using the Leica analysis software, whereas the pixel intensity across the cross-

sectional images was quantified using Image J analysis software.

4.3.2.9 Fluorescence substrate assay

The fluorescence elastase substrate, MeOSuc-Ala-Ala-Pro-Val-AMC consisted of the

AMC fluorophore which was used to study elastase activity. The mode of substrate

hydrolysis by elastase is summarised in figure 43. The fluorescence substrate was

dissolved in 2 ml methanol (10 mM) and was further diluted to 20 ml with 0.1 M

potassium phosphate buffer within the range of pH 7 – 9 with a final concentration of

0.5 mM (1X stock). For enzyme kinetic studies, various concentrations of the

fluorescence substrate were prepared by initially dissolving the fluorescence substrate in

MeOH and then diluting it with 0.1 M potassium phosphate buffer to give a final

concentration that was: 10-fold, 5-fold, 2- fold, 1-fold in comparison to 1X stock. The

fluorescence substrate concentration of 2X (1.0 mM) was made using the same principles

as the 1X stock (Note: due to low solubility of the fluorescence substrate, concentrations

> 5X did not obey Michealis-Menten kinetics).

Elastase (PPE, 0.01 mg/ml) was dissolved in 0.1 M potassium phosphate buffer at pH 7

– 9. The fluorescence substrate assay was performed by incubating 50 µl of the


152

fluorescence substrate with 50 µl of the sample fluid. The sample fluid was either: (1)

Fluids containing elastase (PPE, 0.01 mg/ml); and (2) supernatants collected after

elastase was treated with functionalised PEGA particles at various time points. The

fluorescence for the hydrolysis of the substrate was measured at λexc 383 nm (excitation)

and λem 405 nm (emission) continuously for 14 hours. The rate of elastase activity was

expressed as RFU hr-1. After sample fluids were treated with functionalised PEGA

particles the rate of elastase activity was corrected according to the mass of PEGA

particles used i.e. rate / mass: RFU hr-1/mg. All assays were performed at 34 – 37oC in

a black 96-well plate with duplicate measurements.

NH

CH

NH

NCH3

ONH

CH

NH

CH

O

O CH3

O CH3

O

OCH

3CH

3

O

O

CH3

O

NH2

O

CH3

O

CH3

ONH

CH

O

O CH3

O

OH NH2

CH

OH

CH3

CH3

O

NNH

2

CH

CH3

O

O

OH

Elastase

MeOSuc-Ala-Ala-Pro-Val-AMC

AMC

+

MeOSuc-Ala-OH Ala-Pro

+ +

Valine

Figure 43. The mechanism for the cleaving of the fluorescence-quenched substrate: MeOSuc-Ala-Ala-Pro-Val-AMC by elastase. In this FRET substrate, the AMC (blue) fluorophore is conjugated to the peptide Ala-Ala-Pro-Val via an amide bond between the amine of coumarin and the carboxyl group of the C-terminal valine residue. Hydrolysis of this amide bond, including both the Ala-Ala and Pro-Val bonds in presence of elastase releases a free AMC causing the fluorescence to increase which is then monitored over time. Hydrolysis of peptide bonds is indicated by red lines. Figure tailored from Novabiochem (letters 02/05) and Patrick 2010.

4.3.2.10 Statistics

Data are expressed as the mean value + SEM and statistical analysis was carried out using

SPSS 16.0. Statistical significance was determined using ANOVA test (one-way and two-

way full factorial) and for pair-wise comparisons, post-hoc (Tukey and Duncan) tests

alongside the Student-test were conducted. A significant difference was observed at the

95% confidence level in which P values below 0.05 (i.e. p < 0.05) were considered

hν


153

statistically significant and values above 0.05 (i.e. p > 0.05) were not statistically

significant.


4.4.1 Selective hydrolysis of ECPs by elastase at high ionic strength as a

function of pH

Here, functionalised PEGA particles with the ECPs: RAA, EAA, GAA (test) and RFF,

EFF, GFF (control) were treated with elastase (PPE, 0.1 mg/ml) at pH 7 – 9 and 0.1 M.

Elastase was found to selectively cleave the Fmoc-X-Ala-Ala-PEGA particles (1.50 –

6.23%) significantly more compared to Fmoc-X-Phe-Phe-PEGA particles (0.36 – 0.84%)

(p < 0.05) as summarised in figure 44. On the basis of charge, the selectively of elastase

to cleave RFF, EFF and GFF was not significantly different (p > 0.05). However, as

expected for the Fmoc-X-Ala-Ala-PEGA1900 particles in its cationic form elastase

selectively cleaved EAA more (pH 7 – 8: 6.23 %) compared to both RAA (pH 7: 3.33 %

and pH 8: 4.70 %) and GAA (pH 7: 1.50 % and pH 8: 3.70 %). The opposite effect was

observed at pH 9 wherein elastase in its anionic form selectively cleaved RAA (5.39 %)

more compared to both EAA (4.70 %) and GAA (4.64 %).

It can be seen that the yields of the cleaved products for the Fmoc-X-Ala-Ala-PEGA

particles (figure 44: 1.50 – 6.23 %) are much lower than those obtained in chapter 3

(figure 30). This difference was expected given that in chapter 3 the selective hydrolysis

of RAA, EAA and GAA by elastase was studied at low ionic strength (0.001 M) whereas

in this chapter it is studied at high ionic strength (0.1 M). Previously, the swelling of

RAA, EAA and GAA was significantly more at 0.001 M than 0.1 M (figure 34 in chapter

3). In this way, it is expected that the accessibility of elastase into functionalised PEGA

to reduce at high ionic strength and as a consequence the amount of ECPs cleaved by

elastase would also reduced at 0.1 M (figure 44) in contrast to 0.001 M (figure 30 in

chapter 3).


154

0

2

4

6

8

RAA EAA GAA RFF EFF GFF

Cle

ave

d p

rod

uct

(%)

pH 7.0

pH 8.0

pH 9.0

Figure 44. Selective hydrolysis of Fmoc-X-Ala-Ala-PEGA1900 particles and Fmoc-X-Phe-Phe-PEGA1900 particles by elastase at high ionic strength under the influence of pH. These particles were treated with elastase at a concentration of 0.1 mg/ml within the pH range of 7.0 – 9.0 in 0.1M potassium phosphate buffer for 3 hours on a blood rotator at 34oC. The cleaved products were collected and analysed by rpHPLC. Elastase selectively cleaved the RAA, EAA, and GAA PEGA1900 particles significantly more than RFF, EFF and GFF PEGA1900 particles (p < 0.05). Cationic elastase (pH 7 – 8) had preference of cleaving EAA (negative) significantly more than RAA (positive) and GAA (neutral); and the opposite effect was observed wherein anionic elastase (pH 9) had preference of cleaving RAA more compared to EAA. Statistical analysis was conducted using ANOVA and pair-wise comparisons were conducted by post hoc (Tukey and Duncan) test: a significant difference observed at the 95% confidence level (p < 0.05). Data represents the mean + SE of two measurements (n = 2). .

4.4.2 Fluorescence studies

4.4.2.1 Diffusion of elastase via dansyl chloride

Next, the diffusion of elastase into functionalised PEGA1900 particles was monitored

using dansyl chloride staining. After the ECPs of functionalised PEGA1900 particles were

cleaved by elastase (0.1 mg/ml), the free amine groups of the cleaved PEGA1900 particles

were stained with dansyl chloride (for chemical reaction see figure 41). After that, cross-

sectional equatorial images were obtained using TPM in which the fluorescence of the

dansylated-PEGA particles was used to visualise and measure the accessibility/ diffusion

of elastase into functionalised PEGA particles at 0.1 M as a function of pH. Un-cleaved

functionalised PEGA1900 particles were used as a control (0 minutes, figure 45 and 46) to

demonstrate that the ECP had fully coupled unmodified PEGA1900 particles during SPPS

and to compare the fluorescence observed by the cleaved functionalised PEGA1900

particles between 5 – 60 minutes.


155

0

50

100

150

200

250

RFF EFF GFF

Ave

rag

e F

luore

scen

ce (

RF

U)

0 min 5 min 15 min 30 min 60 min

Figure 45. TPM micrographs (top) and the average fluorescence graph (bottom) depicting the fluorescence labelling after un-cleaved and cleaved Fmoc-X-Phe-Phe-PEGA1900 particles (RFF, EFF and GFF) were stained with dansyl chloride at pH 8 (0.1 M). Note: the fluorescence of the un-cleaved particles at 0 min represented the baseline levels; as a result the fluorescence of the cleaved particles (5 – 60 min) were normalised by subtracting the fluorescence of baseline values for each Fmoc-X-Phe-Phe-PEGA1900 particles. ECPs were fully coupled to unmodified PEGA1900 as the un-cleaved Fmoc-X-Phe-Phe-PEGA1900 particles (0 min) were found to fluoresce to a lesser extent compared to the cleaved Fmoc-X-Phe-Phe-PEGA1900 particles (5 – 60 min). Unexpectedly the cleaved RFF, EFF and GFF PEGA1900 particles were found to fluoresce to a high degree compared to un-cleaved particles. Scale bars range from 50 - 100 µm (as indicated). Data represents the mean + SE of the whole well fluorescence (n = > 15 particles).

The manufacturer (Worthington Biochemical Corporation) reported elastase to have an

optimum pH at 8.0, therefore as a starting point the diffusion of elastase by dansyl

chloride staining was examined at pH 8.0 first. It was observed that the fluorescence of

the un-cleaved functionalised PEGA1900 particles (0 minutes) was low compared to the

cleaved products (5 – 60 minutes). This indicated that the ECPs: RFF, EFF, GFF (figure


RFF

EFF

GFF


156

45) and RAA, EAA, and GAA (figure 46) were fully coupled to unmodified PEGA1900

particles.

0

50

100

150

200

250

RAA EAA GAA

Ave

rag

e F

luore

scen

ce (

RF

U)


Figure 46. TPM micrographs (top) and the average fluorescence (bottom) illustrating the fluorescence observed after un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles (RAA, EAA and GAA) were stained with dansyl chloride at pH 8 (0.1 M). Note: the fluorescence of the un-cleaved particles at 0 min represented the baseline levels; as a result the fluorescence of the cleaved particles (5 – 60 min) were normalised according to the baseline values for each Fmoc-X-Ala-Ala-PEGA1900 particles. ECPs were fully coupled to unmodified PEGA1900 as the un-cleaved Fmoc-X-Ala-Ala-PEGA1900 particles (0 min) were found to fluoresce to a lesser extent compared to cleaved Fmoc-X-Ala-Ala-PEGA1900 particles (5 – 60 min). The fluorescence of all cleaved particles was found to increase over time, and was completely saturated at 60 minutes. Scale bars represent 75 – 250 µm (as indicated). Data represents the mean + SE of the whole well fluorescence (n = > 15 particles).

Since elastase was previously found to cleave RFF, EFF and GFF to a lesser extent than

RAA, EAA and GAA (see figure 44), it was predicted that the cleaved Fmoc-X-Phe-Phe-

PEGA1900 particles should fluoresce at a lower degree than Fmoc-X-Ala-Ala-PEGA1900

particles. However, both the un-cleaved and cleaved Fmoc-X-Phe-Phe-PEGA1900

RAA

EAA

GAA



157

particles (figure 45) were found to fluoresce to a high degree in contrast to Fmoc-X-Ala-

Ala-PEGA1900 particles (figure 46). The un-cleaved Fmoc-X-Phe-Phe-PEGA1900 particles

had an average fluorescence of 35.46 RFU at 0 minutes; whereas the cleaved particles

were found to fluoresce in the range of: 111.36 – 210.93 RFU (RFF), 12.09 – 218.67

RFU (EFF) and 167.40 – 220.41 RFU (GFF) between 5 – 60 minutes (figure 45). In

contrast the average fluorescence of the un-cleaved Fmoc-X-Ala-Ala-PEGA1900 particles

was 28.59 RFU (at 0 minutes) and the cleaved particles were found to fluoresce in the

range of: 66.89 – 223.88 RFU (RAA), 1.03 – 227.03 RFU (EAA) and 8.72 – 224.58 RFU

between 5 – 60 minutes (figure 46). The possible reason for the increase in fluorescence

observed by the cleaved Fmoc-X-Phe-Phe-PEGA1900 particles is because the aromatic

side chains of the Phe residues of the ECPs and the aromatic groups of dansyl chloride

are in close proximity of one another (Creighton 1993). This means that when dansyl

chloride was excited by TPM, it is possible that energy was transferred between the

aromatics groups causing the fluorescence of the cleaved particles to increase (Creighton

1993). Subsequently, in order to avoid obtaining false observations, for the remaining

experiments, the accessibility/ diffusive behaviour of elastase into Fmoc-X-Phe-Phe-

PEGA1900 particles by various fluorophores was abandoned.

From figure 46 it can be seen over the course of time, elastase increasingly penetrated

Fmoc-X-Ala-Ala-PEGA1900 particles since the fluorescence of the cleaved Fmoc-X-Ala-

Ala-PEGA1900 particles was found to gradually increase from 5 – 60 minutes. The

expected observation was only observed at 15 minutes in which elastase in its cationic

form was found to diffuse into the EAA particles more compared to both RAA and

GAA particles. Un-expectantly, at both 5 and 30 minutes cationic elastase was found to

diffuse and cleaved the RAA ECP (5 min: 89.20 RFU and 30 min: 216.31 RFU)

significantly more compared to EAA (5 min: 1.03 RFU and 30 min: 90.70 RFU) (p <

0.05). There was no significant difference in the diffusion of elastase at 60 minutes as

the average fluorescence for all cleaved Fmoc-X-Ala-Ala-PEGA1900 particles was

significantly similar i.e. RAA: 223.88 RFU; EAA: 227.03 RFU and GAA: 224.58 RFU (p

> 0.05). For that reason, > 60 minutes was considered as the sufficient time to cleave all

ECPs from functionalised PEGA1900 particles.

Next, several attempts were carried out to visual that elastase (0.1 mg/ml at pH 8 and

0.1M) firstly cleaved the ECPs of Fmoc-X-Ala-Ala-PEGA1900 particles from the outer


158

surface and then following its diffusion into Fmoc-X-Ala-Ala-PEGA1900 particles elastase

cleaved the ECPs located within the inner-central core of Fmoc-X-Ala-Ala-PEGA1900

particles. It was difficult to capture this by TPM at 0.1 M, as the entire surface of the

Fmoc-X-Ala-Ala-PEGA1900 particles appeared to be cleaved within 5 minutes seeing as

no ring-effect was observed at this time point (figure 46). A brief description illustrating

the meaning of a ‘ring-effect’ has been given in figure 67 (see appendix III). This

suggested that as soon as elastase was treated with Fmoc-X-Ala-Ala-PEGA1900 particles

the elastase reaction was completed within 2 – 5 minutes for the entire surface (i.e. outer

and inner) of the cleaved particles. For that reason, it was envisaged that possibly

slowing down the enzyme reaction may facilitate the formation of rings; consequently the

concentration of elastase was diluted to 0.01 mg/ml. In addition, since the entire surface

appeared to be cleaved at 0.1 M, the ionic strength was reduced as well because

previously in chapter 3 the swelling of Fmoc-X-Ala-Ala-PEGA1900 particles was more

profound at low ionic strength (0.001 M) compared to high ionic strength (0.1 M).

Subsequently, a time course study was then carried out in which Fmoc-X-Ala-Ala-

PEGA1900 particles were cleaved with elastase (0.01 mg/ml at pH 8 and 0.001 M) and the

elastase reaction was stopped at various time points as indicated in figure 47.

From figure 47, the un-cleaved Fmoc-X-Ala-Ala-PEGA1900 particles (0 min) were found

to fluorescence to a lesser extent compared to cleaved Fmoc-X-Ala-Ala-PEGA1900

particles (5 – 180 min). This indicated that all ECPs were successfully coupled to

unmodified PEGA1900 particles during SPPS. As expected, the cationic form of elastase

had preference to diffuse into EAA particles more compared to both RAA and GAA

particles. Interestingly, from the TPM micrographs a ring-effect of fluorescence was

observed for the EAA particles (figure 47a). These micrographs clearly demonstrated

that elastase first diffused into EAA particles and cleaved the ECPs from the outer

surface < 5 minutes. Then, elastase diffused into the inner-core of EAA particles

wherein the ECP was gradually cleaved from the inner surface and then completely from

the centre of EAA particles with 30 minutes compared to RAA and GAA (figure 47a).

However, from the pixel intensity graph (figure 45b) it was measured that the ECP was

completely cleaved from the centre of EAA particles within 1 hour rather than 30

minutes as fluorescence for dansyl chloride staining was found to be homogenous

between 60 – 180 minutes since the average fluorescence was not significantly different


159

(72.72 – 75.79 RFU: p > 0.05; figure 45b). To further confirm this, at each time point

the cleaved products were collected and analysed by rpHPLC. From figure 45c it can be

seen that the cationic form of elastase selectively hydrolysed the negatively charged ECP

(EAA: 20.79 – 60.92 %) more compared to both the positive ECP (RAA: 0.23 – 0.78 %)

and neutral ECP (GAA: 1.65 – 2.68 %).

(a)

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Figure 47. The selective diffusion of elastase into Fmoc-X-Ala-Ala-PEGA1900 particles (RAA, EAA and GAA) followed by the selective hydrolysis of the corresponding ECPs in potassium phosphate buffer at pH 8 and 0.001M monitored over the course of 180 minutes. (a) TPM micrographs demonstrating the fluorescence observed after un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles were stained with dansyl chloride (a). The pixel intensity graph was plotted by measuring the average fluorescence across the equatorial cross-section of the TPM micrographs illustrating the gradual diffusion of elastase into RAA, EAA and GAA particles (b) in which the data represents the mean + SE of the whole well fluorescence (n = 10 particles). At each time point, the cleaved products were collected and analysed by rpHPLC (c) in which the data represents the mean + SE of two measurements (n = 2). In its cationic form, elastase selectively diffused into EAA particles and cleaved its ECP significantly more compared to both RAA and GAA particles (p < 0.05). Scale bar of TPM micrographs represent 75 – 100 µm (as indicated).

0 min 10 min 180 min 60 min 30 min 5 min

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Next, the diffusion of elastase was examined at pH 7 and pH 9 after Fmoc-X-Ala-Ala-

PEGA particles were cleaved for 3 hours with elastase (0.1 mg/ml) at 0.1 M rather than

0.001 M. This is because previously at 0.1 M we encountered that elastase in its cationic

form (pH 8) penetrated RAA particles more compared to EAA and GAA (figure 46).

Consequently, it would be interesting to examine whether the same effect is observed at

pH 7 seeing as elastase is positively charged at both pH 7 and pH 8.

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Figure 48. TPM micrographs demonstrating the diffusion of elastase after Fmoc-X-Ala-Ala-PEGA1900 particles were cleaved with elastase for 3 hours at pH 7 and pH 9 (0.1 M). In its cationic form, elastase unexpectedly diffused and cleaved the RAA particles significantly compared to EAA and GAA particles. However, at pH 9, the expected effect was observed at pH 9, wherein elastase in its anionic forms selectively diffused into RAA more compared to EAA and GAA. Scale bar represents 250 µm; and the data represents the mean + SE of the whole well fluorescence (n = > 15 particles).

RAA EAA GAA

pH 9

pH 7


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Firstly, from figure 48 it can be seen that the expected trends were observed at pH 9 in

which elastase is anionic and was therefore found to penetrate into the RAA particles

and cleave the ECP of RAA significantly more since the fluorescence for dansyl chloride

staining for RAA was 235.82 RFU; whereas the fluorescence was significantly lower for

both EAA (36.70 RFU) and GAA (28.98 RFU) particles (p < 0.05). The average

fluorescence for EAA and GAA particles was not significantly different (p > 0.05) which

is not surprising because previously the selective hydrolysis of the ECPs for both EAA

and GAA were found to be significantly similar i.e. EAA (4.70 %) and GAA (4.64 %) at

pH 9 (figure 44 in section 4.41).

Just like pH 8, un-expectantly figure 48 shows that the fluorescence for dansyl chloride

staining at pH 7 was significantly more for RAA particles (245.35 RFU) compared to

both the EAA particles (77.99 RFU) and the GAA particles (48.22 RFU). This

suggested that possibly the RAA particles were cleaved more compared to both EAA

and GAA particles. In order to test this, the Kaiser test was carried out before and after

Fmoc-X-Ala-Ala-PEGA1900 particles were treated with elastase. From figure 49 it was

observed that all ECPs: RAA, EAA and GAA were fully coupled to unmodified

PEGA1900 particles given that a yellow colour was observed for the Kaiser test for the un-

cleaved Fmoc-X-Ala-Ala-PEGA1900 particles. Additionally, the Kaiser test confirmed

that in its cationic form (i.e. at pH 7) elastase successfully cleaved the ECP of the EAA

particles since a prominent blue adducted was observed for the cleaved EAA particles.

In contrast, the Kaiser test revealed that the cleaved forms of RAA and GAA particles

appeared partly greyish-green this indicated that the ECPs of RAA and GAA were

cleaved to a lesser degree than EAA particles.

Figure 49. Treating both un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles (0.1 M, pH 7.0) with Kaiser test. Fmoc-X-Ala-Ala-PEGA1900 particles: 1: Un-cleaved RAA, 2: cleaved RAA, 3: Un-cleaved EAA, 4: Cleaved EAA, 5: Un-cleaved GAA and 6: cleaved GAA. Elastase in its cationic form was found to cleave EAA (4: blue colour) compared to RAA and GAA (2 and 6: greyish-green colour).

1 2 3 4 5 6


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Next, it was envisaged that possibly the Arg residues were localised by dansyl chloride at

pH 7. In this way, the problem was further investigated by staining unmodified and

ionic PEGA1900 particles with dansyl chloride. The ionic particles were coupled with

Fmoc-Arg(+) (positively charged) and Fmoc-Glu(–) (negatively charged). From figure 50,

it was observed that the unmodified PEGA1900 particles were found to fluoresce

significantly more (9.50 RFU) than both the Fmoc-Arg(+)-PEGA1900 particles (6.24 RFU)

and the Fmoc-Glu(–)-PEGA1900 particles (1.00 RFU). This was expected because

unmodified PEGA particles contain a lot more free amino groups (NH2) compared to

functionalised PEGA1900 particles as the amine groups of these particles are protected by

a terminal Fmoc-group which cannot be stained by dansyl chloride.

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Figure 50. Comparing the pixel fluorescence intensity at pH 7 (0.1 M) after unmodified and ionic PEGA1900 particles were stained with dansyl chloride. Unmodified PEGA1900 particles were found to fluorescence significantly more because these particles contained more free amine groups compared to ionic PEGA1900 particles. The amine groups of the ionic PEGA1900 particles (Fmoc-Arg(+)-PEGA and Fmoc-Glu(-)-PEGA) are protected by the terminal Fmoc-group during SPPS causing the fluorescence to decrease. An increase in fluorescence was observed for Fmoc-Arg(+)-PEGA particles compared to Fmoc-Glu(-)-PEGA particles. This is because there is an effect of localised pH on the dansyl group with the Arg residues for Fmoc-Arg(+)-PEGA particles. Length of scale bar for: PEGA (69 µm), Fmoc-Arg(+)-PEGA (73.24 µm) and Fmoc-Glu(-)-PEGA (62.84 µm). Data represents the mean + SE of the whole well fluorescence (n = 10 particles).


163

Studies have shown that the dansyl chloride reaction with peptides and amino acids is

susceptible to hydrolysis depending on the pH. It has been reported that the rate of

hydrolysis for dansyl chloride reaction is stable up to pH 9.5 (Gros and Labouesse 1969;

Walker 1996). However, other studies have reported that in order to successfully react

free amine groups with dansyl chloride, then the dansyl chloride reaction must be carried

out at a pH > 8.5 (Gray 1967) or more specifically in the range of pH 9.5 – 10.5 (Walker

1996). This is because if the pH increases above this pH range then the hydrolytic

reaction for dansyl chloride increases via base catalysed reaction (Gros and Labouesse

1969). Therefore, since the Arg side chain has a pKa of 12.48

this means that it provides a basic environment as a result it is possible that the

hydrolytic reaction increased via base catalysed reaction which consequently hydrolysed

the dansyl chloride reagent (Gros and Labouesse 1969). In this manner, it is most likely

that the Fmoc-group was hydrolysed via OH– catalysed reaction which therefore caused

the fluorescence of the Arg containing particles i.e. Fmoc-Arg(+)-Ala-Ala-PEGA1900

particles and Fmoc-Arg(+)-PEGA1900 particles to increase.

Moreover, at the required pH range (pH > 8.5 – 10. 5) the N-terminal amino group

remains in the unprotonated i.e. NH2 form and this facilitates the labelling reaction with

dansyl chloride (Walker 1996; Gros and Labouesse 1969). In contrast, the unreactive

protonated form of an N-terminal amino group (NH3+) could potentially alter the pH

value outside the required pH range of the dansyl chloride reaction (Walker 1996) or

slow the labelling reaction down if the pH is lower than pH 8 (Sandhu and Robbins

1989). Since the labelling reaction for dansyl chloride was slow at pH 7 and pH 8, this

explains why the fluorescence of the Glu(–) containing particles i.e. Fmoc-Glu(–)-Ala-Ala-

PEGA1900 particles (figures 44 and 46) and Fmoc-Glu(–)-PEGA1900 particles (figure 48)

was found to decrease. Overall, it was found the dansyl chloride approach was not

suitable for assessing the diffusion of elastase into PEGA particles containing both acidic

and basic groups and for that reason this method was abandoned.

4.4.2.2 Accessibility of a FITC-labelled dextran into PEGA particles

As the aim of this thesis involved designing a wound dressing that would selectively

remove elastase by entrapping it inside functionalised PEGA1900 particles, this section

investigated whether a FITC-labelled dextran with a molecular weight as low as 20 kDa


164

could access Fmoc-X-Ala-Ala-PEGA1900 particles at high ionic strength after these

functionalised PEGA1900 particles had been treated with elastase. The 20 kDa FITC-

labelled dextran was chosen as its molecular weight was lower than the molecular weight

of elastase. Firstly, the molecular weight of elastase was confirmed by SDS-page, and the

active form of PPE had a molecular weight of 25.9 kDa (figure 68: appendix III).

Consequently, this suggests that elastase has the ability of penetrating into PEGA1900

particles since these particles have a molecular weight cut-off of 35 kDa (Kress et al.

2002; Polymer Laboratories Ltd). Nevertheless, the accessibility of proteases into PEGA

particles can vary owing to the swelling of PEGA particles depending on the ionic

strength in addition to the selective hydrolysis of ECPs by protease.

Un-cleaved and cleaved PEGA1900 particles (5 mg of RAA, EAA and GAA) were washed

and swollen in 0.1 M potassium phosphate buffer at pH 8 and they were then incubated

with the 20 kDa FITC-labelled dextran in which its accessibility into the PEGA1900

particles was monitored using TPM. For the un-cleaved PEGA1900 particles, it can be

seen from the pixel intensity graphs that within 10 minutes the 20 kDa FITC-labelled

dextran diffused into the charged PEGA particles (RAA and EAA) more compared to

neutral particles (GAA) as the fluorescence was found to increase from 0 – 10 minutes

(figure 51: green).

However, after elastase cleaved the ECP of the Fmoc-X-Ala-Ala-PEGA1900 particles it

was expected that the molecular accessibility of the cleaved Fmoc-X-Ala-Ala-PEGA1900

PEGA particles decreased by limiting the amount of the 20 kDa FITC-labelled dextran

diffusing into the cleaved PEGA particles. From the pixel intensity graph the

fluorescence for the RAA particles was to found to increase (figure 51: top, red) this

showed that the 20 kDa FITC-labelled dextran penetrated into the cleaved RAA particles

at 0.1 M. In contrast, the accessibility of the 20 kDa FITC-labelled dextran into the

cleaved EAA particles was found to reduce (figure 51: middle, red). These observations

were expected seeing as elastase in its cationic form has the preference of selectively

cleaving the negative ECP (EAA) more than the positive ECP (RAA) as previously

observed in figure 44 (HPLC analysis). Lastly, it was seen that the 20 kDa FITC-labelled

dextran was unable to penetrate into the GAA particles after they had been treated with

elastase (figure 51: bottom, red). Additionally, there was no difference in the accessibility

of the 20 kDa FITC-labelled dextran into both the un-cleaved and cleaved GAA


165

particles, as the fluorescence intensity for the un-cleaved and cleaved GAA particles was

the same (figure 51: bottom).

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Figure 51. TPM micrographs and pixel intensity graphs demonstrating the penetration of a 20 kDa FITC-labelled dextran into both un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles at 0.1 M (pH 8). Un-cleaved Fmoc-X-Ala-Ala-PEGA1900 particles were treated with a 20 kDa FITC-labelled dextran and its penetration into PEGA particles was monitored via TPM at 0 minutes (un-cleaved: 0 min; black:�) and up to 10 minutes (un-cleaved: 10 min; green:�). After Fmoc-X-Ala-Ala-PEGA1900 particles were cleaved with elastase (cleaved) the penetration of the 20 kDa FITC-labelled dextran was again monitored up to 10 minutes (cleaved: 10 min; red:�). Scale bars are either 75 µm or 100 µm (as indicated); and the data represents the pixel intensity across the diameter of each particle.

Seeing as the 20 kDa FITC-labelled dextran did not diffuse into both un-cleaved and

cleaved Fmoc-X-Ala-Ala-PEGA1900 particles to a high degree at 0.1 M, it was predicted

that possibly the swelling of the functionalised PEGA1900 particles at high ionic strength

(0.1 M) hindered the complete accessibility of the 20 kDa FITC-labelled dextran into the

central-core of both un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles.

Consequently, comparative studies were then carried out using unmodified PEGA

particles to investigate the difference in the penetration of the 20 kDa FITC-labelled

dextran into the PEGA1900 particles at 0.1 M and 0.001 M (figure 52). Unmodified

PEGA1900 particles were first washed and then swollen in potassium phosphate buffer at

Un-cleaved: 0 min

Un-cleaved: 10 min

Cleaved: 10 min

RAA

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GAA


166

0.1 M and 0.001 M (pH 8.0) for 5 minutes. After that the unmodified PEGA1900 particles

were incubated with FITC-labelled 20 kDa dextran at the appropriate ionic strength (as

indicated), the penetration of the FITC-labelled dextran was monitored via TPM up to

10 minutes. Figure 52 clearly illustrates that the 20 kDa FITC-labelled dextran

significantly penetrated into unmodified PEGA1900 particles at 0.001 M in contrast to

0.1M.

Figure 52. TPM micrographs (left and middle) and pixel intensity graphs (right) demonstrating the accessibility and diffusion of FITC-labelled 20 kDa dextran into unmodified PEGA1900 particles at an ionic strength of 0.1 M and 0.001 M for the duration of 10 minutes. Unmodified PEGA1900 particles were swollen in 0.1 M or 0.001 M potassium phosphate buffer (pH 8.0) and then incubated with a 20 kDa FITC-labelled dextran (1 mg/ml: pH 8 at 0.1 M or 0.001 M). Without a doubt the 20 kDa FITC-labelled dextran penetrated PEGA1900 particles significantly more at 0.001 M compared to 0.1 M (p < 0.05). In the pixel intensity graphs 0 min is indicated by black diamonds (�) and 10 min by green squares (�). Scale bars are 75 µm (0.1 M) and 100 µm (0.001 M); and the data represents the pixel intensity across the diameter of each particle.

Accordingly, the penetration of the 20 kDa FITC-labelled dextran into Fmoc-X-Ala-Ala-

PEGA1900 particles was repeated at 0.001M. Evidently, figure 53 shows that for the un-

cleaved Fmoc-X-Ala-Ala-PEGA1900 particles the 20 kDa FITC-labelled dextran

penetrated into RAA particles more compared to both EAA and GAA particles. Un-

expectantly the 20 kDa FITC-labelled dextran was found to penetrate GAA more

compared to EAA. This is because FITC has a pKa of 5.93 or 6.5 (Moding et al. 2009;

Marchetti et al. 2009) meaning that it is negatively charged at pH 8 and for that reason

the 20 kDa FITC-labelled dextran was unable to penetrate into un-cleaved EAA particles

due to electrostatic repulsion between the negatives charges for both FITC and EAA.

0.1 M

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For the cleaved Fmoc-X-Ala-Ala-PEGA1900 particles, after 10 minutes the accessibility of

the 20 kDa FITC-labelled 20 kDa dextran became restricted for both RAA and GAA

since the fluorescence within the pixel intensity graphs was found to decrease for both of

these PEGA1900 particles (figure 53: red). This is because the molecular accessibility of

RAA and GAA PEGA1900 particles became reduced after elastase cleaved the ECP for

both the positive and neutral Fmoc-X-Ala-Ala-PEGA1900 particles. However, it can be

seen that the molecular accessibility of RAA was profoundly reduced compared to GAA,

which is expected since elastase was previously found to selectively hydrolyse the ECP of

the RAA particles significantly more than the GAA particles (figure 53: pH 8, p < 0.05).

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Figure 53. TPM micrographs and pixel intensity graphs demonstrating the penetration of a 20 kDa FITC-labelled dextran into both un-cleaved and cleaved Fmoc-X-Ala-Ala-PEGA1900 particles at 0.001M (pH 8). Un-cleaved Fmoc-X-Ala-Ala-PEGA1900 particles were treated with 20 kDa FITC-labelled dextran and its penetration into PEGA particles was monitored via TPM at 0 minutes (un-cleaved: 0 min; black:�) and up to 10 minutes (un-cleaved: 10 min; green:�). After Fmoc-X-Ala-Ala-PEGA1900 particles were cleaved with elastase (cleaved) the penetration of the 20 kDa FITC-labelled dextran was again monitored up to 10 minutes (cleaved: 10 min; red:�). Scale bars are either 75 µm or 100 µm (as indicated); and the data represents the pixel intensity across the diameter of each particle.

Additionally, it can be deduced that after elastase cleaved the ECP of the RAA particles,

removal of the positively charged Arg(+) residues caused the pores of the RAA particles

to collapse and therefore the accessibility of the FITC-labelled 20 kDa dextran became

Un-cleaved: 0 min

Un-cleaved: 10 min

Cleaved: 10 min

RAA

EAA

GAA


168

limited within 10 minutes. Furthermore, it can be concluded that if the accessibility of a

FITC-labelled dextran with a molecular weight as low as 20 kDa becomes restricted after

the positive particles were cleaved by elastase, subsequently due to polymer collapse this

confirms that elastase becomes trapped inside the RAA particles as it has a molecular

weight of 25.9 kDa which is greater than 20 kDa.

As mentioned above, the FITC-labelled 20 kDa dextran did not penetrate un-cleaved

EAA particles. Interestingly, once the ECP of the EAA particles was cleaved by elastase,

the FITC-labelled 20 kDa dextran was found to slightly penetrate the EAA particles as

the fluorescence within the pixel intensity graph was found to increase slightly (figure 53:

middle, red). This is because some of the Glu residues, hence negative charges of the

EAA particles were removed by elastase causing the negatively charged FITC-labelled

dextran (at pH 8) to diffuse into the cleaved EAA particles. It is not surprising that only

a small fraction of FITC-labelled 20 kDa dextran penetrated into the cleaved EAA

particles since previously in chapter 3 the swelling of cleaved EAA particles was found to

decrease at 0.001 M (figure 37).

4.4.2.3 FITC-elastase

The penetration of FITC-labelled dextran of known molecular weight into both un-

cleaved and cleaved PEGA particles can be considered as an indirect approach of

visualising the penetration followed by entrapment of the enzyme into PEGA particles.

Therefore, in order to directly visualise the penetration of elastase into Fmoc-X-Ala-Ala-

PEGA particles under real-time conditions, elastase was labelled with FITC to generate

FITC-elastase in PBS buffer (0.15 M and pH 7.4). Figure 54 depicts the TPM

micrographs along with the average fluorescence intensity graph for the accessibility and

penetration of FITC-elastase into RAA, EAA and GAA particles. FITC-elastase was

found to penetrate the RAA and EAA significantly more compared to GAA (p < 0.05).

However, since FITC-elastase existed in its cationic form, as expected FITC-elastase was

highly proficient in accessing and cleaving the ECP of EAA particles more quickly

compared to both RAA and GAA (p < 0.05).. The ECP of EAA particles was cleaved

from the inner-core of PEGA within 2 minutes (101.05 RFU) and there was no

significant difference in the fluorescence between 2 – 15 minutes for the total reaction

time (i.e. 101.05 – 104.65 RFU).


169

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Figure 54. TPM micrographs (top) and the average fluorescence graph (bottom) depicting the real-time penetration of FITC-elastase into Fmoc-X-Ala-Ala-PEGA1900 particles in PBS buffer (0.15 M and pH 7.4). Sequential cross-sectional equatorial images of functionalised PEGA particles were obtained over the course of 15 minutes (as indicated). The fluorescence was found to increase for the charged particles, this means that FITC-elastase was encapsulated by both RAA and EAA particles to a greater extent compared to GAA particles. Since elastase predominantely in its cationic form under these conditions, it selectively cleaved and became trapped into EAA particles within two minutes compared to RAA particles (x minutes). For the neutral particles, GAA (light grey) over the course of time the fluorescence inside the PEGA1900 particles was found to decrease illustrating that the GAA particles were unable to retained elastase. Scale bar represents 75 µm; and the data represents the mean + SE of fluorescence across the diameter of each particle.

It was difficult to monitor the actual time required for FITC-elastase to cleave the ECP

from the outer surface of the EAA particles. As soon as a drop of FITC-elastase was

added to EAA particles and straight after that the TPM was quickly placed into focus it

0 min 2.0 min 15 min 7.5 min 5.0 min 0.5 min

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170

can be seen that majority of the ECP had already been cleaved by FITC-elastase (figure

54: 0 min). In contrast, at 2.0 minutes the fluorescence intensity of RAA was 84.11 RFU

and for GAA it was 102.55 RFU (figure 54). Compared to the charged particles (RAA

and EAA) the fluorescence intensity for the penetration of FITC-elastase inside the

GAA particles was found to decrease over time from 124.25 RFU at time zero to 73.25

RFU at 15 minutes.

Previously in chapter 3, it was shown that the swelling of GAA at high ionic strength i.e.

0.2 M was much lower than the charged PEGA1900 particles. It was envisaged that

possibly the swelling due to the high ionic strength restricted the accessibility of FITC-

elastase into GAA. Subsequently, the penetration of FITC-elastase into GAA particles

was repeated at 0.1 M (pH 7.0) and monitored up to 30 minutes.

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FU

)

Figure 55. TPM micrographs (top) and the average fluorescence graph (bottom) depicting the real-time penetration of FITC-elastase into GAA particles at 0.1 M (pH 7.0). Sequential cross-sectional equatorial images of the GAA particles were obtained over the course of 30 minutes (as indicated). As the fluorescence for the penetration of FITC-elastase was found to decrease over the course of time this indicated that the GAA particles were unable to retain and entrap elastase. Scale bar represents 75 µm; and the data represents the mean + SE of fluorescence across the diameter of each particle.

Figure 55 demonstrated that the fluorescence intensity was again found to decrease

inside the GAA particles over time from 101.47 RFU at time zero to 28.53 RFU at 30

minutes. Therefore, it was concluded that although elastase has the ability to cleave the


171

neutral ECP possibly from the surface of the GAA particles, elastase is not retained by

GAA particles. This observation is further confirmed by the fluorescence assay whereby

sample fluids containing elastase are treated with Fmoc-X-Ala-Ala-PEGA1900 particles at

0.1 M (at pH 7-8: figure 57) and at 0.15 M (PBS buffer, at pH 7.4: figure 65). In each

case, elastase activity is found to be reduced by the charged particles (RAA and EAA)

compared to the neutral particles (GAA). The latter particles are found not to reduce

elastase activity at pH 7 – 8 (0.1 – 0.15 M). Detailed descriptions of these observations

are given in section 4.4.2.4 (figure 57) and chapter 5 (figure 65).


In this section the elastase activity in sample fluids mimicking the environment of

chronic wounds before and after the sample fluids had been treated with functionalised

PEGA particles was examined. This was achieved by enzyme kinetics which involved

measuring the initial rates of elastase activity of PPE using the fluorescence substrate

MeOSuc-Ala-Ala-Pro-Val-AMC.

Firstly, it was essential to optimise the initial rates of elastase-catalysed reactions of PPE

and to determine the Michaelis-Menten constant (KM). In enzyme kinetics, KM is an

important parameter, it is the concentration of a substrate when the active site of an

enzyme is half filled and therefore yields half maximal velocity wherein significant

enzyme catalysis takes place (Stryer et al. 2002). Additionally, the KM directly influences

the fluorescence intensity of the enzyme-catalysed reaction depending on the

environment of the substrate under the influence of pH, ionic strength and temperature

(Stryer et al. 2002). These parameters were selected according to the environment

observed in chronic wounds and depending on the optimal conditions for elastase.

Firstly, elastase was dissolved at a concentration of 0.01 mg/ml as this concentration of

elastase is observed in chronic wounds (as reported previously in chapter 2). Secondly,

given that elastase is optimal at pH 8 (Worthington Biochemical Corporation), the

fluorescence substrate assay was optimised at this pH wherein both the fluorescence

substrate and elastase were both dissolved in potassium phosphate buffer at pH 8 and

0.1 M. Finally, to mimic the temperature observed in chronic wounds the fluorescence

substrate assay was carried out in the range of 34 – 37oC. The fluorescence substrate was

dissolved in buffer within the range of 0.00 – 1.00 mM and within this range elastase


172

activity of PPE was found to obey Michealis-Menten kinetics as a hyperbolic curve was

observed (figure 56a) which is derived from Michaelis-Menten equation (see equation 3:

figure 56a).

(a)

Equation 3. Michaelis-Menten equation:

(b)

Equation 4. Derivation of the Michaelis-Menten equation:

y = 0.0000297x + 0.0002262

R2 = 0.992

-0.0004

-0.0002

0.0000

0.0002

0.0004

0.0006

0.0008

0.0010

-20 -10 0 10 20 30

1/[S] (mM-1)

1/V

o

Figure 56. Optimisation of the fluorescence substrate assay using Michaelis-Menten kinetics. Elastase (PPE, 0.01 mg/ml) and the fluorescence substrate MeOsuc-Ala-Ala-Pro-Val-AMC (0 – 1.00 mM) were dissolved in potassium phosphate buffer at pH 8 (0.1 M). The increase in fluorescence was measured after elastase cleaved the fluorescence substrate releasing AMC over the course of 14 hours. The initial velocity (Vo) for the elastase activity of PPE at various concentrations of the fluorescence substrate (0 – 1.00 mM) was found to obey Michaelis-Menten kinetics as a hyperbolic curve was observed (a). From the Lineweaver-Burk plot (b) the kinetic constants: Vmax (4420.87 µmol hr-1) and KM (0.133 mM) were calculated.

Vo = Vmax[S]

+[S] KM

Vo Vmax

K M1=

Vmax[S]

1+. 1

0

1125

2250

3375

4500

0.00 0.25 0.50 0.75 1.00

[S] (mM)

Vo

(µm

ol h

r-1)


173

From figure 56a it can be seen that at constant concentration of elastase (0.01 mg/ ml)

increasing the concentration of the fluorescence substrate was found to increase the

velocity of the enzyme reaction. As a result this caused the fluorescence for the elastase

activity of PPE to increase at which point the elastase reaction was at first order (Zubay

et al. 1995). Eventually, it was found that increasing the concentration of the

fluorescence substrate caused the elastase reaction of PPE to reach zero order whereby

the elastase reaction became saturated. This is most commonly referred to as Vmax i.e.

maximum velocity (Stryer et al. 2002). Vmax is the point at which elastase had

completely complexed with the fluorescence substrate to form the enzyme substrate (ES)

complex i.e. elastase/ MeOSuc-Ala-Ala-Pro-Val-AMC and at this point the products (see

figure 43) are constant (Stryer 1995; Zubay et al. 1995). Due to the solubility of the

fluorescence substrate, high concentrations of the fluorescence substrate were avoided in

the range of > 2.50 – 5.00 mM as Michaelis-Menten kinetics was not obeyed since the

elastase activity of PPE was found to decrease. Seeing as it was impossible to calculate

Vmax from the hyperbolic plot, the Lineweaver-Burk plot (figure 56b) was used along

with equation 4 (given in figure 56b) which is the derivation of the Michaelis-Menten

equation (equation 3) to determine both the kinetic constants: Vmax and KM. These

were calculated as 4420.87 µmol hr-1 (Vmax) and 0.133 mM (KM). For subsequent

experiments, 0.133 mM was used as the optimal substrate concentration for the

fluorescence assay.

Next the reduction of elastase activity of PPE was examined when sample fluids

containing elastase were treated with Fmoc-X-Ala-Ala-PEGA1900 particles. Sample fluids

were prepared by dissolving elastase (0.01 mg/ml) in 0.1 M potassium phosphate buffer

at pH 7 – 9. Subsequently, after SPPS Fmoc-X-Ala-Ala-PEGA1900 particles were washed

with H2O and then with 0.1 M potassium phosphate buffer at the appropriate pH (as

indicated). After that, Fmoc-X-Ala-Ala-PEGA1900 particles were transferred (10 mg) to

individual centrifuges vials and then swollen in 0.1 M potassium phosphate buffer (at pH

7 – 9; 50 µl) for 5 minutes. Next, sample fluids containing elastase were treated with the

pre-washed and swollen Fmoc-X-Ala-Ala-PEGA1900 particles (RAA, EAA and GAA)

over the course of 20 minutes. At each time point, the elastase reaction was stopped and

the supernatants were examined for the residual elastase activity of PPE in the sample

fluids after they had been treated with Fmoc-X-Ala-Ala-PEGA1900 particles using the

fluorescence substrate assay.


174

(a)

30

60

90

120

150

180

0 5 10 15 20

Time (minutes)

Ra

te (

RF

U h

r-1)

0

3

6

9

12

15

0 5 10 15 20

Time (minutes)

Rat

e (

RF

U h

r-1/

mg

)

(b)

30

60

90

120

150

180

0 5 10 15 20Time (minutes)

Rate

(R

FU

hr-1

)

0

3

6

9

12

15

0 5 10 15 20Time (minutes)

Rat

e (R

FU

hr-1

/m

g)

(c)

30

60

90

120

150

180

0 5 10 15 20

Time (minutes)

Rat

e (

RF

U h

r-1)

0

3

6

9

12

15

0 5 10 15 20

Time (minutes)

Rate

(R

FU

hr-1

/m

g)

RAA EAA GAA

Figure 57. Rates of residual elastase activity of PPE after sample fluids containing elastase were treated with Fmoc-X-Ala-Ala-PEGA particles at 0.1 M and at various pH values: pH 7 (a), pH 8 (b) and pH 9 (c). Elastase (PPE: 0.01 mg/ml) was incubated with Fmoc-X-Ala-Ala-PEGA particles over the course of 20 minutes. At each time point the residual elastase activity was measured using the fluorescence substrate assay. Left plots: demonstrate the initial rates from elastase-substrate reaction versus the time in which elastase was incubated with functionalised PEGA particles. Right plots: demonstrate the initial rates of elastase activity per mass of functionalised PEGA particles. Data represents the mean + SE of two measurements (n = 2).

Figure 57 shows the elastase activity of PPE in sample fluids (at time zero) and the

residual elastase activity of PPE after the sample fluids were treated with Fmoc-X-Ala-


175

Ala-PEGA1900 particles over the course of 20 minutes. The initial rates of residual

elastase activity of PPE are expressed as RFU hr-1 (figure 57a-c: left side) and according

the initial rates of the residual elastase activity of PPE in the sample fluids were corrected

according to the mass of Fmoc-X-Ala-Ala-PEGA1900 particles used, in which the initial

rates were expressed as Rate/ mass (RFU hr-1/mg, figure 57a-c: right side).

It was established that elastase activity of PPE at each pH varied (figure 57) and as

expected the initial rate elastase activity of PPE was optimal at pH 8 (161.75 RFU hr-1)

compared to pH 7 (105.36 RFU hr-1) and pH 9 (129.48 RFU hr-1). When elastase was in

its cationic form at pH 7-8, collectively on average the elastase activity of PPE was found

to reduce more by the EAA (pH 7: 57.89 RFU hr-1 and pH 8: 105.07 RFU hr-1)

compared to both RAA (pH 7: 70.22 RFU hr-1 and pH 8: 126.417 RFU hr-1) and GAA

(pH 7: 100.40 RFU hr-1 and pH 8: 127.48 RFU hr-1) PEGA particles (p > 0.05, figure

57a-b: left). When elastase was in its anionic form at pH 9, although elastase activity of

PPE was collectively reduced more by RAA (average: 114.64 RFU hr-1) compared to

both EAA (average: 122.57 RFU hr-1) and GAA (average: 127.20 RFU hr-1) PEGA

particles (figure 57c: left) on the contrary the initial rates of elastase activity for PPE were

not significantly different between RAA, EAA and GAA PEGA particles (p > 0.05).

When taking the mass of Fmoc-X-Ala-Ala-PEGA1900 particles into account, the residual

elastase activity of PPE in samples fluids after they had been treated with Fmoc-X-Ala-

Ala-PEGA particles were compared (figure 57a-c: right). The elastase activity was

reduced by: 21.26 – 35.68 % at pH 7; 20.83 – 43.70 % at pH 8 and 6.95 – 29.79 % at pH

9 (5 – 20 minutes) after samples fluids were treated with EAA particles. In contrast,

when samples fluids were treated with RAA particles, the elastase activity of PPE was

reduced by -5.92 – 13.55 % at pH 7; 13.75 – 27.06 % at pH 8 and 10.57 – 36.06 % at pH

9 (5 – 20 minutes). Although, the residual elastase activity of PPE remained fairly

constant for the GAA particles at pH 7 – 8, within this pH range the level of reduction

for elastase activity of PPE was very small i.e. 6.04 – 2.74 % at pH 7 and 7.03 – 6.43 % at

pH 8 (5 – 20 minutes); whereas at pH 9 it was found to increase from 6.34 – 31.22 (5 –

20 minutes). Overall, from figure 55 it was deduced that the residual cationic form of

elastase activity for PPE in samples fluids was highly reduced by EAA particles at pH 7 –

8; whereas the residual anionic form of elastase activity for PPE in sample fluids was

reduced by all Fmoc-X-Ala-Ala-PEGA1900 particles (RAA, EAA and GAA) at pH 9.


176

4.5 CONCLUSION

The results in this chapter demonstrate the accessibility and diffusion of elastase into

Fmoc-X-Ala-Ala-PEGA1900 particles at 0.1 M under the influence of pH by exploiting the

used of various fluorophores (dansyl chloride, FITC-labelled dextran and FITC). It was

found that assessing the diffusion of elastase by staining cleaved PEGA1900 particles with

dansyl chloride was not appropriate method for PEGA1900 particles containing both

acidic and basic amino acids due to the effect of localised pH on the dansyl chloride

group. The FITC-labelled dextran studies showed that elastase significantly penetrated

into RAA particles more compared to the GAA particles. Additionally the FITC-labelled

dextran showed that after the CMR was removed from the charged PEGA1900 particles

the pores of PEGA1900 particles became restricted as a 20 kDa FITC-labelled dextran was

unable to penetrate into PEGA1900 particles. This demonstrated that since elastase has a

molecular weight of 25.9 kDa it became trapped inside the PEGA1900 particles due to the

collapsing of the pores within the PEGA1900 particles.

Subsequently, for real-time monitoring elastase was labelled with FITC to demonstrate

the direct penetration and entrapment of FITC-elastase into PEGA particles in PBS at

physiological pH and ionic strength (at pH 7.4 and 0.15 M, respectively). FITC-elastase

was studied in its cationic form and was found to penetrate and eventually became

trapped into both EAA and RAA particles more compared to GAA particles. Cationic

FITC-elastase was found to totally cleave the ECP of the EAA particles within 2 minutes

and subsequently FITC-elastase became trapped into EAA particles within 2 minutes. In

contrast, the neutral particles were unable to retain and encapsulate FITC-elastase at 0.1 -

0.15 M as the fluorescence inside the GAA particles was found decrease.

Finally, it was demonstrated that Fmoc-X-Ala-Ala-PEGA1900 particles were found to

reduce elastase activity of PPE after sample fluids containing elastase were treated with

Fmoc-X-Ala-Ala-PEGA1900 particles. At high ionic strength i.e. 0.1 M the residual

elastase activity of PPE in sample fluids was reduced more by the charged PEGA

particles (RAA and EAA) compared to neutral particles (GAA) within the pH range of 7

– 8, although all Fmoc-X-Ala-Ala-PEGA1900 particles were found to reduce elastase

activity of PPE at pH 9.

CHAPTER 5: Reduction of Fibroblast Elastase Activity by PEGA Particles

177

CCHHAAPPTTEERR 55

RReedduuccttiioonn ooff FFiibbrroobbllaasstt EEllaassttaassee AAccttiivvii ttyy bbyy PPEEGGAA PPaarrttiicclleess


178

5.1 INTRODUCTION

In chapter 1, skin ulcers were reported as the largest and most frequently occuring types

of chronic wounds (Eaglstein and Falanga 1997; Stadelmann et al. 1998). Previously in

chapter 2 we encountered that there are various human proteases that express

elastinolytic activity (for examples see table 4). Neutrophils and macrophages were

reported as the two major cells that express elastase(s) during the wound healing process.

However, since the late eighties skin fibroblasts, specifically human dermal fibroblasts

(HDF) have been reported to express elastase activity (Homsy et al. 1988; Croute et al.

1991; Tsukahara et al. 2001; Tsuji et al. 2001; Isnard et al. 2002; Suganuma et al. 2010).

For that reason, in this chapter HDF were chosen as the cells to study the elastase

activity expressed inside or outside the cell. Elastase or elastase activity that is expressed

intracellularly or extracellularly by fibroblast is sometimes referred to as human skin

fibroblast elastase activity (Homsy et al. 1988), elastase-type activity (Isnard et al. 2002) or

skin fibroblast elastase (Suganuma et al. 2010). In this thesis, it is designated as HDF-

elastase and the elastase activity as HDF-elastase activity. Interestingly, some

investigators have shown elastase localised at the surface of fibroblasts in which elastase

binds to receptors on the surface of fibroblasts (Cunningham et al. 1986; Campbell and

Cunningham 1987) and the research carried out by Robert and colleagues shows elastase

activity bound to the membrane of dermal fibroblasts (Archillas-Marcos and Robert

1993; Beranger et al. 1994; Isnard et al. 2002). The class of enzymes responsible for this

elastinolytic activity by fibroblasts has been stated as an endopeptidase of MMP origin, a

metalloendopeptidase (Homsy et al. 1988) such as the metalloelastase-type protease:

MMP-2 and MMP-9 (Beranger et al. 1994; Isnard et al. 2002) as well as a serineprotease

(Schmidt et al. 2009).

Fibroblasts are considered as the most important cell type during the wound healing

process for the reason that proper healing requires the migration and proliferation of

fibroblasts into the wound bed. The prominent role of fibroblast is laying down and re-

organising the ECM by synthesising and secreting high levels of newly synthesised

collagen (main component of the ECM) alongside other ECM components e.g. elastin,

fibronectin, glycoaminoglycans, proteoglycans, growth factors (Singer and Clark 1999;

Porter 2007; Schultz and Mast 1998; Dasu et al. 2003) in the attempt of replacing and

regenerating the provisional matrix of the fibrin clot. Fibroblast successfully achieve this


179

by controlling the levels of MMPs which degrade the old ECM and TIMPs (Mutsaers et

al. 1997; Enoch and Harding 2003; Schultz and Mast 1998). Eventually, fibroblasts

generate a compact and contracting-ECM around occupying cells (Singer and Clark 1999;

Porter 2007; Lauffenburger and Wells 2003). A detailed description for the function of

fibroblasts during the wound healing process was previously covered in chapter 2

(section 2.2).

Chronic wounds are known to exhibit elevated levels of various pro-inflammatory

cytokines such as IL-1α (Barone et al. 1998), IL-1β and TNF-α (Tarnuzzer and Schultz

1996; Trengrove et al. 2000) and under in-vitro conditions cultured normal human

fibroblast cells have been stimulated with the cytokine, IL-1β to produce elastase activity

(Croute et al. 1991). IL-1β is a polypeptide, a potent pleiotropic pro-inflammatory

cytokine that mediates processes in host defense, inflammation and in-response to injury.

In healing wounds, even though activated monocytes/macrophages are considered as the

main source for IL-1β, fibroblasts alongside other cells are reported to produce or

secrete IL-1β (Manninen et al. 1992; Robson et al. 1994; Robbins et al. 1999; Chung

2001). TNF-α self-regulates its own synthesis by macrophages and then macrophages

and fibroblasts enhance the generation of IL-1β (Mast and Schultz 1996; Bryant and Nix

2007). Cytokines are stated to be extremely potent and active at picomolar-to-nanomolar

concentrations (Henry and Garner 2003). During the wound healing process binding of

these cytokines to cell surface receptors results in signal transduction wherein IL-1β and

TNF-α act synergistically to exert many functions: stimulate endothelial cells to express

cell adhesion molecules, influence collagen synthesis by fibroblasts by up-regulating the

expression of MMPs, however for the latter role IL-1β is said to be mitogenic for

fibroblast MMP expression (Elias et al. 1989; Chung 2001; Robson 2003; Mast and

Schultz 1996; Schultz and Mast 1998; Bryant and Nix 2007; Robbins et al. 1999; Dasu et

al. 2003). Additionally, IL-1β and TNF-α promote chemotaxis (Schultz et al. 2003)

alongside the migration, increase mitosis and proliferation of fibroblasts (Mast and

Schultz 1996; Robbins et al. 1999; Rumalla and Borah 2001; Chung 2001; Robson 2003;

Henry and Garner 2003) and are known to downregulate the expression of TIMPs (Mast

and Schultz 1996; Schultz and Mast 1998; Bryant and Nix 2007).

When designing a wound dressing (i.e. functionalised PEGA particles) it is important to

assess the surface of the wound dressing in the presence of cells. The biocompatibility


180

of PEGA has previously been addressed by Ulijn and co-workers in which PEGA

surfaces and particles were demonstrated as being biocompatible with the growth of

both fibroblast and osteoblast cells (Zourob et al. 2006; Patrick 2010; Todd et al. 2007).

However, for the clinical use of functionalised PEGA particles as a chronic wound

dressing for mopping-up elastase it is essential to examine the efficacy of the

functionalised PEGA particles under in-vitro conditions to reduce/ remove levels of

elastase that is expressed both outside (extracellular) or inside (intracellular) HDF cells

within sample fluids mimicking the environment of a wound exudate. This chapter

highlights the reduction of HDF-elastase activity from HDF cells by functionalised

PEGA particles at physiological pH and ionic strength.

5.2 OBJECTIVES Having demonstrated in chapters 3 and 4, the recognition of ECPs by elastase followed

by its accessibility into functionalised PEGA particles containing the Ala-Ala peptide at

both low and high ionic strength, accordingly the main focus of this final experimental

chapter was to analyse the reduction of elastase activity that was expressed by HDF cells

under in-vitro conditions at physiological pH and ionic strength (pH 7.4, 0.15 M

respectively). The experimental study was undertaken in two parts: firstly HDF cells

were stimulated with IL-1β to express HDF-elastase both outside or inside (extracellular

or intracellular) HDF cells and this stimulation process was optimised to achieve

detectable levels of HDF-elastase activity from HDF cells; and secondly sample fluids

containing HDF-elastase activity were treated with functionalised PEGA particles (RAA,

EAA, and GAA) and residual HDF-elastase activity in the sample fluids was examined to

demonstrate the mopping-up of HDF-elastase by functionalised PEGA particles.

In the literature it was confusing to understand whether authors made reference to HDF-

elastase or HDF-elastase activity that was expressed outside (extracellular) or inside

(intracellular) fibroblasts. Consequently to avoid confusion in this chapter, the elastase

activity of HDF-elastase expressed outside HDF cells is designated as the ‘HDF-elastase

activity expressed outside HDF cells (extracellular) whereas the elastase activity of the

HDF-elastase expressed inside HDF cells is designated as the HDF-elastase activity

expressed inside HDF cells (intracellular)’.


181


5.3.1 Materials

Dulbecco’s Modified Eagle Medium (DMEM + 1000 mg/L glucose, + GlutaMAX 1, +

pyruvate), Fetal Bovine Serum (FBS), Antibiotic/antimycotic, DPBS (phosphate buffer

saline), DH2O, Trypsin (0.25 % trypsin, 1 mM EDTA.4Na+) were all purchased from

Invitrogen (UK). Human dermal fibroblasts (adult) were from Cascade Biologics

(Invitrogen). IL-1β was supplied by Sigma (UK). Fmoc-protected amino acids (Fmoc-

Ala-OH.H2O, Fmoc-Arg(pbf)-OH, Fmoc-Glu(OtBu)-OH, Fmoc-Gly-OH) and

MeOSuc-Ala-Ala-Pro-Val-AMC were all purchased from Bachem (UK). Porcine

pancreatic elastase (PPE, 3.9 mg per protein) was purchased from Worthington

Biochemical Corporation (USA). Culture flasks, centrifuge tubes, pipettes and 96-well

plates (black) were purchased from Fisher Scientific (UK). Well-plates (48) were

purchased from VMR (UK). Microplate reader FLUORstar OPTIMA was supplied by

BMG LABTECH. Class II laminar flow microbiological cabinets (Bio2+) were from

Envair Ltd.

5.3.2 Methods

5.3.2.1 Fmoc SPPS

Functionalised PEGA particles i.e. Fmoc-tripeptide-PEGA particles with the

configuration Fmoc-X-Ala-Ala-PEGA (abbreviated as RAA, EAA, GAA) were prepared

using the protocol as previously described in chapter 3, section 3.2.2.2. The successful

coupling of amino acids and de-protecting of the Fmoc groups during SPPS was

monitored using the Kaiser test (see section 3.3.2.3).

5.3.2.2 Cell culture of human dermal fibroblast (HDF)

For cell culture studies asceptic techniques were carried out in microbiological laminar

flow cabinets (class II). HDF cells were cultured in DMEM (supplemented with 10%

heat inactivated FBS and 1% (v/v) antibiotic/ antimycotic) and the culture flasks (75

cm2) were incubated at 37oC in a humidified atmosphere containing 95% air and 5%


182

CO2. After every 2 days the DMEM was replaced with fresh DMEM. Near confluent

(> 95%) HDF cells were rinsed with DPBS and then detached from the culture flask by

incubating the HDF cells with trypsin for 5 minutes at 37oC in a humidified atmosphere

containing 95% air and 5% CO2. The bottom of the culture flask was gently tapped to

completely detach all HDF cells from the surface of the culture flask. The HDF cells

were then re-suspended in fresh DMEM to deactivate trypsin activity and then

centrifuged for 5 minutes at 1.5 x 1000 rpm to produce a pellet of HDF cells. The

supernatant was removed by aspiration and the pellet of HDF cells were re-suspended in

fresh DMEM (as described below). Passages 10 – 12 were used for the experiments (as

indicated).

5.3.2.3 Activation of HDF cells by IL-1ß to express HDF-elastase

HDF cells were re-suspended in DMEM i.e. serum-free media (SFM) or supplemented

with either 10% or 25% FBS). When DMEM was supplemented with FBS, in all

experiments heat inactivated FBS was used. For each experiment in a 48 well-plate, 0.5

ml HDF cells (40, 000 cells/ ml) were seeded in each well and incubated at 37oC in a

humidified atmosphere containing 95% air: 5% CO2. IL-1β (25 – 500 nM) was dissolved

in the appropriate DMEM i.e. SFM or supplemented with either 10% or 25% FBS.

After 4 hours, HDF cells were stimulated with IL-1β (0.25 ml at various concentrations)

over the course of 7 days.

After activation of HDF cells by IL-1β, each day the cell culture medium was examined

for HDF-elastase activity expressed outside (extracellular) HDF cells using the protocol

described in section 5.3.2.5. In order to test whether HDF cells expressed elastase

activity inside (intracellular) HDF cells, the HDF cells were lysed to release all their

intracellular contents by freeze-thaw using two approaches. In the first approach, the

incubated DMEM (supplemented with/ without both FBS and IL-1β) was not removed

and the HDF cells were lysed by freeze-thaw directly after the activation process by IL-

1β. The second approach involved removing the incubated DMEM (supplemented

with/ without both FBS and IL-1β) and subsequently the HDF cells were firstly washed

with DPBS (twice) and then incubated with DH2O (0.75 mL) and then lysed by freeze-

thaw. Freeze-thaw involved first freezing HDF cells (incubated with both DMEM and

DH2O) at –80 oC and once frozen they were then thawed at 37oC in a humidified


183

atmosphere containing 95% air: 5% CO2. In order to efficiently lyse the HDF cells the

freeze-thaw procedure was repeated twice. After lysing the HDF cells, it was essential

that the collected lysate was not centrifuged as previous studies reported that the surface

of fibroblasts contains bound elastase (Cunningham et al. 1986; Campbell and

Cunningham 1987). Instead the lysate of HDF cells was thoroughly mixed by pipetting

it in-and-out using a micropipette to efficiently mix both bound and unbound HDF-

elastase and after that the expressed HDF-elastase activity by HDF cells (intracellular)

was monitored each day using the protocol described in section 5.3.2.5.

5.3.2.4 Elastase treated with functionalised PEGA particles

After SPPS, functionalised PEGA particles (i.e. RAA, EAA, GAA) were thoroughly

washed with H2O, DH2O, DPBS and DMEM + 25% FBS. Then PEGA particles (10

mg) were transferred to a microcentrifuge tube and swollen in 50 µl DMEM + 25% FBS.

The positive control, elastase (PPE, 0.01 mg/mL) was also dissolved in DMEM + 25%

FBS. In the microcentrifude tubes, 250 µl of the sample fluid i.e. elastase (PPE) or the

HDF cell lysate was added to the swollen PEGA particles. The microcentrifuge vials

were then mixed vigorously on a vortex for 1 minute; and then centrifuged for 4 and 14

minutes, resulting in an overall incubation time of 5 and 15 minutes, respectively. At

each time point, the supernatant of the sample fluids after they had been treated with

each functionalised PEGA particle (RAA, EAA, and GAA) were tested for the residual

elastase activity in the sample fluids using the protocol described in section 5.3.2.5.


The fluorescence substrate, MeOSuc-Ala-Ala-Pro-Val-AMC was dissolved in 2 ml

methanol and then further diluted with DPBS giving a final concentration of 0.133 mM

(as determined in chapter 4). Elastase assay was carried out by incubating 50 µl of the

substrate with 50 µl of the sample fluid. The sample fluid was either: (1) cell suspension

of the unlysed HDF cells and the HDF cell lysate of the lysed HDF cells treated with/

without IL-1β; and (2) the supernatants collected after the cell suspension and HDF cell

lysate had been treated with each functionalised PEGA particle (RAA, EAA, and GAA).

In this assay, elastase i.e. PPE was used as a positive control and the negative controls

were DMEM, DH2O and IL-1β. The fluorescence of the substrate hydrolysis was


184

measured at λexc 383 nm (excitation) and λem 405 nm (emission) and rate of elastase

activity was expressed as RFU hr-1 and when the latter was corrected according to the

mass of PEGA particles used, the rate of elastase activity was expressed as rate/ mass i.e.

RFU hr-1/mg. The assay was carried out using a black 96-well plate with triplicate

measurements and the temperature of the assay was maintained within the range of 34 oC

– 37oC.

5.3.2.6 Statistics

Data are expressed as the mean value + SEM and statistical analysis was carried out using

SPSS 16.0. Statistical significance was determined using ANOVA test (one-way and two-

way full factorial) and for pair-wise comparisons, post-hoc (Tukey and Duncan) tests

were conducted. A significant difference was observed at the 95% confidence level in

which P values below 0.05 (i.e. p < 0.05) were considered statistically significant and

values above 0.05 (i.e. p > 0.05) were not statistically significant.


5.4.1 IL-1β induced HDF-elastase expression in HDF cells

This section discusses the optimisation for the expression of extracellular and

intracellular HDF-elastase by HDF cells with or without the influence of IL-1β. After

cell culture, confluent HDF cells (> 95%, passage 10) were firstly re-suspended in fresh

DMEM supplemented with 10% heat inactivated FBS (refer as DMEM + 10% FBS) and

seeded at a concentration of 40, 000 cells/ml per well. The expressed HDF-elastase

(extracellular and intracellular) was examined by stimulating treated HDF cells with

various concentrations of IL-1β (25 – 500 nM in DMEM + 10% FBS). For the control

experiment (untreated HDF cells), IL-1β was omitted. It is evident from figure 58 that

IL-1β increased the proliferation of treated HDF cells (figure 58b, d) compared to

untreated HDF cells (figure 58a, c) and this confirmed that IL-1β plays a role in

proliferation of fibroblast cells (Rumalla and Borah 2001; Henry and Garner 2003).

Additionally, it was observed that the growth of both untreated and treated HDF cells

(with or without IL-1β) increased from day 1 (figure 58a-b) to day 7 (figure 58c-d).


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(a) Control HDF cells: Day 1 (b) Treated HDF cells (IL-1β): Day 1

(c) Control HDF cells: Day 7 (d) Treated HDF cells (IL-1β): Day 7

Figure 58. Photomicrographs of HDF cultures (passage 10) showing the effect of IL-1β on the proliferation of HDF cells after 1 day and 7 days growth in DMEM + 10% FBS. Growth of HDF cells after 1 day (a, b) and 7 days (c, d). Control HDF cells in DMEM + 10% FBS without IL-1β (a, c) and increased number of HDF cells in DMEM + 10% FBS supplemented with IL-1β (500 nM; b, d). Scale bar = 100 µm.

Next, the in-vitro cell conditions for HDF cells to express HDF-elastase activity were

optimised. For the positive control experiment, the commercially bought elastase i.e.

PPE was used to demonstrate elastase activity; and the negative controls (DMEM + 10%

FBS, DH2O and 500 nM IL-1β) were used to demonstrate no elastase activity.

In comparison to the negative controls (figure 59a) low amounts of HDF-elastase activity

was produced by the HDF-elastase expressed outside HDF cells (extracellular) as the

initial rates were in the range of 0.26 – 10.46 RFU hr-1 (figure 59b) and on average the

rates for the negative controls were < 9.29 RFU hr-1. Additionally, over the 7 day period

there was no significant difference in the HDF-elastase activity of the extracellular HDF-

elastase expressed by HDF cells depending on the concentration of IL-1β (p > 0.05).


186

(a) (b)

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Figure 59. Rate of HDF-elastase activity expressed by HDF cells in DMEM + 10% FBS. (a): initial rates of elastase activity for both negative (DMEM, DH2O and IL-1β) and the positive (elastase: PPE) controls. HDF cells (passage 10: 40, 000 cells/ml) were stimulated with various concentrations of IL-1β (25 – 500 nM) to examine the rate of HDF-elastase activity expressed extracellular (b) or intracellular (c-d) by HDF cells in DMEM + 10% FBS. Low amounts of HDF-elastase activity were expressed extracellular by both untreated (0 nM IL-1β) and treated (25-500 nM IL-1β) HDF cells (b). In contrast, when HDF cells were lysed, HDF cells were found to express HDF-elastase activity when the surrounding media of HDF cells was water (d: after day 4) rather than DMEM + 10% FBS incubated with/without IL-1β (c). The data for the negative/ positive controls are expressed as the average rate + SE for 7 days with 3 measurements observed per day. The data for the HDF-elastase activity are expressed as the mean rate per day for 3 measurements (n = 3).

In order to determine whether HDF cells expressed elastase activity inside the cells

(intracellular), the cell membrane of HDF cells were lysed releasing the internal contents

into the surrounding media. After HDF cells were stimulated with IL-1β, both untreated

and treated HDF cells were lysed by freeze-thaw and two approaches of the surrounding

media were used in which the HDF cells were incubated.

0

2

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te (

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(a) Unlysed HDF cells: DMEM + 10%FBS (b) Lysed HDF cells: DMEM + 10%FBS

(c) Swollen HDF cells: DH2O (d) Lysed HDF cells: DH2O

Figure 60. Photomicrographs of lysing HDF cells depending on the surrounding media after 5 days growth. The surrounding media was serum containing medium (a, b) and water (c, d). HDF cells prior to lysis (a, c) and after lysis (b, d). The small arrow in photomicrograph b: indicates unlysed HDF cells which has been expanded in the top right side of the photomicrograph. Scale bar = 100 µm.

After the activation process by IL-1β was completed, in the first approach the

surrounding culture medium i.e. DMEM + 10% FBS (supplemented with or without IL-

1β) was not removed and the HDF cells (untreated and treated) were immediately placed

into a freezer (-80oC) to start the lysing of the HDF cells via freeze-thaw. In contrast, in

the second approach the surrounding culture medium DMEM + 10% FBS

(supplemented with or without IL-1β) was removed and subsequently the HDF cells

(untreated and treated) were washed with DPBS, then submerged in water (i.e. DH2O as

the fresh surrounding media) and then the HDF cells were lysed by freeze-thaw.

It is evident from figure 59d that when the HDF cells were lysed with water, after day 4

there was a gradual increase in HDF-elastase activity expressed by HDF cells from days 4

– 7 (as the initial rates increased from 6.37 – 280.23 RFU hr-1); whereas when the

surrounding media i.e. DMEM + 10 % FBS (with and without IL-1ß) was not removed

it is evident from figure 59c that the initial rates of HDF-elastase activity expressed by


188

HDF cells was significantly lower as the initial rates were in the range of 0.27 – 10.06

RFU hr-1 (p < 0.05).

Using the first approach, there are two possible reasons as to why HDF cells did not

express HDF-elastase activity and these are now considered. Firstly, it was possibly that

the HDF cells were not efficiently lysed with DMEM + 10% FBS (figure 60b) in

comparison to water (figure 60d). It was observed that when water was added to HDF

cells they instantly swelled (figure 60c) which is expected because in the presence of

water the ionic strength of the surrounding media was lowered making it easier for water

to penetrate into the HDF cells and thus allowing the HDF cells to efficiently burst

releasing their internal contents into surrounding media via freeze-thaw (figure 60d). As

a result, for the remaining experiments in this chapter, water was used as the surrounding

media to lyse HDF cells via freeze-thaw.

Secondly, for the reason that HDF cells were found to express HDF-elastase activity

when lysed with water, it was envisaged that possibly the serum (10% FBS) in DMEM

may inhibit HDF-elastase activity (Meyer et al. 1975) or the HDF cells were secreting

something resulting in the inhibition of HDF-elastase activity outside the HDF cells.

Accordingly, the experiment was repeated in serum-free medium (SFM): again the HDF

cells were cultured in DMEM + 10% FBS and then near confluent HDF cells (>95%,

passage 12) were re-suspended in SFM and 40, 000 cells/ml were seeded per well. IL-1β

was dissolved at various concentrations (25 – 500 nM) in SFM and subsequently after 4

hours the treated HDF cells were stimulated with IL-1β (25 – 500 nM) whereas SFM was

added to the untreated HDF cells instead of IL-1β. Next, the extracellular and

intracellular expression of HDF-elastase activity by both untreated and treated HDF cells

was monitored over 7 days.

Collectively, over the course of 7 days, in comparison to the negative controls the HDF-

elastase activity of the extracellular or intracellular HDF-elastase expressed by HDF cells

was not significantly different (p > 0.05). Additionally, the HDF-elastase activity of the

extracellular HDF-elastase expressed by HDF cells (untreated and treated) in SFM had

initial rates in the range of 0.83 – 6.69 RFU hr-1 (figure 61b) and these initial rates were

not significantly different from the HDF-elastase activity of the intracellular HDF-

elastase expressed inside HDF cells (untreated and treated) in SFM as the initial rates


189

were in the range of 0.86 – 10.07 RFU hr-1 (figure 61c, p > 0.05). Furthermore, on

average there was no significant difference in the HDF-elastase activity of the

extracellular or intracellular HDF-elastase depending on the concentration of IL-1β in

SFM (p > 0.05).

(a) (b)

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Next the HDF-elastase activity of the extracellular and intracellular HDF-elastase by

both untreated and treated HDF cells in SFM (figure 61b-c) and in DMEM + 10% FBS

(figure 59a, d) were compared. The percentage decrease of HDF-elastase in SFM in

comparison to DMEM + 10% FBS is summarised in table 14: it can be seen that the

HDF-elastase activity of the extracellular or intracellular HDF-elastase expressed by

HDF cells in SFM was significantly lower than in DMEM + 10% FBS (p < 0.05). It was

therefore evident that serum was required for HDF-elastase activity.

Figure 61. Rate of HDF-elastase activity expressed by HDF cells in SFM. (a): the intial rates of elastase activity for both the negative (DMEM, DH2O and IL-1β) and positive (elastase: PPE) controls are expressed as the average rate + SE for 7 days with 3 measurements observed per day. HDF cells (passage 12: 40, 000 cells/ml) were stimulated by IL-1β (25-500 nM) in SFM and the initial rates of HDF-elastase activity expressed extracellularly (b) or intracellularly (c) by both untreated (0 nM IL-1β) and treated (25-500 nM IL-1β) HDF cells. The data for the expression of HDF-elastase activity (b-c) are expressed as the mean rate per day for 3 measurements (n = 3).

0

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190

Table 14. Percentage decrease of HDF-elastase activity in SMF compared to DMEM + 10% FBS for both untreated and treated HDF cells with or without of IL-1β

Average Rate (RFU hr-1) ** HDF-elastase activity

HDF cells *

IL-1 β

SFM DMEM + 10 % FBS

Decrease HDF-elastase activity in SFM (%) #

p-value #

(ANOVA)

Untreated None 3.80 4.07 6.63 p < 0.05

Secreted Treated Added 2.95 4.63 36.37 p < 0.05

Untreated None 2.84 17.13 83.42 p < 0.05

Expressed Treated Added 4.02 58.62 93.14 p < 0.05

* Untreated HDF cells (control: no IL-1β); treated HDF cells (test: added IL-1β i.e. 25 – 500 nM)

** The collective average rate (RFU hr-1) of HDF-elastase activity over 7 days # Comparison between the rate of HDF-elastase activity for SFM versus DMEM + 10% FBS

Consequently, a positive control experiment was then conducted to determine whether

supplementing DMEM with 25% heat inactivated FBS (DMEM + 25% FBS) was

sufficient to increase elastase activity. As the commercially bought elastase i.e. PPE was

used as a positive control for all experiments in this chapter it was therefore used as the

positive model standard to mimic the elastase activity that would normally be expressed

intracellularly or extracellularly by any particular cell such as HDF cells within the

environment of a wound. As we encountered earlier, various researchers have reported

that HDF-elastase is an MMP (Homsy et al. 1988; Beranger et al. 1994; Isnard et al.

2002) whereas PPE is a serine protease and in a previous study Meyer et al. showed

elastase activity of pure PPE to be inhibited by human serum (Meyer et al. 1975). As a

consequence, in order to make direct comparisons between the elastase activity of PPE

and HDF-elastase it was essential to measure the effect of elastase activity for PPE in the

presence of SFM and DMEM + 10% FBS. Therefore, PPE was dissolved in DMEM

(i.e. SFM or supplemented with 10% or 25% FBS) at a concentration of 0.01 mg/ml.

Furthermore, after the PPE was prepared in the above experiments (referred as the

positive control) it was stored at 4oC until it was required for analysis and was

subsequently added to the wells of a 96 well-plate just before the fluorescence substrate

assay was carried out. It was predicted that possibly the storage temperature of PPE

(positive control) may affect the fluorescence output reading. For that reason, it was

essential to measure elastase activity of PPE after it had been stored at 4 oC or incubated

at 37 oC (incubated in a humidified atmosphere containing 5% CO2) to determine the

effect of temperature on the elastase activity of PPE.


191

750

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2250

2750

SFM 10% FBS 25% FBS

Rate

(R

FU

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4 oC 37 oC

Figure 62. Variation of elastase activity in DMEM supplemented with or without FBS against temperature. Elastase (PPE, 0.01 mg/ml) was dissolved in DMEM i.e. serum-free medium (SFM) or supplemented with either 10% or 25% heat inactivated FBS and then stored at 4 oC or incubated at 37 oC for 15 minutes prior to analysis via the fluorescence assay. Initial rates of elastase-substrate reaction versus DMEM (i.e. SFM, +10% FBS and + 25% FBS) demonstrate elastase activity to increase with increasing serum concentration. Elastase activity was significantly high for DMEM + 25% FBS at 37oC compared to SFM and DMEM + 10% FBS (p < 0.05). Data are expressed as the mean + SE of five measurements (n = 5).

Table 15. Statistical differences and similarities between the elastase activity of PPE in DMEM depending on both the concentration of FBS and temperature

Entry DMEM SFM:

4oC

SFM:

37oC

10% FBS:

4oC

10% FBS:

37oC

25% FBS:

4oC

25% FBS:

37oC

1 SFM: 4oC n/a NSD SD SD SD SD

2 SFM: 37oC NSD n/a SD SD SD SD

3 10% FBS: 4oC SD SD n/a NSD NSD NSD

4 10% FBS: 37oC SD SD NSD n/a NSD SD

5 25% FBS: 4oC SD SD NSD NSD n/a NSD

6 25% FBS: 37oC SD SD NSD SD NSD n/a

Abbreviations: SFM (serum-free medium), FBS (fetal bovine serum), n/a (not applicable), NSD (not significantly different, p > 0.05); SD (significantly different, p < 0.05). Note for each row/ column the shaded grey is the same as the un-shaded parts (except for when it states n/a).

Figure 62 shows that the initial rates of elastase activity of PPE significantly depended on

both concentration of serum (FBS) in DMEM and the storage temperature of PPE after

its preparation. Depending on the concentration of serum in DMEM, elastase activity

was found to increase with increasing serum concentration at both 4oC (figure 62: blue

bars) and 37oC (figure 62: maroon bars). High elastase activity of PPE was observed

when PPE was dissolved in DMEM + 25% FBS at 37oC compared to SFM and DMEM

+ 10% FBS. Statistically, elastase activity of PPE was significantly lower for SFM

compared to both 10% and 25% FBS containing DMEM at both 4oC and 37oC (entries

1-2: table 15, p < 0.05). On the contrary, when the initial rates of elastase activity of PPE


192

for both 10% and 25% FBS containing DMEM at both 4oC and 37oC were compared,

elastase activity of PPE was not significantly different as summarised in table 15 (entries

3 – 6); whereas at 37oC elastase activity of PPE for DMEM + 25% FBS was significantly

different from DMEM + 10% FBS (p < 0.05).

(a) (b)

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As elastase activity of PPE was found to increase with increasing serum, it was therefore

predicted that HDF cells required more serum (i.e. 25% FBS) in DMEM to possibly

increase the HDF-elastase activity of the secreted HDF-elastase by HDF cells.

Subsequently, the experiment was repeated in DMEM + 25% FBS: near confluent HDF

cells (>95%, passage 12) were re-suspended in DMEM + 25% FBS and 40, 000 cells/ ml

were seeded per well, IL-1β (25 – 500 nM) was dissolved in DMEM + 25% FBS and

subsequently after 4 hours the treated HDF cells were stimulated with IL-1β (i.e. 25 –

500 nM) whereas DMEM + 25% FBS was added to untreated HDF cells instead of IL-

Figure 63. Rate of HDF-elastase activity expressed by HDF cells in DMEM + 25% FBS. (a): Initial rates of elastase activity for both negative (DMEM, DH2O and IL-1β) and positive (elastase: PPE) controls are expressed as the average rate + SE for 7 days with 3 measurements observed per day. HDF cells (passage 12: 40, 000 cells/ml) were stimulated by IL-1β (25-500 nM) in DMEM + 25% FBS and the initial rates of the HDF-elastase activity expressed extracellularly (b) or intraceullarly (c) by both untreated (0 nM IL-1β) and treated HDF cells (25-500 nM IL-1β) demonstrate that HDF cells are found to intracellularly express elastionolytic activity (c) rather than express HDF-elastase extracellularly. The data for the HDF-elastase activity (b-c) are expressed as the mean rate per day for 3 measurements (n = 3).

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1β. After that the HDF-elastase activity of the expressed extracellular or intracellular

HDF-elastase was again monitored for both untreated and treated HDF cells in DMEM

+ 25% FBS over 7 days (figure 63).

Additionally, in order to keep the temperature consistent for the fluorescence assay, both

the positive (PPE) and negative controls were all equilibrated for 15 minutes at 37oC (in a

humidified atmosphere containing 5% CO2) prior to analysis via the fluorescence

substrate assay.

Although, the serum concentration of DMEM was increased to 25% FBS, collectively

the HDF-elastase activity of the expressed extracellular HDF-elastase by HDF cells was

< 6.00 RFU hr-1 (figure 63b: initial rates ranged from 0.77 – 5.57 RFU hr-1) over the

period of 7 days. As a result, this confirmed that HDF cells were unable to express

extracellular HDF-elastase in the absence or presence of IL-1β (figure 63b). Previously it

was proposed that HDF cells were secreting elastase inhibitors. As mention earlier,

various reports have illustrated that the elastase secreted by fibroblasts is an MMP such

as MMP-2 or MMP-9 (Homsy et al. 1988; Beranger et al. 1994; Isnard et al. 2002).

Additionally, fibroblasts have been reported to secrete MMP-1 (Lobmann et al. 2005;

Lovejoy et al. 1994) and IL-1β has been reported as being linked with the production of

MMP-1 (Herny and Garner 2003; Dasu et al. 2003) in which Dasu et al. have showed

that stimulated dermal fibroblasts primarily produce MMP-1 alongside TIMP-1 under the

influence of IL-1β (Dasu et al. 2003) and Mast et al. have also reported that fibroblasts

secrete TIMPs (Mast and Schultz 1996) which as we have encountered in chapter 2 are

the natural inhibitors of MMPs. This would suggest that if the extracellular HDF-

elastase expressed by HDF cells is an MMP and if the HDF cells are also secreting

TIMPs, then it is most likely that the extracellular HDF-elastase expressed by HDF cells

is inhibited by TIMPs.

Yet again, HDF cells were found to express (intracellular) HDF-elastase activity in

DMEM + 25% FBS (figure 63c) and as expected HDF-elastase activity was expressed

more by the treated HDF cells (i.e. 4.04 – 306.53 RFU hr-1) compared to untreated HDF

cells (i.e. 4.03 – 8.98 RFU hr-1). Although, the HDF-elastase activity of the expressed

intracellular HDF-elastase gradually increased from days 4 – 7 (i.e. initial rates increased

from 25.43 – 306.53 RFU hr-1) statistically the initial rates of HDF-elastase activity for


194

days 5 – 7 i.e. 130.27 – 306.53 RFU hr-1 were significantly higher than days 1 – 4 i.e. 4.04

– 55.81 RFU hr-1 (p < 0.05). On average over the 7 day period, the HDF-elastase activity

for the treated HDF cells was independent on the concentration of IL-1β (p > 0.05).

However, 250 nM of IL-1β gave the highest HDF-elastase activity for the expressed

intracellular HDF-elastase inside HDF cells and was therefore selected for subsequent

experiments.

Next the HDF-elastase activity of the expressed intracellular HDF-elastase in HDF cells

in DMEM + 25% FBS (figure 63b) was compared with DMEM + 10% FBS (figure 59d)

and SFM (figure 61b) only for the treated HDF cells (with 25 – 500 nM IL-1ß) as they

expressed more HDF-elastase activity. Comparisons were made using ANOVA

(oneway) followed by pair comparisons via post hoc Tukey and Duncan tests. As

expected the HDF-elastase activity of the expressed intracellular HDF-elastase in HDF

cells was significantly higher in DMEM + 25% FBS compared to SFM (p < 0.05: figure

64; entry 1: table 16).

Figure 64. Comparisons between the initial rate for HDF-elastase activity in DMEM + 25% FBS versus both SFM and DMEM + 10% FBS. Statistical analysis was carried out using oneway ANOVA followed by pairwise comparison using post hoc (Tukey and Duncan) tests.

However, mixed observations were obtained when HDF-elastase activity in DMEM +

25% FBS was compared with that in DMEM + 10% FBS as summarised in table 16

(entry 2): the Tukey post hoc test indicated that HDF-elastase activity in both 10% and

25% containing DMEM was not significantly different (p > 0.05: figure 64; entry 2: table

16); whereas the Duncan post hoc test indicated that HDF-elastase activity in DMEM +

Table 16. Statistical comparisons of HDF-elastase activity in DMEM (SFM, or supplemented with 10% or 25% FBS) using one-way ANOVA

Entry DMEM 10% FBS 25% FBS

1 SFM SD */# SD */#

2 10% FBS n/a NSD*

SD#

Abbreviations: FBS (fetal bovine serum), SFM (serum-free medium), SD (significantly different, p < 0.05); NSD (not significantly different, p > 0.05), * post hoc: Tukey test and # post hoc: Duncan test.


195

25% FBS was significantly different from that in DMEM + 10% FBS (p < 0.05: figure

64; entry 2: table 16).

Collectively, the initial mean rates of HDF-elastase activity of the expressed intracellular

HDF-elastase in HDF cells in DMEM + 10% FBS (58.62 RFU hr-1) was lower than in

DMEM + 25% FBS (96.97 RFU hr-1) for all IL-1β concentrations over the 7 day period.

As a result, DMEM + 25% FBS was selected for subsequent experiments.

5.4.2 Reduction of HDF-elastase activity by PEGA particles

This section demonstrates the reduction of elastase activity when sample fluids or HDF

cell lysate containing elastase activity are treated with functionalised PEGA particles.

Firstly, a positive control experiment was carried out in which PPE was again used as the

model elastase standard to demonstrate the feasibility of reducing elastase activity in

samples fluids containing PPE at physiological pH and ionic strength. Sample fluids

were prepared by dissolving elastase (PPE, 0.01 mg/ml) in DMEM + 25% FBS (pH 7.4

and 0.15M). After SPPS, functionalised PEGA particles (RAA, EAA, GAA) were

washed with H2O, DH2O, DPBS and then with DMEM + 25% FBS. Functionalised

PEGA particles (10 mg) were transferred to individual centrifuges vials and then swollen

in DMEM + 25% FBS (50 µl) for 5 minutes. Next sample fluids containing PPE were

treated with the prewashed and swollen functionalised PEGA particles (RAA, EAA,

GAA) for 5 and 15 minutes. After that, at each time point the supernatants were

examined with the fluorescence elastase substrate (MeOSuc-Ala-Ala-Pro-Val-AMC) for

the residual elastase activity of PPE in the sample fluids after they had been treated with

PEGA particles using the fluorescence substrate assay.

Figure 65 shows the elastase activity of PPE in sample fluids and the residual elastase

activity of PPE after the sample fluids were treated with PEGA particles for 5 and 15

minutes and the initial rates of elastase for PPE are expressed as RFU hr-1 (figure 65a).

However, as the mass for each PEGA particle varied, the initial rates of residual elastase

activity of PPE in the sample fluids was corrected according to the mass of PEGA

particles used, in which the initial rates were expressed as Rate/ mass (RFU hr-1/mg,

figure 65b).


196

(a) (b)

1500

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2500

0 5 10 15Time (minutes)

Rat

e (

RF

U h

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Elastase RAAEAA GAA

120

140

160

180

200

RAA EAA GAA

Ra

te/

Ma

ss (

RF

U h

r-1/

mg

)

5 minutes 15 minutes

Figure 65. Rates of residual elastase activity remaining in sample fluids (at physiological pH 7.4 and ionic strength 0.15M) after the removal of functionalised PEGA particles (RAA, EAA, GAA). Elastase (PPE: 0.01 mg/ml dissolved in DMEM + 25% FBS) was incubated with functionalised PEGA particles for 5 and 15 minutes. Plots demonstrate (a) the intial rates from elastase-substrate reaction versus the time in which elastase was incubated with PEGA particles, and (b) initial rates of elastase activity per mass of PEGA particles. Data represents the mean + SE of three measurements (n = 3).

Collectively at both 5 minutes and 15 minutes, on average the elastase activity of PPE

(initial rate: 2223.78 RFU hr-1) was significantly reduced more by EAA (negative: 1702.51

RFU hr-1) and RAA (positive: 1881.29 RFU hr-1) PEGA particles (p < 0.05, figure 65a)

compared to GAA (neutral: 2015.35 RFU hr-1) PEGA particles (p > 0.05, figure 65a).

Next, depending on the mass of PEGA particles used, the residual elastase activity of

PPE in samples fluids after they had been treatment with PEGA particles were

compared: elastase activity of PPE was reduced by 20.8 – 26.1% (5 – 15 minutes) after

samples fluids were treated with EAA; whereas when treated with RAA elastase activity

of PPE reduced by 15.0 – 15.8% (5 – 15 minutes) and for GAA it reduced by 11.6 –

7.1% (5 – 15 minutes). Statistically, the elastase activity of PPE was significantly reduced

more by EAA compared to both RAA and GAA (p < 0.05) which was expected since at

pH 7.4 elastase i.e. PPE exists in its cationic form (pI < 8.31) which means that it has

preference of accessing PEGA particles of opposite charge such as EAA (negative)

PEGA particles more compared to both RAA (positive) and GAA (neutral) PEGA

particles. Additionally, as we learnt in chapter 3, after elastase (PPE) cleaves the ECP

from EAA, the negative charge (Glu(–)) is removed causing elastase (PPE) to become

trapped into the matrix of the negative PEGA particles (EAA) and as a result this would

cause a drop in the residual elastase activity of PPE in sample fluids compared to both

the positive (RAA) and neutral (GAA) PEGA particles.


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Depending on the mass of PEGA particles (figure 65b), statistically the residual elastase

activity of PPE in sample fluids was significantly lower for EAA compared to GAA at

both 5 and 15 minutes (p < 0.05) and RAA at 15 minutes (p < 0.05; figure 65b).

However, the residual elastase activity of PPE for the charged PEGA particles (EAA

versus RAA) was not significantly different at 5 minutes (p > 0.05). Similarly the residual

elastase activity of PPE in sample fluids was not significantly different between RAA

versus GAA at both 5 and 15 minutes (p > 0.05). When each PEGA particle (RAA,

EAA and GAA) was compared to time, for each PEGA particle the residual elastase

activity of PPE in all samples fluids was not significantly different at both 5 and 15

minutes (figure 65b). However, the residual elastase activity of PPE was high in sample

fluids that had been treated with the neutral PEGA particles (GAA) and from figure 65a

it was observed that the initial rate of elastase activity of PPE was found to increase at 15

minutes, whereas when taking the mass of GAA particles into account, the residual

elastase activity of PPE were roughly the same for both 5 and 15 minutes (figure 63b),

although it was observed that at 15 minutes the residual elastase activity of PPE was

found to increase by 4.5% in respect of 5 minutes. These observations for GAA

correspond to findings observed in chapter 4 (i.e. fluorescence substrate assay and FITC-

elastase) in which the mopping-up of elastase activity for PPE in sample fluids at pH 7 –

8 and with an ionic strength of 0.1 M did not change over time, there was a slight

increase in elastase activity observed from 5 minutes to 15 minutes (using fluorescence

substrate assay). Similarly, the penetration of FITC-elastase into GAA showed that

fluorescence was found to decrease inside the GAA particles over time. The results in

this chapter and chapter 4 confirm the reason why elastase (PPE) is not retained by the

neutral (GAA) PEGA particles: after elastase i.e. PPE cleaves the neutral ECP it

becomes trapped for the first 5 minutes and subsequently elastase leaks back out of the

GAA particles. This is because with the GAA particles there is no initial charge to be

removed. Instead after elastase (PPE) cleaves the Ala-Ala bond of the neutral ECP

(GA~A) the terminal amine group of the Ala residue becomes protonated (NH3+), hence

causing the swelling of the (+)Ala-PEGA particles to increase (see chapter 3) which

possibly explains why elastase (PPE) is not retained by the neutral GAA particles.

Additionally, it is possible that initial swelling of the neutral GAA particles can only

accommodate and trap some of the elastase (PPE) from sample fluids whereas the

remaining elastase (PPE) remains in the sample fluid.


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Having established that HDF cells express intracellular HDF-elastase, hence HDF-

elastase activity under the influence of IL-1β in DMEM + 25% FBS, the remaining part

of this section highlights the reduction of HDF-elastase activity of the expressed

intracellular HDF-elastase in HDF cells by incubating and treating it with functionalised

PEGA particles (RAA, EAA, and GAA). After HDF cells were lysed, the HDF cell

lysate containing HDF-elastase was collected and thoroughly mixed by drawing the HDF

cell lysate ‘in-and-out’ of a micropipette. Subsequently, the pH of the collected HDF cell

lysate was examined (pH 7.6) and it was then incubated and treated with functionalised

PEGA particles (RAA, EAA, GAA) for 5 and 15 minutes.

After centrifugation, at each time point the supernatant were assayed with the synthetic

fluorescence elastase substrate (MeOSuc-Ala-Ala-Pro-Val-AMC) to examine the residual

HDF-elastase activity in the supernatants of the HDF cell lysate after it had been treated

with each functionalised PEGA particles (RAA, EAA, GAA).

Figure 66 illustrates the mopping-up of HDF-elastase activity by all functionalised PEGA

particles. The elastase activity of both the positive control (PPE) and the negative

controls (DMEM, DH2O and 250 nM IL-1β) are given by figure 66a. Figure 66b shows

the HDF-elastase activity of the expressed HDF-elastase in HDF cells that was added to

PEGA particles and the same trend was previously seen by figure 63b; in figure 66b

HDF-elastase gradually increased from days 4 onwards (> 99.91 – 419.83 RFU hr-1) and

statistically the initial rates of HDF-elastase activity were significantly higher for both

days 6 (419.83 RFU hr-1) and day 7 (296.70 RFU hr-1) compared to days 1 – 5 (p < 0.05).

When HDF cell lysate containing the HDF-elastase activity was treated with PEGA

particles, comparing the initial HDF-elastase activity (figure 66b) with the residual HDF-

elastase activity for all functionalised PEGA particles i.e. RAA (positive: figure 66c) EAA

(negative: figure 66e) and GAA (neutral: figure 66g) the HDF-elastase activity was

significantly reduced by all functionalised PEGA particles at both 5 and 15 minutes (p <

0.05). The initial rates for residual HDF-elastase activity were significantly similar for all

PEGA particles (figure 66c, e, g: p > 0.05).


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Figure 66. Rates of residual HDF-elastase activity remaining in sample fluids after treatment with functionalised PEGA particles (RAA, EAA, GAA) swollen in DMEM + 25% FBS. Initial rates of elastase activity for both the negative and positive controls (a). The HDF-elastase activity of the expressed HDF-elastase in HDF cells (b) was treated with functionalised PEGA particles for 5 and 15 minutes over the course 7 days (c-h). The initial rates of HDF-elastase-substrate reaction are expressed as RFU hr-1 (c, e, g) and then corrected per mass of PEGA particles (d, e, f). PEGA particles: RAA (c, d), EAA (e, f), GAA (g, h). Data represents the mean + SE of three measurements (n = 3).

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However, differences were achieved when the mass of PEGA was taken into account, in

which the initial rate of HDF-elastase-substrate complex were expressed as the rate/mass

i.e. RFU hr-1/mg (figure 66d, f, h). When the initial rates of HDF-elastase activity were

compared with all PEGA particles collectively at both 5 and 15 minutes, on average the

reduction of HDF-elastase activity was significantly similar between the positive and

negative PEGA particles (RAA: figure 66d and EAA: figure 66f, p > 0.05) as well as the

negative and neutral PEGA particles (EAA: figure 66f; and GAA: figure 66h, p > 0.05),

but was significantly different between the positive and neutral PEGA particles (RAA:

figure 66d; and GAA: figure 66h, p < 0.05). RAA was found to reduce HDF-elastase

activity significantly more compared to GAA (figures 66d and 66h respectively, p <

0.05). There are two possible reasons for these observations, firstly as we observed in

chapter 3 at high ionic strength the positive PEGA particles (RAA) were found to swell

significantly more than both the negative (EAA) and neutral (GAA) PEGA particles (p <

0.05) whereas the swelling for EAA and GAA was significantly similar; and for that

reason HDF-elastase was able to penetrate RAA more compared to EAA and GAA.

Secondly, as mentioned previously it is possible the intracellular HDF-elastase expressed

by HDF cells is in fact an MMP such as MMP-1, MMP-2 or MMP-9 (Homsy et al. 1988;

Lobmann et al. 2005; Lovejoy et al. 1994; Herny and Garner 2003; Dasu et al. 2003;

Beranger et al. 1994; Isnard et al. 2002). From table 17 (given in appendix I) the pI

values of these proteases are:- MMP-1: pI 6.47 (entry 4, table 17), MMP-2: pI 5.26 (entry

7, table 17) and MMP-9: pI 5.69 (entry 8, table 17) and since the pH of the HDF cell

lysate containing HDF-elastase that was added to the PEGA particles was pH 7.6, this

therefore means that if HDF-elastase is an MMP then the pI value for each MMP-1,

MMP-2 and MMP-9 indicates that intracellular HDF-elastase expressed by HDF cells

existed in its anionic form and as expected it would access, cleave and become trapped

more in the positive (RAA) PEGA particles and as a result the residual HDF-elastase

activity would be reduce more by RAA compared to both the negative (EAA) and

neutral (GAA) PEGA particles.

5.5 CONCLUSION

The results in this chapter demonstrate that HDF cells were unable to express HDF

elastase extracellularly, but when the cell membrane of HDF cells were lysed in the


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presence of water, the HDF cells were found to express HDF-elastase (intracellularly)

under the influence of IL-1β and the levels of HDF-elastase activity were found to

increase from day 4 onwards. HDF-elastase activity was elevated when the

concentration of serum was increased from 10% to 25% FBS in DMEM and HDF-

elastase activity was found to decrease in SFM.

After sample fluids containing cationic elastase (i.e. PPE) were treated with

functionalised PEGA particles (RAA, EAA and GAA) at physiological pH 7.4 and ionic

strength 0.15M, the residual elastase of PPE (cationic) was significantly reduced more by

EAA (negative) PEGA particles compared to both RAA (positive) and GAA (neutral)

PEGA particles. Similarly, when the supernatants of HDF cell lysate containing HDF-

elastase activity from HDF cells were treated with functionalised PEGA particles

(prewashed and swollen in PBS at pH 7.4, 0.15 M) for 5 and 15 minutes. The residual

HDF-elastase activity in the supernatants of HDF cell lysate was reduced by all

functionalised PEGA particles (RAA, EAA and GAA) over 7 days. Although, it was

found that both the charged particles (RAA and EAA) contained low residual HDF-

elastase activity in the supernatants of HDF cell lysate compared to GAA, the positive

particles were found to significantly reduce HDF-elastase activity expressed by HDF cells

more compared to both the negative (EAA) and neutral (GAA) PEGA particles.

CHAPTER 6: Conclusions & Further Studies

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CCHHAAPPTTEERR 66

CCoonncclluussiioonnss && FFuurrtthheerr SSttuuddiieess


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6.1 CONCLUSIONS

The management of chronic wounds has entered a new era of developing sophisticate

‘responsive’ mechanism-based care products for treating chronic wounds. Abundant

levels of elastase (namely NE) are documented as the main culprit of developing

unbearable chronic wounds as a result of causing the normal wound healing process to

become deadlocked. The aim of this thesis, therefore, was to design an elastase-

responsive chronic wound dressing based on the hydrogel polymer, PEGA in the form

of particles to exclusively and selectively mop-up excess elastase activity in sample fluids

mimicking the environment of chronic wounds. In order to address this, the

experimental study was split into to separate chapters and the findings are summarised

below.

In the first experimental chapter (chapter 3) PEGA(800, 1900) particles were functionalised

with various ECPs using SPPS consisting of the following configurations i.e. Fmoc-X-

Ala-Ala and Fmoc-X-Ala. Comparative studies demonstrated that elastase has a high

preference of hydrolysing Fmoc-X-Ala-Ala-PEGA(800, 1900) particles more whereas

thermolysin had a high preference to hydrolyse both the Fmoc-X-Ala-Ala-PEGA(800, 1900)

particles and Fmoc-X-Ala-PEGA(800, 1900) particles. At low ionic strength, it was

demonstrated that under the influence of pH (6.0 – 9.0) and charge, both thermolysin

and elastase in their cationic and anionic forms selectively penetrated functionalised

PEGA particles and selectively hydrolyse ECPs of opposite charge to that of the

thermolysin or elastase. Thermolysin was studied in its anionic form and was found to

readily hydrolyse the positively charged ECP (RAA or RA) compared to a negatively

charged ECP (EAA or EA). In contrast, elastase was studied both in its cationic and

anionic forms, cationic elastase had a high preference to cleave a negatively charged ECP

(EAA) whereas anionic elastase had the opposite effect. Additionally, the introduction

of ionic charges (positive: Arg(+) and negative: Glu(–)) into PEGA particles were found to

induce an increase in swelling compared to PEGA particles with a neutral charge

(achieved by Gly residue). Once the CMR was removed by a protease from both RAA

and EAA the swelling of each particle was found to reduce at low ionic strength

compared to GAA which contained no CMR. In contrast at high ionic strength the

swelling was found to increase for all ionic and neutral PEGA1900 particles i.e Fmoc-X-

Ala-Ala-PEGA1900 and Fmoc-X-Phe-Phe-PEGA1900. Overall, chapter 3 demonstrated the


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potential of selectively removing excess elastase from sample fluids mimicking the

environment of chronic wounds and this was then investigated further in chapter 4 and

5.

Firstly, in chapter 4 it was confirmed that in its cationic form at pH 7 – 8 elastase had

preference of cleaving the ECP of EAA significantly more than the ECPs of both RAA

and GAA at 0.1 M. The opposite effect was observed; wherein in its anionic form at pH

9 elastase was found to cleave RAA more compared to EAA and GAA at 0.1 M. Next,

the diffusion and accessibility of elastase into Fmoc-X-Ala-Ala-PEGA1900 particles was

explored with the use of various fluorescence techniques. As a consequence of localised

pH effect on the dansyl group, the fluorophore dansyl chloride was found not to be a

suitable staining approach for assessing the diffusion of elastase at various pH into

PEGA particles containing both acidic (e.g. Glu(–)) and basic (e.g. Arg(+)) groups. The

localised pH effect was found to cause base hydrolysis of the dansyl group causing the

fluorescence to increase for the Arg containing particles. Additionally, it is possible that

the dansyl labelling method was slow below pH 8 due to the presence of protonated

molecules causing the fluorescence to decrease for acidic residues.

Comparing the penetration of a 20 kDa FITC-labelled dextran into both un-cleaved and

cleaved PEGA particles, demonstrated the penetration of elastase followed by its

entrapment into charged particles (i.e. RAA particles) compared to neutral particles

(GAA). Removal of the CMR e.g. Arg(+) from the charged PEGA particles by elastase

activated the pores of PEGA particles to become restricted. In doing so, the 20 kDa

FITC-labelled dextran was unable to penetrate into the cleaved PEGA particles. This

collapsing of the pores within the matrix of PEGA1900 particles caused elastase to remain

trapped inside PEGA particles as it has a molecular weight of 25.9 kDa. Interestingly,

the direct penetration and entrapment of elastase into Fmoc-X-Ala-Ala-PEGA1900

particles was achieved by labelling elastase with FITC. At physiological pH 7.4 and ionic

strength 0.15 M, it was demonstrated that FITC-elastase became trapped in both EAA

and RAA particles more compared to GAA particles. In its cationic form, FITC-elastase

was found to become trapped into EAA particles within 2 minutes as FITC-elastase

managed to completely cleave the anionic ECP of the EAA particles within 2 minutes.

In contrast, at 0.1 - 0.15 M it was demonstrated that the neutral particles (GAA) were

unable to retain or encapsulate elastase as the fluorescence of the FITC-elastase inside


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the GAA particles was found to decrease. Coinciding with the observations achieved

from the FITC-dextran and FITC-elastase studies, it was observed that when sample

fluids containing elastase were treated with Fmoc-X-Ala-Ala-PEGA1900 particles, the

residual elastase activity of PPE in the sample fluids was reduced more by both RAA and

EAA compared to GAA particles.

In chapter 5, it was initially demonstrated that after treating sample fluids containing

cationic elastase with functionalised PEGA particles at physiological pH 7.4 and ionic

strength 0.15M, the residual elastase activity of PPE (cationic) was significantly reduced

more by negative PEGA particles compared to both positive and neutral PEGA

particles. Next, after HDF cells were activated by IL-1β and then lysed they were found

to express intracellular HDF-elastase. High HDF-elastase activity was expressed by

HDF cells when DMEM was supplemented with 25% FBS rather than 10% FBS or

SFM. Subsequently, supernatants of HDF cell lysate containing HDF-elastase were

treated with functionalised PEGA particles; over the course of 7 days the residual HDF-

elastase activity was found to decrease more by the ionic particles (RAA and EAA)

compared to GAA. Nevertheless, it was demonstrated that RAA were found to

significantly reduce HDF-elastase activity expressed by HDF cells more compared to

both the EAA and GAA PEGA1900 particles.

Overall, this thesis demonstrates the reduction of elastase activity by mopping it up into

both ionic (positive: RAA and negative: EAA) and neutral (GAA) PEGA particles.

Depending on the charge of elastase (i.e. PPE or HDF-elastase), PEGA particles of

opposite charge (RAA and EAA) were able to reduce elastase more compared to neutral

particles (GAA). Additionally, the charged particles (RAA and EAA) were found to

retain and entrap elastase, unlike the neutral GAA particles which were unable to retain/

entrap elastase.

6.1.1 Limitations of the studies

During the course of this thesis, there were a few limitations associate with the

experimental design of some methods, which are now discussed according to each

chapter.


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In chapter 3, due to time constraints the swelling of the cleaved particles was predicted in

which unmodified PEGA particles were functionalised with either (+)Ala or (+)Phe to

generate (+)Ala-PEGA particles and (+)Phe-PEGA particles, respectively. These particles

represented the cleaved products as schematically summarised in figure 36; and the

swelling of these particles was measured at various pH and ionic strength (figure 37a:

(+)Ala-PEGA; figure 38a: (+)Phe-PEGA). After that the difference in swelling was

calculated between the swelling of the un-cleaved particles (Fmoc-X-Ala-Ala-PEGA:

figure 34; Fmoc-X-Phe-Phe-PEGA: figure 35) and the cleaved particles ((+)Ala-PEGA:

figure 37a; and (+)Phe-PEGA: figure 38a) generating either an increase or decrease in

swelling as demonstrated by figure 37b-d and figure 38b-d, respectively. The method

could be improved by directly measuring the swelling of Fmoc-X-Ala-Ala-PEGA and

Fmoc-X-Phe-Phe-PEGA particles after they have been treated and cleaved with

proteases (e.g. thermolysin and elastase: PPE).

In chapter 4, after the functionalised particles were cleaved with elastase and then stained

with dansyl chloride the ring-effect for the accessibility and diffusion of elastase from

outer surface in the core of PEGA particles was successfully shown by reducing both the

ionic strength and enzyme concentration (figure 47). Seeing as both factors (i.e. ionic

strength and elastase concentration) were studied together, this raises the question as to

which factor influences the accessibility and diffusion of elastase into functionalised

PEGA particles:

• Does reducing ionic strength increase the accessibility of PEGA particles to

enable elastase to diffuse into PEGA particles?

• Does reducing the enzyme concentration slow the reaction to facilitate the

diffusion of elastase into PEGA particles?

To answer these questions, the experiment could be improved by examining each factor

(i.e. reducing the ionic strength or reducing elastase concentration) individually. Then

after dansyl chloride staining, the fluorescence of the TPM images for each factor could

be quantified and compared with one another including the fluorescence observed in

figure 47 in order to determine if one factor or both factors are required to be reduced to

influence the accessibility and diffusion of elastase in functionalised PEGA particles.

In chapter 5, after the HDF cells were lysed, the total cellular protein content should

have been determined using the Biorad protein assay or BCA protein assays in order to


207

accurately determine whether or not the HDF elastase had completely released from the

lysed cells. Additionally, the cell culture conditions of the fluorescence substrate assay

for elastase activity should have been optimised by standardisation of the intracellular

concentration of HDF elastase with the cell number of HDF.

Additionally, during the cell culture studies in chapter 5 it became apparent that the

intracellular HDF elastase expressed by HDF cells maybe an MMP (e.g. MMP-1, MMP-2

or MMP-9: see the ending of section 5.4.2). The identification on whether this

predication was true was not investigated in this thesis. Accordingly, further work could

be done to confirm if the intracellular HDF-elastase was an MMP by reacting the HDF

cell lysate with commercially available MMP inhibitor's as this is a more specific

approach of distinguishing between which MMP it could be. For instance, the

commercial available MMPs inhibitors could be: tetracycline derivatives, hydroxamate/

heterocyclic base MMP inhibitors (as reviewed by Hayashi et al. 2007) and the

bisphosphonate: alendronate (as previously mentioned in chapter 2, see section 2.6.3.3).

6.2 FURTHER STUDIES

Having proven in this thesis that collapsing of pores within the matrix of PEGA particles

entraps elastase within the interior of charged Fmoc-X-Ala-Ala-PEGA particles after the

CMR of the ECP is selectively removed by elastase; clinically it is fundamental to take

other considerations into account before these elastase responsive hydrogel particles can

be applied as a chronic wound dressing to treat chronic wounds. For instance, since

PEGA particles cannot be degraded by the body, the next important factor is to consider

designing a biodegradable dressing or introducing a degradable element/ system into the

polymer PEGA particles dressing which will enable the body to degrade the polymer

PEGA particles to remove them.

Additionally, on a clinical basis before the functionalised PEGA particles can be use to

treat chronic wounds, in-vitro studies should be conducted in which wound fluids

obtained from chronic wounds patients are treated with functionalised PEGA particles.

Consequently, the fluorescence substrate assay as described in chapter 4 could be used to

measure elastase activity observed in chronic wound fluids before and after it has been


208

treated with functionalised PEGA particles. Furthermore, the elastase activity present in

chronic wound fluids can be compared with the elastase activity observed in acute

wound fluids. In relation to acute wounds, the length of time required to decrease the

elastase activity observed in chronic wounds could be determined to normal levels;

bearing in mind that the levels of elastase cannot be totally depleted, as this in turn may

impair the wound healing process.

There are other possibilities of future work in which these particles could be designed for

other applications. In chapter 2, other proteases (such as MMPs, plasmin and thrombin)

were also reported as being elevated in chronic wounds. In order to control the

imbalance levels of these chronic wounds proteases by mopping and entrapping them

into the enzyme responsive PEGA particles as described in this thesis, the P1-P1′ scissile

bond of the ECP would have to be modified so that it mimics the substrate sequence

that is specifically recognised by the target protease(s). A list of substrates/ substrate

sequences recognised by MMPs, plasmin and thrombin including the P1-P1′ scisscle

bond for each substrate sequence are provided in appendix I (table 17). For other

clinical proteases, the same principles would have to be applied in which the P1-P1′

scissile bond would be modified to suit clinically found proteases.

Furthermore, the enzyme responsive PEGA particles could also be designed to have a

dual approach so that the particles have the ability to remove excess proteases as well as

control bacterial infection observed in chronic wounds. This could be achieved by

replacing the terminal Fmoc-group with an anti-inflammatory component that could

release a bioactive group into the wound to combat bacterial infection to facilitate the

healing of chronic wounds.

Additionally, elevated levels of elastase and MMPs have been reported to be associated

with the development of wrinkles (Tsuji et al. 2001; Tsukahara et al. 2001).

Consequently, the enzyme responsive PEGA particles as described within this thesis

could potentially in the same way be used to design cosmetic anti-wrinkle skin creams to

reduce the formation of wrinkles by mopping-up or reducing the levels of elastase or

MMPs by entrapping them into the matrix of these enzyme responsive PEGA particles.

CHAPTER 7: References

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232

AAPPPPEENNDDIIXX II

TTaabbllee 1177

233

Table

17.

Subst

rate

spec

ific

ity

of pro

teoly

tic

pro

tease

s in

volv

ed in the

pro

cess

of wound h

ealing

Pro

tease

s E

nzy

me

Nam

e(s

)/E

C

MW

(kD

a)

Entry

Fam

ily

Typ

e M

MP

Oth

er n

am

es

late

nt

act

ive

Subst

rate

s Subst

rate

spec

ific

ity

pI

1 Ser

ine

E

last

ase

N

/A

N

eutr

op

hil

elas

tase

, PM

N

elas

tase

, HN

E,

LN

E, l

euko

cyte

en

asta

se,

En

asta

se-2

,

(EC

3.4

.21.3

7)

28.6

E

last

in, c

olla

gen

s (I

, II,

II

I, I

V),

cro

sslin

ked

fi

bri

n, f

ibro

nec

tin

, la

min

in, p

rote

ogl

ycan

s,

vitr

on

ecti

n, I

L-2

, IL

-1β,

TN

F-α

, IL

-8, T

NF

r,

CD

4, C

D8,

CD

2, c

lott

ing

fact

ors

, co

mp

lem

ent

fact

ors

, im

mun

ogl

ob

ulin

s

Val

~X

> A

la~

X

X-A

la-A

la-P

ro-V

al~

Y

(X =

MeO

-Suc,

Y =

NH

Ph

NO

2/SB

zl)

X-A

la-A

la-A

la~

Y (

X =

Suc,

Y =

NH

Ph

NO

2)

Val

-Pro

-Val

9.31

2 Ser

ine

Thro

mbin

N

/A

Co

agula

tio

n

fact

or

II,

Fib

rin

oge

nas

e,

pro

thro

mb

in

pre

curs

or,

th

rom

bin

lig

ht/

h

eavy

ch

ain

, α-

Th

rom

bin

,

(EC

3.4

.21.5

)

36

.5

Fib

rin

oge

n, f

ibri

n,

colla

gen

s, c

oag

ula

tio

n

fact

ors

(V

, V

III,

XII

I),

thro

mb

in r

ecep

tors

(P

AR

1, P

AR

3),

hep

arin

co

fact

or

II,

thro

mb

om

od

ulin

, pla

tele

t gl

yco

pro

tein

1b

, p

rote

in

C, a

ctiv

ates

dif

fere

nt

cell

typ

es a

nd

pla

tele

ts

Met

-Pro

-Arg

~Ser

-Ph

e-A

rg

Gly

-Gly

-Val

-Arg

~G

ly-P

ro-A

rg

Ph

e-Ser

-Ala

-Arg

~G

ly-H

is-A

rg

Leu

-Gly

-Ile

-Arg

~Ser

-Ph

e-A

rg

Leu

-Ser

-Pro

-Arg

~T

hr-

Ph

e-H

is

Trp

-Tyr

-Leu

-Arg

~Ser

-Asn

-Asn

Il

e-G

ln-I

le-A

rg~

Ser

-Val

-Ala

Il

e-G

lu-P

ro-A

rg~

Ser

-Ph

e-Ser

G

ln-S

er-P

ro-A

rg~

Ser

-Ph

e-G

ln

Gly

-Val

-Pro

-Arg

~G

ly-V

al-A

sn

Leu

-Asp

-Pro

-Arg

~Ser

-Ph

e-L

eu

Pro

-Ala

-Pro

-Arg

~G

ly-T

yr-P

ro

Ile-

Lys

-Pro

-Arg

~Il

e-V

al-G

ly

Val

-Asp

-Pro

-Arg

~L

eu-I

le-A

sp

Val

-Ser

-Pro

-Arg

~A

la-S

er-A

la

Ile-

Ala

-Gly

-Arg

~Ser

-Leu

-Asn

P

he-

Met

-Pro

-Leu

~Ser

-Th

r-G

ln

Ben

zoyl

-Ph

e-V

al-A

rg-p

NA

,HC

l

3 Ser

ine

Pla

smin

N

/A

Fib

rin

ase,

F

ibri

no

lysi

n,

(EC

3.4

.21.7

)

90

.6

Fib

rin

, lam

inin

, fi

bro

nec

tin

, vit

ron

ecti

n,

coag

ula

tio

n f

acto

rs

(V,

Va,

VII

I, V

IIIa

), p

lasm

in,

colla

gen

s

Lys

~X

> A

rg~

X

Ac-

Lys

-Th

r-X

-Lys

-AM

C (

X =

Tyr

, Ph

e, T

rp,

Ser

) A

c-P

he-

Th

r-T

yr-L

ys-A

MC

A

c-L

eu-T

hr-

Ph

e-L

ys-A

MC

A

c-L

eu-G

lu-P

he-

Lys

-AM

C

7.04

234

Tab

le 1

7. C

on

tinue…

Pro

tease

s E

nzy

me

Nam

e(s

)/E

C

MW

(kD

a)

Entry

Fam

ily

Typ

e M

MP

Oth

er n

am

es

late

nt

act

ive

Subst

rate

s Subst

rate

spec

ific

ity

pI

Pla

smin

(c

on

tin

ue)

T

hr-

Glu

-Tyr

-Arg

L

ys-G

ly-T

ry-A

rg

Glu

-Ala

-Tyr

-Arg

L

ys-T

hr-

Ph

e-L

ys-(

X) 6

-Cys

(X

= G

ly)

Bo

c-G

lu-L

ys-L

ys~

AM

C

4 M

MP

Collagen

ase

s M

MP

-1

Co

llage

nas

e-1,

in

ters

tita

l co

llage

nas

e,

(EC

3.4

.24.7

)

52

43

Co

llage

ns

(I, II

, II

I, V

II,

VII

I, X

, XI)

; ge

lati

n;

aggr

ecan

; ten

asci

n, L

-se

lect

in; I

L-1

β;

pro

teo

glyc

ans;

en

tact

in;

ovo

stat

in; M

MP

-2; M

MP

-9

Ac-

Pro

-Leu

-Gly

-Ser

~L

eu-L

eu-G

ly-O

Et

Mca

-Pro

-Leu

-Gly

~L

eu-D

pa-

Ala

-Arg

-NH

2

Pro

-Met

-Ala

~L

eu-T

rp-A

la-T

hr

Leu

-Pro

-Met

~P

he-

Ser

-Pro

A

c-P

ro-L

eu-A

la-S

~N

va-T

rp-

NH

2 18

3A

rg-T

rp-T

hr-

Asn

-Asn

-Ph

e-A

rg-G

lu-T

ry19

1 P

ro-G

lu-G

ly~

Ile-

Ala

-Gly

P

ro-G

lu-G

ly~

Leu

-L

eu-G

ly

6.47

5 M

MP

Collagen

ase

s M

MP

-8

Co

llage

nas

e-2,

n

eutr

op

hil

colla

gen

ase,

(EC

3.4

.24.3

4)

75

58

Co

llage

ns

(I, II

, II

I, V

, V

II, V

III,

an

d X

); g

elat

in;

enta

ctin

; agg

reca

n;

ten

asci

n,

fib

ron

ecti

n,

Pro

MM

P-1

,2

Gly

-Pro

-Gln

-Gly

~Il

e-T

rp-G

ly-G

ln

Pro

-Leu

-Glu

/A

la~

Tyr

-Trp

-Ser

2,

4-D

np

-Pro

-Gln

-Gly

~Il

e-A

la-G

ly-D

-Arg

-OH

6.38

6 M

MP

Collagen

ase

s M

MP

-13

C

olla

gen

ase-

3,

Rat

in

ters

tita

l co

llage

nas

e,

(EC

3.4

.24.?

)

52

42

Co

llage

ns

(I, II

, II

I, I

V,

IX, X

, XIV

); g

elat

in;

enta

ctin

; agg

reca

n;

ten

asci

n,

pla

smin

oge

n;a

ggre

can

; p

erle

can

; fib

ron

ecti

n;

fib

rin

oge

n/

fib

rin

; o

steo

nec

tin

; MM

P-9

, P

roM

MP

-9,1

3

Mca

-Pro

-Leu

-Gly

~L

eu-D

pa-

Ala

- A

rg-N

H2

Gly

-Pro

-Gln

-Gly

~L

eu-A

la-G

ly-G

ln779

Asp

-Val

-Gly

-Glu

~T

yr-A

sn-V

al-P

he8

8

5.32

235

Tab

le 1

7. C

on

tinue…

Pro

tease

s E

nzy

me

Nam

e(s

)/E

C

MW

(kD

a)

Entry

Fam

ily

Typ

e M

MP

Oth

er n

am

es

late

nt

act

ive

Subst

rate

s Subst

rate

spec

ific

ity

pI

7 M

MP

Gel

atinase

s M

MP

-2

Gel

atin

ase

A,

72-k

Da

gela

tin

ase,

Typ

e IV

co

llage

nas

e,

(EC

3.4

.24.2

4)

71

62

Co

llage

ns

(I, IV

, V

, VII

, X

, XI)

; ge

lati

ns;

fi

bro

nec

tin

; ela

stin

; la

min

in; a

ggre

can

; vi

tro

nec

tin

; dec

ori

n;

IGF

BP

-3/

5; p

ro-M

MP

s (1

, 9 a

nd

13)

Gly

-Pro

-Gln

-Gly

~Il

e-P

he-

Gly

-Gln

M

ca-P

ro-L

eu-G

ly~

Leu

-Trp

-Ala

- A

rg-N

H2

Ac-

Pro

-Leu

-Ala

~SN

va-T

rp-N

H2

Pro

-Gln

-Gly

~Il

e-A

la-G

ly-G

ln

5.26

8 M

MPs

Gel

atinase

s M

MP

-9

Gel

atin

ase

B,

92-k

Da

ge

lati

nas

e, T

ype

V c

olla

gen

ase,

(E

C 3

.4.2

4.3

5)

76

67

Co

llage

ns

(IV

, V, V

II, X

, X

IV, X

VII

); g

elat

in;

enta

ctin

; agg

reca

n; e

last

in;

fib

ron

ecti

n;

fib

rin

oge

n/

fib

rin

; o

steo

nec

tin

; pla

smin

oge

n;

MB

P; I

L-1

β

Gly

-Pro

-Leu

-Gly

~Il

e-A

la-G

ly-G

ln

Pro

-Leu

-Gly

~M

et-L

eu-S

er-H

is

5.69

9 M

MPs

Strom

elys

ins

MM

P-3

Str

om

elys

in-1

, T

ran

sin

, CA

P,

pro

teo

glyc

anas

e,

(EC

3.4

.24.1

7)

52

43

Co

llage

ns

(III

, IV

, V

, an

d

IX);

gel

atin

; agg

reca

n;

per

leca

n; d

eco

rin

; fi

bro

nec

tin

; lam

inin

; el

asti

n; p

rote

ogl

ycan

s;

caes

in;

ost

eon

ecti

n;

ovo

stat

in; e

nta

ctin

; p

lasm

ino

gen

; MB

P;

IL-1

β; M

MP

-2/

TIM

P-2

; p

ro-M

MP

s (1

, 7, 8

, 9, 1

3)

Mca

-Arg

-Arg

-Lys

-Pro

-Val

-Glu

~Z

-Trp

-Arg

-L

ys(d

np

)- N

H2

Dn

p-A

rg-P

ro-L

eu-A

la~

X-T

rp-A

rg-S

er, w

her

e X

equal

s: L

eu, P

he,

Tyr

, Trp

5.77

10

MM

P

Strom

elys

ins

MM

P-

10

Str

om

elys

in-2

, T

ran

sin

-2,

(EC

3.4

.24.2

2)

52

44

Co

llage

ns

(II-

V);

gel

atin

; ca

sein

; agg

reca

n; e

nta

ctin

; el

asti

n;

fib

ron

ecti

n;

vitr

on

ecti

n; l

amin

in,

fib

rin

oge

n/

fib

rin

; M

MP

-1; M

MP

-8

Asp

-Val

-Gly

-His

~P

he-

Ser

-Ser

-Ph

e85

Gly

-Pro

-His

-Leu

~L

eu-V

al-G

lu-A

la29

5.

49

236

Tab

le 1

7. C

on

tinue…

Pro

tease

s E

nzy

me

Nam

e(s

)/E

C

MW

(kD

a)

Entry

Fam

ily

Typ

e M

MP

Oth

er n

am

es

late

nt

act

ive

Subst

rate

s Subst

rate

spec

ific

ity

pI

11

MM

P

Strom

elys

ins

MM

P-

11

Str

om

elys

in-3

, F

uri

n m

oti

f,

(E

C 3

.4.2

4.?

)

51

46

Lam

inin

; fib

ron

ecti

n;

aggr

ecan

;; IG

FB

P-1

; α

1-p

rote

inas

e in

hib

ito

r

Ala

-Ala

-Gly

-Ala

~M

et-P

he-

Leu

-Glu

354

Arg

-Val

-Gly

-Ph

e~T

yr-G

lu-S

er-A

sp68

8 L

ys-A

la-L

eu-H

is~

Val

-Th

r-A

sn-I

le14

4 D

ns-

Pro

-Leu

-Ala

~C

ys(O

meB

zl)-

Trp

-Ala

-Arg

-N

H2

6.25

12

MM

P

Oth

er

MM

Ps

or el

ast

ase

MM

P-

12

Mac

rop

hag

e m

etal

loel

asta

se,

met

allo

elas

tase

, (E

C 3

.4.2

4.6

5)

52

20

Inso

lub

le e

last

in; c

olla

gen

IV

; gel

atin

; cas

ein

; fi

bro

nec

tin

; vit

ron

ecti

n;

lam

inin

; en

tact

in; M

BP

; p

rote

ogl

ycan

s;

fib

rin

oge

n/

fib

rin

; p

lasm

ino

gen

; hep

arin

su

lph

ates

Leu

-Val

-Glu

-Ala

~L

eu-T

yr~

Leu

-Val

-Cys

O3H

-G

ly20

D

np

-Arg

-Pro

-Leu

-Ala

~L

eu-T

rp-A

rg-S

er-N

H2

Arg

-Pro

-Ph

e-G

lu~

Val

-Lys

-Asp

-Th

r203

G

ly-A

la-M

et-P

he~

Leu

-Glu

-Ala

-Ile

356

8.75

En

trie

s 1

, 2

, 4

– 1

2 w

ere

pre

sen

ted

in

pa

rt o

f a

pa

ten

t a

ppl

ica

tion

(Ulij

n,

R.V

., et

al.,

200

7b);

in

the

ab

ove

ta

ble

so

me o

f th

e i

nfo

rma

tion

fo

r th

ese

en

trie

s h

as

been

up

date

d.

Mod

ified

fro

m:

Pu

rich

and

Alli

son

2002

; D

ovi

et

al.

2004

; B

axt

er

199

4;

Pra

ger

et

al.

199

1;

Ed

wa

rds

et

al.

199

9;

Bia

nc

hin

i et

al.

200

2;

Fent

on 1

981

; H

unt

ing

ton

2005

; B

od

e 20

06;

Ko

lev

et

al.

1995

; P

arf

yono

va e

t a

l. 20

02;

Ba

ckes

et

al.

200

0;

Oh

tsu

ka e

t a

l. 20

09;

McC

raw

ley

and

Ma

tris

ian

200

1; S

tern

lich

t an

d W

erb

200

1;

Chu

ng

et

al.

2000

; W

oess

ner

and

Na

ga

se 2

002

; S

yro

vets

and

S

imm

et

200

4;

Xu

e e

t al

. 20

06;

Va

day

and

Lid

er

2000

. T

he i

soele

ctric

po

int

(pI)

of

all

en

zym

es

wa

s ca

lcu

late

d u

sin

g ‘

Co

mpu

te p

I/M

W’

thro

ugh

th

e d

ata

base

pro

vid

ed

by

Sw

issP

rot

(ww

w.e

bi.a

c.u

k/sw

issp

rot/

) fo

r th

e h

um

an

speci

es.

C

lea

vag

e b

etw

een

sci

ssile

P1

–P1

′ b

on

d (~

, i.e

. P

1~P

1′ ),

AM

C (

7-a

min

o-4

-meth

ylco

um

arin

), Dn

p (d

initr

oph

enyl

), D

pa

(N3 -dn

p-L

-2,3

-d

iam

inop

ropi

onic

aci

d).

237

AAPPPPEENNDDIIXX IIII

TTaabbllee 1188

238

Table

18. H

ydro

gel dre

ssin

gs designed to tre

at chro

nic w

ounds

Hydro

gel Pro

duct

Entry

Nam

e

Desc

ription

Manufactu

rers

M

anagem

ent of action

Indication

Contrain

dications

1 A

cryD

erm

Mo

ist

Hyd

rop

hill

ic W

oun

d

Dre

ssin

g

Sh

eet

A

cryM

ed, I

nc.

Sem

iper

mea

ble

en

vir

on

men

t P

rom

ote

mo

ist

wo

un

d h

ealin

g P

rote

ct b

oth

gra

nula

tio

n a

nd

ep

ith

elia

l ti

ssue

• P

ress

ure

ulc

ers

(sta

ges

1-3

)

• P

arti

al –

th

ickn

ess

wo

un

ds

• M

od

erat

e d

rain

age

wo

un

ds

• P

ress

ure

ulc

er (

stag

e 4)

• F

ull-

thic

kn

ess

wo

un

ds

• H

eavy

dra

inin

g w

oun

ds

• T

un

nel

ling

wo

un

ds

2

Am

erig

el O

intm

ent

1o

z T

ub

e

Am

erx

Hea

lth

C

are

Co

rpo

rati

on

T

hic

k p

ale

gel,

liquef

ies

at b

od

y te

mp

erat

ure

.

Co

nta

ins:

oak

ext

ract

, mea

do

wsw

eet

extr

act,

zi

nc

acet

ate,

PE

G 4

00

and

335

0, w

ater

. ↑

pro

tein

s to

wo

un

d s

ite

↓ c

ell m

emb

ran

e p

erm

eab

ility

P

rom

ote

wo

un

d c

on

trac

ture

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• D

iab

etic

ski

n u

lcer

s

• P

ost

-surg

ical

in

cisi

on

s

• B

urn

s:

1st ,

2n

d d

egre

e

• A

llerg

y to

oak

• A

llerg

y to

oak

po

llen

3

Aquaf

lo

Dis

c K

end

all H

ealt

h

Car

e P

rod

uct

s C

om

pan

y

Flu

id m

anag

emen

t M

ildly

– m

od

erat

e ex

ud

ing

wo

un

ds

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• D

erm

al u

lcer

s (s

tage

s 1-

3)

• D

iab

etic

leg

ulc

ers

• B

urn

s:

1st ,

2n

d d

egre

e

• H

eavi

ly d

rain

ing

wo

un

ds

4

AquaG

auze

H

ydro

gel

Imp

regn

ated

Gau

ze

Dre

ssin

g

Pad

s D

eRo

yal

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

Pro

tect

wo

un

d f

rom

deh

ydra

tio

n

No

ad

her

ence

to

wo

un

d b

ed

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• U

lcer

s:

Pre

ssure

(st

ages

2-4

),

der

mal

, leg

, dia

bet

ic

• 3r

d d

egre

e b

urn

s

5

Aquas

ite:

A

mo

rph

ous

Hyd

roge

l

Imp

regn

ated

Gau

ze

hyd

roge

l

Imp

regn

ated

No

n-

Wo

ven

H

ydro

gel

Sh

eet

hyd

roge

l

1

or

3 fl

uid

oz

(cle

ar s

eale

d

hyd

roge

ls)

G

auze

(10

0%

cott

on

sp

on

ge)

G

auze

(n

on

-wo

ven

sp

on

ge)

Sh

eets

(cl

ear)

Dum

ex M

edic

al

All

pro

duct

s:

Fill

wo

un

d s

pac

e M

ain

tain

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

Ab

sorb

sm

all –

mo

der

ate

exud

ates

M

ain

tain

so

oth

ing

and

co

olin

g ef

fect

↓

Pai

n

2nd d

ress

ing

for

secu

rin

g d

ress

ing

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

(d

rain

age

is m

inim

al –

mo

der

ate)

•

3rd d

egre

e b

urn

s

239

Tab

le 1

8. C

on

tin

ue

…

Hydro

gel Pro

duct

Entry

Nam

e

Desc

ription

Manufactu

rers

M

anagem

ent of action

Indication

Contrain

dications

6

Aquas

orb

Hyd

roge

l W

oun

d D

ress

ing

Sh

eet

(tra

nsp

aren

t)

DeR

oya

l P

rim

ary

dre

ssin

g M

ois

ture

vap

our

dif

fusi

on

en

able

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent.

P

rote

ct w

oun

d f

rom

deh

ydra

tio

n

Bac

teri

al b

arri

er

Ab

sorb

ent

(rem

ove

dra

inag

e fr

om

wo

un

d)

Mai

nta

ins

soo

thin

g an

d c

oo

ling

effe

ct

↓ P

ain

• L

eg u

lcer

s

• P

ress

ure

ulc

ers

( 1-

4)

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• B

urn

s:

1st ,

2n

d d

egre

e

• M

od

erat

e –

hea

vy d

rain

age

wo

un

ds

• In

fect

ed, n

on

-in

fect

ed w

oun

ds

• 3r

d d

egre

e b

urn

s

7

Bio

lex

Wo

un

d G

el

Tub

e B

ard

Med

ical

D

ivis

ion

C.R

. B

ard

, In

c.

En

able

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

En

coura

ges

hea

ling

as its

aci

dic

•

Pre

ssure

ulc

ers

(sta

ges

2-4

)

• D

erm

al u

lcer

s

• B

urn

s:

1st ,

2n

d d

egre

e

•

• 3r

d d

egre

e b

urn

s

8

Car

raD

res

Cle

ar

Hyd

roge

l Sh

eet

Sh

eets

(8

9.5%

wat

er p

lus

cro

ss-l

inked

p

oly

eth

elen

e m

atri

x)

Car

rin

gto

n

Lab

ora

tori

es,

Inc.

Ab

sorb

ent

hyd

rop

hili

c d

ress

ings

: ab

sorb

s 3x

w

eigh

t w

ater

, ser

um

, blo

od

R

elea

ses

hea

t to

mai

nta

in c

oo

ling

effe

ct

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• D

erm

al u

lcer

s

• B

urn

s:

1st ,

2n

d d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• In

fect

ed, n

on

-in

fect

ed w

oun

ds

• N

on

e sp

ecif

ied

9

Car

rGau

ze P

ads

and

Str

ips

wit

h

Ace

man

nan

H

ydro

gels

Pad

s Str

ips

Car

rin

gto

n

Lab

ora

tori

es,

Inc.

Pri

mar

y co

ver

or

fille

r A

bso

rb e

xud

ates

M

ain

tain

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• D

erm

al u

lcer

s

• B

urn

s:

1st ,

2n

d d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• In

fect

ed, n

on

-in

fect

ed w

oun

ds

• A

llerg

ic t

o A

cem

ann

an

Hyd

roge

l/ o

ther

co

mp

on

ents

10

Car

rSm

art

Gel

w

oun

d d

ress

ing

wit

h

Ace

man

nan

H

ydro

gels

Tub

e C

arri

ngt

on

L

abo

rato

ries

, In

c.

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

(pro

vid

es m

ois

t to

dry

wo

un

ds

& a

bso

rbs

mo

ist

fro

m w

et w

oun

ds)

C

ove

r an

d p

rote

ct w

oun

ds,

th

eref

ore

↓ p

ain

b

y co

olin

g ef

fect

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• B

urn

s:

1st ,

2n

d d

egre

e

• F

oo

t ulc

ers

• Sta

sis

ulc

ers

• A

llerg

ic t

o A

cem

ann

an

Hyd

roge

l/ o

ther

co

mp

on

ents

240

Tab

le 1

8. C

on

tin

ue

…

Hydro

gel Pro

duct

Entry

Nam

e

Desc

ription

Manufactu

rers

M

anagem

ent of action

Indication

Contrain

dications

11

Car

rasy

n G

el w

oun

d

dre

ssin

g w

ith

A

cem

ann

an

Hyd

roge

ls

C

arra

syn

Sp

ray

Gel

w

oun

d d

ress

ing

wit

h

Ace

man

nan

H

ydro

gels

Car

rasy

n V

wit

h

Ace

man

nan

H

ydro

gels

Tub

e

Bo

ttle

Tub

e

Car

rin

gto

n

Lab

ora

tori

es, I

nc.

A

ll p

rod

uct

s:

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

En

able

s au

toly

tic

deb

rid

emen

t

All

pro

duct

s ar

e use

d t

o t

reat

:

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• V

eno

us

stas

is u

lcer

s

• B

urn

s:

1st ,

2n

d d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• In

fect

ed, n

on

-in

fect

ed w

oun

ds

Car

rasy

n G

el w

oun

d d

ress

ing,

C

arra

syn

V:

• D

iab

etic

ulc

er

• F

oo

t ulc

ers

• P

ost

surg

ical

in

cisi

on

s

• A

llerg

ic t

o:

alo

e ve

ra/

Ace

man

nan

h

ydro

gel

12

Co

mfo

rt-A

id

Sh

eet

wit

h

adh

esiv

e b

ord

er

(glt

ceri

n c

on

ten

t)

So

uth

wes

t T

ech

no

logi

es,

Inc.

En

able

s co

ol an

d s

oo

thin

g ef

fect

P

reve

nts

dry

ing

out

Bac

teri

ost

atic

& f

un

gist

atic

• Surg

ical

in

cisi

on

s

• F

oo

t ulc

ers

• L

eg u

lcer

s

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• B

urn

s:

1st ,

2n

d d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• In

fect

ed, n

on

-in

fect

ed w

oun

ds

• H

igh

ly e

xud

ing

wo

un

ds

• O

ther

ab

sorb

ent

mat

eria

ls

13

Cura

fil G

el w

oun

d

dre

ssin

g &

im

pre

gnat

ed s

trip

s

Tub

e

Imp

regn

ated

pad

Ken

dal

l H

ealt

h

Car

e P

rod

uct

s C

om

pan

y

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

Aid

gra

nula

tio

n, e

pit

hel

ialis

atio

n

En

able

s au

toly

tic

deb

rid

emen

t

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• N

on

e sp

ecif

ied

14

Cura

gel

Dre

ssin

g K

end

all H

ealt

h

Car

e P

rod

uct

s C

om

pan

y

Cle

ar g

el e

nab

les

hyd

rati

on

M

anag

e fl

uid

s •

Der

mal

ulc

ers

(sta

ges

1-3)

• D

iab

etic

leg

ulc

ers

• B

urn

s:

1st ,

2n

d d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• H

igh

ly e

xud

ing

wo

un

ds

241

Tab

le 1

8. C

on

tin

ue

…

Hydro

gel Pro

duct

Entry

Nam

e

Desc

ription

Manufactu

rers

M

anagem

ent of action

Indication

Contrain

dications

15

Cura

sol ge

l w

oun

d

dre

ssin

g T

ub

e P

ad

Hea

lth

po

int

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

Pro

tect

s w

oun

d f

rom

fo

reig

n c

on

tam

inat

ion

•

Surg

ical

in

cisi

on

s

• F

oo

t ulc

ers

• D

iab

etic

ulc

ers

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• Sta

sis

ulc

ers

• B

urn

s:

1st ,

2n

d d

egre

e

• N

on

e sp

ecif

ied

16

Der

maG

el H

ydro

gel

Sh

eet

Sh

eet

Med

line

Ind

ust

ries

, In

c.

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

Ab

sorb

s ex

ud

ates

~ 5

x o

wn

wei

ght

Bac

teri

ost

atic

&

fun

gist

atic

• L

eg u

lcer

s

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• H

igh

ly e

xud

ing

wo

un

ds

17

Der

mag

ram

H

ydro

ph

ilic

Wo

un

d

Dre

ssin

g

Tub

e Im

pre

gnat

ed g

auze

D

erm

a Sci

ence

s,

Inc.

P

rim

ary

cover

or

fille

r A

bso

rb e

xud

ates

(m

ild)

Mai

nta

ins

an a

cid

ic (

mild

) en

viro

nm

ent

• Surg

ical

in

cisi

on

s

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• D

iab

etic

ulc

ers

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• V

eno

us

stas

is u

lcer

s

• B

urn

s:

Par

tial

-th

ickn

ess

• N

on

e sp

ecif

ied

18

Der

mag

ran

Zin

c-Sal

ine

Hyd

roge

l T

ub

e D

erm

a Sci

ence

s,

Inc.

P

rim

ary

cover

or

fille

r M

ain

tain

s m

ois

t fo

r h

ealin

g gr

anula

tio

n t

issu

e

• Surg

ical

in

cisi

on

s

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• V

eno

us

stas

is u

lcer

s

• B

urn

s:

Par

tial

-th

ickn

ess

ther

mal

b

urn

s

• N

on

e sp

ecif

ied

19

Der

maS

yn

D

erm

aGau

ze

Tub

e/Sp

ray

Ste

rile

pad

Der

maR

ite

ind

ust

ries

P

rim

ary

cover

or

fille

r M

ain

tain

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

Fill

ing

dea

d s

pac

e o

f si

nus

trac

ts

Man

age

dee

p w

oun

ds

• P

ress

ure

ulc

ers

• V

eno

us

ulc

ers

• D

iab

etic

ulc

ers

• A

rter

ial ulc

ers

• 3r

d d

egre

e b

urn

s

20

Dia

B G

el w

ith

A

cem

ann

an

Hyd

roge

l

Tub

e C

arri

ngt

on

L

abo

rato

ries

, In

c.

• N

on

-oily

pre

par

atio

n f

or

dia

bet

ic f

oo

t ulc

ers

• D

iab

etic

fo

ot

ulc

ers

• A

llerg

ic t

o:

- A

cem

ann

an H

ydro

gel

• o

ther

co

mp

on

ents

of

pro

duct

242

Tab

le 1

8. C

on

tin

ue

…

Hydro

gel Pro

duct

Entry

Nam

e

Desc

ription

Manufactu

rers

M

anagem

ent of action

Indication

Contrain

dications

21

Ela

sto

-Gel

Ela

sto

-Gel

Plu

s

Sh

eet

(wit

ho

ut

tap

e)

H

ors

esh

oe

shap

e (a

dh

esiv

e b

ord

er)

Sh

eet

(tap

e)

So

uth

wes

t T

ech

no

logi

es,

Inc.

All

pro

duct

s:

Ab

sorb

exu

dat

es

Wo

un

ds

are:

sea

led

, pro

tect

ed, c

ush

ion

ed

Bac

teri

ost

atic

&

fun

gist

atic

• Surg

ical

in

cisi

on

s

• F

oo

t u

lcer

s

• L

eg u

lcer

s

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• H

igh

ly e

xud

ing

wo

un

ds

22

Fle

xiG

el

Gel

sh

eet

dre

ssin

g

(po

lyac

ylam

ide

mat

rix

wit

h

emb

edd

ed

hyd

rop

hili

c p

oly

sacc

har

ide

par

ticl

es)

Sm

ith

&

Nep

hew

, In

c.

Ab

sorb

&

dis

solv

e ex

ud

ates

T

ran

sfer

s m

ois

t vap

our

Pre

ven

ts b

acte

rial

co

nta

min

atio

n b

y o

ffer

ing

a p

hys

ical

sep

arat

ion

bet

wee

n t

he

wo

un

d a

nd

ex

tern

al e

nvi

ron

men

t

• F

ull-

thic

kn

ess

wo

un

ds

• D

iab

etic

fo

ot

ulc

ers

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• V

eno

us

leg

ulc

ers

• A

rter

ial ulc

ers

• Surg

ical

wo

un

ds

• B

urn

s (1

st a

nd

2n

d d

egre

e)

• 3r

d d

egre

e b

urn

s

23

Gen

tell

Hyd

roge

l Im

pre

gnat

ed 2

-ply

ga

uze

pad

s

Sp

ray

gel

T

ub

e

Gen

tell,

Inc.

G

uar

d w

oun

d b

ed

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• V

eno

us

ulc

ers

• B

urn

s: 1

st, 2

nd d

egre

e

• N

on

-dra

inin

g w

oun

ds

• N

on

e sp

ecif

ied

24

Hyp

erge

l T

ub

e M

oln

lyck

e H

ealt

h C

are

Hyp

erto

nic

en

viro

nm

ent

(NaC

l d

isso

lves

an

d

infi

ltra

te n

ecro

tic

tiss

ue)

E

nab

les

auto

lyti

c d

ebri

dem

ent

Dra

ws

out

of

wo

un

d: d

rain

age,

deb

ris,

ed

ema,

b

acte

ria

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

• P

ress

ure

ulc

ers

(sta

ges

3-4

)

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• T

un

nel

ling

wo

un

ds

• D

o n

ot

use

fo

r d

ead

ti

ssue

25

Hyp

erio

n H

ydro

gel

Gau

ze D

ress

ing

Pad

H

yper

ion

M

edic

al, I

nc.

M

ain

tain

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

(full-

thic

kn

ess

wo

un

ds)

If

by

mis

take

th

e p

rod

uct

co

nta

cts

inta

ct s

kin

th

en g

lyce

rin

e p

reve

nts

mac

erat

ion

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• Sta

sis

ulc

ers

• B

urn

s: 1

st, 2

nd d

egre

e

• M

od

erat

e –

hea

vy

exud

ates

243

Tab

le 1

8. C

on

tin

ue

…

Hydro

gel Pro

duct

Entry

Nam

e

Desc

ription

Manufactu

rers

M

anagem

ent of action

Indication

Contrain

dications

26

Hyp

erio

n

Hyd

rop

hill

ic

Wo

un

d G

el

H

yper

ion

H

ydro

ph

ilic

Wo

un

d

Dre

ssin

g

Tub

e

Imp

regn

ated

gau

ze

Hyp

erio

n

Med

ical

, In

c.

Bo

th p

rod

uct

s:

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

by

hyd

rate

wo

un

d b

ed

En

able

osm

oti

c gr

adie

nt

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• Sta

sis

ulc

ers

• B

urn

s: 1

st, 2

nd d

egre

e

• M

od

erat

e –

hea

vy

exud

ates

27

Lam

in H

ydra

tin

g G

el

Tub

e B

ard

Med

ical

D

ivis

ion

C

.R.B

ard

, In

c.

Mai

nta

ins

mo

ist

envi

ron

men

t fo

r w

oun

ds/

burn

s •

Pre

ssure

ulc

ers

(sta

ges

1-4

)

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• D

iab

etic

ulc

ers

• Sta

sis

ulc

ers

• B

urn

s: 1

st, 2

nd d

egre

e

• A

rter

ial ulc

ers

• P

ost

op

erat

ive

inci

sio

ns

• 3r

d d

egre

e b

urn

s

28

Intr

aSit

e G

el

Ap

plip

aks

(w

ater

, ca

rbo

xym

eth

ylce

llose

p

oly

mer

, so

diu

m a

nd

p

rop

ylen

e gl

yco

l)

Sm

ith

&

Nep

hew

, In

c.

Hyd

rate

s: g

ran

ula

tio

n t

issu

e R

ehyd

rate

s: d

ry e

sch

ar, s

lough

D

isso

lve

nec

roti

c ti

ssue

to liq

uid

A

id a

uto

lysi

s M

ain

tain

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• T

un

nel

ling

wo

un

ds

• 3r

d d

egre

e b

urn

s

29

MP

M E

xcel

Gel

MP

M H

ydro

gel

Tub

e

Tub

e

MP

M M

edic

al,

Inc.

M

ain

tain

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

E

nab

les

auto

lyti

c d

ebri

dem

ent

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• V

eno

us

stas

is u

lcer

s

• B

urn

s: 1

st, 2

nd d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• D

o n

ot

use

fo

r d

rain

ing

wo

un

ds

30

MP

M G

elP

ad

Hyd

roge

l Sat

ura

ted

D

ress

ing

Ste

rile

Pad

M

PM

Med

ical

, In

c.

Pri

mar

y co

ver

M

ain

tain

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

E

nab

les

auto

lyti

c d

ebri

dem

ent

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• B

urn

s: 1

st, 2

nd d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• D

o n

ot

use

fo

r d

rain

ing

wo

un

ds

31

MP

M R

egen

ecar

e T

ub

e M

PM

Med

ical

, In

c.

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

E

nab

les

auto

lyti

c d

ebri

dem

ent

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• V

eno

us

stas

is u

lcer

s

• B

urn

s: 1

st, 2

nd d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• N

on

e sp

ecif

ied

244

Tab

le 1

8. C

on

tin

ue

…

Hydro

gel Pro

duct

Entry

Nam

e

Desc

ription

Manufactu

rers

M

anagem

ent of action

Indication

Contrain

dications

32

No

rmlg

el

Ste

rile

tub

e M

őln

lyck

e H

ealt

h C

are

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• In

fect

ed w

oun

ds

33

NU

-GE

L

Tub

e

(co

nta

ins

wat

er,

carb

oxy

met

hyl

cello

se

po

lym

er, so

diu

m

algi

nat

e an

d p

rop

ylen

e gl

yco

l)

Joh

nso

n &

Jo

hn

son

, In

c.

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

E

nab

les

auto

lyti

c d

ebri

dem

ent

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• V

eno

us

stas

is u

lcer

s

• B

urn

s: 1

st, 2

nd d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• 3r

d d

egre

e b

urn

s

34

Pan

oP

lex

Hyd

roge

l W

oun

d D

ress

ing

P

ano

Gau

ze N

on

W

ove

n H

ydro

gel

Dre

ssin

g

Syr

inge

/T

ub

e

Imp

regn

ated

Gau

ze P

ad

Sag

e P

har

mac

euti

cals

B

oth

pro

duct

s:

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

C

ove

r w

oun

d t

her

efo

re p

reven

t fo

reig

n

mat

eria

l en

teri

ng

wo

un

d a

rea

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• D

erm

al u

lcer

s

• B

urn

s: 1

st, 2

nd d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• P

ost

-op

erat

ive

inci

sio

ns

• N

on

e sp

ecif

ied

35

Ph

yto

Der

ma

Wo

un

d G

el

Squee

ze B

ott

le

Alo

e L

ife

Inte

rnat

ion

al

Aid

hea

ling

by

mo

ist

and

aci

dic

en

viro

nm

ents

•

Pre

ssure

ulc

ers

(sta

ges

1-4

)

• N

on

e sp

ecif

ied

36

Puri

lon

Gel

A

cco

rdio

n P

ack

(co

nta

ins

wat

er,

carb

oxy

met

hyl

cellu

lose

an

d c

alci

um

alg

inat

e)

Co

lop

last

C

orp

ora

tio

n

En

able

s au

toly

tic

deb

rid

emen

t o

f n

ecro

tic

tiss

ue

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• L

eg u

lcer

s

• 3r

d d

egre

e b

urn

s

37

Res

tore

Hyd

roge

l D

ress

ing

Am

orp

ho

us

tub

e/

imp

rega

nat

ed g

auze

sp

on

ge/

im

pre

gnat

ed g

auze

p

acki

ng

stri

p

Ho

llist

er

Inco

rpo

rate

d

Mai

nta

ins

mo

ist

wo

un

d h

ealin

g en

viro

nm

ent

F

illin

g d

ead

sp

ace

of

sin

us

trac

ts

Man

age

dee

p w

oun

ds

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• A

rter

ial st

asis

ulc

ers

• B

urn

s: 1

st, 2

nd d

egre

e

• N

on

e sp

ecif

ied

38

SA

F-G

el H

ydra

tin

g D

erm

al W

oun

d

Dre

ssin

g

Tub

e (a

lgin

ate-

con

tain

ing)

C

on

vaT

ec

Mai

nta

ins

max

imum

mo

ist

envi

ron

men

t

•

Pre

ssure

ulc

ers

(sta

ges

1-4

)

• Sta

sis

ulc

ers

• B

urn

s: 1

st, 2

nd d

egre

e

• N

on

e sp

ecif

ied

245

Tab

le 1

8. C

on

tin

ue

…

Hydro

gel Pro

duct

Entry

Nam

e

Desc

ription

Manufactu

rers

M

anagem

ent of action

Indication

Contrain

dications

39

Ski

nte

grit

y A

mo

rph

ous

Hyd

roge

l

Ski

nte

grit

y H

ydro

gel

Imp

regn

ated

Gau

ze

Bel

low

s b

ott

le/

T

ub

e/

Imp

regn

ated

gau

ze

Med

line

Ind

ust

ries

, In

c.

Mai

nta

ins

mo

ist

envi

ron

men

t

•

Pre

ssure

ulc

ers

(sta

ges

2-4

)

• V

eno

us

stas

is u

lcer

s

• B

urn

s: 1

st, 2

nd d

egre

e

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• P

ost

-op

erat

ive

inci

sio

ns

• H

yper

sen

siti

ve

to

pro

duct

40

So

loSit

e W

oun

d G

el

So

loSit

e G

el

Co

nfo

rmab

le

Wo

un

d D

ress

ing

Tub

e/

Push

-butt

on

ap

plic

ato

rs

G

el p

ad

Sm

ith

&

Nep

hew

, In

c.

.

Hyd

rate

s: g

ran

ula

tio

n t

issu

e R

ehyd

rate

s: d

ry e

sch

ar, s

lough

A

dd

itio

n o

f p

rese

rvat

ives

in

hib

it b

acte

rial

gr

ow

th

Ab

sorb

exc

ess

exud

ate

En

able

s au

toly

tic

deb

rid

emen

t M

ain

tain

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

chro

nic

wo

un

ds:

• P

ress

ure

ulc

ers

• V

eno

us

leg

ulc

ers

• 3r

d d

egre

e b

urn

s

41

Ten

der

Wet

Gel

Pad

P

ad

Med

line

Ind

ust

ries

, In

c.

Mult

i-la

yer

gel,

wit

h a

co

re a

bso

rben

t.

Hyd

rop

ho

bic

co

veri

ng

laye

r al

low

s w

oun

d

exud

ates

to

pen

etra

te g

el w

ith

out

adh

erin

g to

th

e w

oun

d.

Mai

nta

ins

mo

ist

wo

un

d e

nvi

ron

men

t d

ue

to

the

Ten

der

Wet

(R

inge

r’s)

so

luti

on

.

En

able

s au

toly

tic

deb

rid

emen

t (e

rad

icat

e n

ecro

tic

tiss

ue

and

deb

ris)

• P

arti

al-,

full-

thic

kn

ess

wo

un

ds

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• V

eno

us

ulc

ers

• A

rter

ial ulc

ers

• D

iab

etic

ulc

ers

• B

urn

s: 1

st, 2

nd d

egre

e

• 3r

d d

egre

e b

urn

s

42

3M T

egag

el

Hyd

roge

l W

oun

d

Fill

ers

Pad

/T

ub

e 3M

Hea

lth

Car

e E

nab

les

auto

lyti

c d

ebri

dem

ent

(hyd

rati

ng

dea

d t

issu

e)

Fill

s d

ead

sp

ace

of

thic

knes

s w

oun

ds

• P

arti

al-,

th

ickn

ess

der

mal

w

oun

ds:

• P

ress

ure

ulc

ers

• V

eno

us

ulc

ers

(in

suff

icie

ncy

)

• A

rter

ial ulc

ers

• D

iab

etic

ulc

ers

• B

urn

s: 1

st, 2

nd d

egre

e

• P

ost

-surg

ical

in

cisi

on

s

• 3r

d d

egre

e b

urn

s

• A

llerg

ic t

o p

rod

uct

in

gred

ien

t

246

Tab

le 1

8. C

on

tin

ue

…

Hydro

gel Pro

duct

Entry

Nam

e

Desc

ription

Manufactu

rers

M

anagem

ent of action

Indication

Contrain

dications

43

Ult

rex

Gel

Wo

un

d

Dre

ssin

g w

ith

A

cem

ann

an

Hyd

roge

l

Tub

e (c

on

tain

s A

cem

ann

an

Hyd

roge

l)

Car

rin

gto

n

Lab

ora

tori

es, I

nc.

M

ain

tain

s m

ois

t w

oun

d h

ealin

g en

viro

nm

ent

• P

ress

ure

ulc

ers

(sta

ges

1-4

)

• Sta

sis

ulc

ers

• F

oo

t ulc

ers

• B

urn

s: 1

st, 2

nd d

egre

e

• P

ost

-surg

ical

in

cisi

on

s

• A

llerg

ic t

o:

- A

cem

ann

an H

ydro

gel

- o

ther

co

mp

on

ents

of

pro

duct

44

WO

UN

’DR

ES

Co

llage

n H

ydro

gel

Tub

e C

olo

pla

st

Co

rpo

rati

on

M

ain

tain

s m

ois

t w

oun

d e

nvi

ron

men

t G

ives

pro

tect

ion

to

gra

nula

tio

n t

issu

e E

nab

les

auto

lyti

c d

ebri

dem

ent

• P

ress

ure

ulc

ers

(sta

ges

2-4

)

• V

eno

us

stas

is u

lcer

s

• P

arti

al-,

th

ickn

ess

wo

un

ds

• B

urn

s: 1

st, 2

nd d

egre

e

• 3r

d d

egre

e b

urn

s

45

Wo

un

d D

ress

ing

wit

h C

lear

Sit

e

Ban

dag

e R

oll

wit

h

Cle

arSit

e

Bo

rder

ed

dre

ssin

g/

Bo

rder

less

d

ress

ing/

Is

lan

d d

ress

ing

B

and

age

roll

CO

NM

ED

C

orp

ora

tio

n

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Ring-effect:

From the cross-sectional images obtained from TPM, the diffusion of an enzyme into

PEGA particles can be monitored from the outer surface into the central-core of PEGA

particles by observing a ‘ring-effect’ as schematically illustrated by figure 67.

Figure 67. Diffusion of an enzyme into PEGA particles via a ‘ring-effect’. When unmodified PEGA particles are stained by dansyl chloride they fluoresce blue (1), coupling of an ECP (2) generates functionalised PEGA particles (3) which are viewed as a black image via TPM (3). After the functionalised particles are treated with an enzyme at various time points (4) the cleaving of the ECP exposes free NH2 groups (blue) over the course of time, generating a ring-effect (5) which thickens inwards as the enzyme diffuses into the particles and cleaves the ECPs from within the PEGA particles until the entire ECP is cleaved from the inner-central core of PEGA particles.

(1)

(2)

(3)

(4)

(5)

249

Figure 68. Separation of elastase by SDS-page via 1-D electrophoresis. Lane 1: standards of known molecular weight. Lane 2: separation of elastase (PPE) gave 2 bands with a molecular weight of 31.35 kDa (top band: inactive PPE) and 25.95 kDa (bottom band: active PPE).

250 kDa

150 kDa

100 kDa

75 kDa

50 kDa

37 kDa

25 kDa

20 kDa

15 kDa

10 kDa

PPE (inactive)

PPE (active)

Lane 1 Lane 2

Elastase Responsive Hydrogel Dressing for Chronic Wounds

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