pre clinical evaluation of amphiphilic silicone...

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PRE‐CLINICAL EVALUATION OF AMPHIPHILIC SILICONE

OLIGOMERS FOR SCAR REMEDIATION

By

Emily C Lynam

Bachelor of Applied Science (Hons), QUT

Cell and Molecular Biosciences Discipline

Faculty and Science and Technology

Queensland University of technology

Brisbane, Australia

A thesis submitted for the degree of Doctor of Philosophy of the

Queensland University of Technology

2011

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KEY WORDS

Wound Healing

Hypertrophic Scar

Silicone

Fibroblasts

Keratinocytes

Apoptosis

Gene Expression

STATEMENT OF ORIGINALITY

The work contained in this thesis has not been previously submitted for a degree or

diploma at any other higher education institution. To the best of this authors knowledge

this thesis contains no material which has been previously published or written by any

other person except where due references are made.

SIGNED:

DATE:

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ACKNOWLEDGEMENTS

Firstly, I would like to thank my wonderful husband, Paul. There have been many

challenging times over the past few years, but you have always supported me and put my

studies first. You are my rock and I simply would not have made it through without you.

I would also like to thank Prof. Zee Upton and Prof. Graeme George for being my

supervisors and for providing endless guidance throughout my PhD. Zee has always been a

great source of help and wisdom for both experimental and personal needs. I have always

respected Zee’s drive and passion for students and to help others reach their own goals.

This has been evident in many ways at different times through the last three years.

Graeme, your input in our fortnightly silicone group meetings was invaluable, especially

since you have always left me with questions to think about thereafter. I always seem to

understand subjects relating to chemistry better after talking to Graeme. The amount of

respect I have for both Zee and Graeme is immense and it has been a pleasure to be your

student.

To the silicone group: thank you Tim Dargaville, Marilla Dickfos, Babak Radi, Daniel Keddie

and Brooke Farrugia. It has been a pleasure to meet with you on a regular basis and to

learn from you. Thank you for your patience in the small amount of chemistry knowledge

that I had at the beginning. Thank you also for your diligence in learning and understanding

the biology side of things.

A lot of thanks also need to go to Jacqui McGovern, Helen McCosker and Matt Hadaway.

We have been through an undergraduate degree, an honours year and now a PhD

together. While we really only started to get to know each other towards the end of our

honours year by eating lunch together every day, it has been great getting to know you.

What started small has now grown and I would therefore also like to thank Dan Broszczak,

Amanda Marks, Jess Heinemann and James Broadbent, our current ‘lunch crew’. You all

mean the world to me and I have a great deal of respect for you. It has been a pleasure

getting to know you and to share all of our IHBI experiences together. I hope our friendship

lasts beyond our time together at IHBI.

To all the post‐docs and other PhD students belonging to the TRR team, there are just too

many to thank but you know who you are. The TRR team is an interactive group and there

has always been someone to help out and answer any of my questions when required.

Furthermore, the administration support that has been provided by Abrona Bugler and

Amanda Marks, as well as more recently by Shally Ho and Nicky Gillott, has been

exceptional. I know that you have all helped me with various administration issues and I

genuinely thank you for that.

My scholarship has been funded by an Australian Postgraduate Award with a QUT Vice‐

Chancellor’s top‐up fund. My PhD would have been very difficult to complete without this

financial support. The project I undertook was funded by an ARC Discovery Grant, DP

0877988 and the research performed herein would not have been possible without this.

Thank you also to the Institute of Health and Biomedical Innovation for providing me space

in which I have performed my research. I am also particularly grateful for Zee’s financial

support in allowing me to attend a number of conferences, both in Australia and overseas.

Lastly, I would also like to thank the rest of my family, Mum, Dad, Lou, Rob, Pam, Dan,

Sarah, Marty and Thea as well as their respective partners, who have provided me with

endless support and encouragement over the last couple of years. While at times it has

been difficult, I thank you for always being there for me, even when you didn’t understand

why I was stressed out or going crazy. Together with Paul, it has been an absolute privilege

to share this journey with you.

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ABSTRACT

The formation of hypertrophic scars is a frequent outcome of wound repair and often

requires further therapy with treatments such as silicone gel sheets (SGS; Perkins et al.,

1983). Although widely used, knowledge regarding SGS and their mechanism of action on

hypertrophic scars is limited. Furthermore, SGS require consistent application for at least

twelve hours a day for up to twelve consecutive months, beginning as soon as wound

reepithelialisation has occurred. Preliminary research at QUT has shown that some species

of silicone present in SGS have the ability to permeate into collagen gel skin mimetics upon

exposure. An analogue of these species, GP226, was found to decrease both collagen

synthesis and the total amount of collagen present following exposure to cultures of cells

derived from hypertrophic scars. This silicone of interest was a crude mixture of silicone

species, which resolved into five fractions of different molecular weight. These five

fractions were found to have differing effects on collagen synthesis and cell viability

following exposure to fibroblasts derived from hypertrophic scars (HSF), keloid scars (KF)

and normal skin (nHSF and nKF). The research performed herein continues to further assess

the potential of GP226 and its fractions for scar remediation by determining in more detail

its effects on HSF, KF, nHSF, nKF and human keratinocytes (HK) in terms of cell viability and

proliferation at various time points. Through these studies it was revealed that Fraction IV

was the most active fraction as it induced a reduction in cell viability and proliferation most

similar to that observed with GP226. Cells undergoing apoptosis were also detected in HSF

cultures exposed to GP226 and Fraction IV using the Tunel assay (Roche). These

investigations were difficult to pursue further as the fractionation process used for GP226

was labour‐intensive and time inefficient. Therefore a number of silicones with similar

structure to Fraction IV were synthesised and screened for their effect following application

to HSF and nHSF. PDMS7‐g‐PEG7, a silicone‐PEG copolymer of low molecular weight and low

hydrophilic‐lipophilic balance factor, was found to be the most effective at reducing cell

proliferation and inducing apoptosis in cultures of HSF, nHSF and HK. Further studies

investigated gene expression through microarray and superarray techniques and

demonstrated that many genes are differentially expressed in HSF following treatment with

GP226, Fraction IV and PDMS7‐g‐PEG7. In brief, it was demonstrated that genes for TGFβ1

and TNF are not differentially regulated while genes for AIFM2, IL8, NSMAF, SMAD7, TRAF3

and IGF2R show increased expression (>1.8 fold change) following treatment with PDMS7‐

g‐PEG7. In addition, genes for αSMA, TRAF2, COL1A1 and COL3A1 have decreased

expression (>‐1.8 fold change) following treatment with GP226, Fraction IV and PDMS7‐g‐

PEG7. The data obtained suggest that many different pathways related to apoptosis and

collagen synthesis are affected in HSF following exposure to PDMS7‐g‐PEG7. The

significance is that silicone‐PEG copolymers, such as GP226, Fraction IV and PDMS7‐g‐PEG7,

could potentially be a non‐invasive substitute to apoptosis‐inducing chemical agents that

are currently used as scar treatments. It is anticipated that these findings will ultimately

contribute to the development of a novel scar therapy with faster action and improved

outcomes for patients suffering from hypertrophic scars.

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TABLE OF CONTENTS

Key Words iii

Statement of Originality iv

Acknowledgements v

Abstract vii

Table of Contents ix

List of Figures xiv

List of Tables xvi

List of Abbreviations xvii

List of Publications and Presentations xx

CHAPTER 1.0 LITERATURE REVIEW 1

1.1 Skin Homeostasis 1

1.2 Abnormal Scars 1

1.3 Pathophysiology of Abnormal Scars 3

1.3.1 Dermal Fibroblast Heterogeneity 5

1.3.2 Role of Epidermis in Wound Healing 6

1.3.3 Scarring and Apoptosis 7

1.4 Management of Abnormal Scars 8

1.5 Silicone Gel Treatments 9

1.5.1 Mechanism of Silicone Gel Sheet Action 10

1.5.2 The Role of Silicone in Scar Prevention 12

1.5.3 Adverse Effects of Silicone Treatment 13

1.5.4 Type of Silicone Gel Treatment 13

1.5.5 Comparing other Therapies to Silicone 14

1.5.6 Limitations of Research 15

1.6 Returning to the Beginning 17

1.6.1 Research undertaken at QUT 17

1.6.2 Silicone and Apoptosis? 19

1.6.3 In Vitro to In Vivo 20

1.7 Project Hypothesis and Aims 21

1.7.1 Hypothesis 21

1.7.2 Aims 21

CHAPTER 2.0 MATERIALS AND METHODS 23

2.1 Introduction 23

2.2 Silicone Preparation 23

2.2.1 Fractionation of GP226 23

2.2.2 Synthesis of PDMS‐PEG oligomers 24

2.3 Cell Culture 25

2.3.1 Fibroblast Cell Culture 25

2.3.2 Keratinocyte Cell Culture 25

2.4 Functional Assays 26

2.4.1 WST‐1 Assay for Determination of Cell Viability 26

2.4.2 Cyquant Assay for Determination of Cell Proliferation 27

2.4.3 Analysis of Silicone and Protein Interaction 28

2.4.4 Analysis of Cell Morphology via Real‐Time Microscopy 29

2.4.5 Tunel Assay for the Detection of Apoptosis 30

2.5 Gene Analysis 31

2.5.1 Extraction of RNA from Silicone‐Treated Fibroblasts 31

2.5.2 HumanHT‐12 v3 Expression BeadChip Microarray 31

2.5.3 Microarray Data Analysis using GeneSpring GX 10.0 32

2.5.4 Gene Ontology and Canonical Pathway Analysis using Ingenuity

Pathway Analysis Tools 33

2.5.5 Apoptosis Super Arrays 33

2.6 Confirmation of Differential Gene Expression using Quantitative RT‐PCR 34

2.6.1 Standard PCR Conditions 34

2.6.2 Primer Design 35

2.6.3 Reverse Transcription (RT) for qRT‐PCR 37

2.6.4 PCR and Amplicon purification 37

2.6.5 qRT‐PCR 37

2.7 Statistical Analysis 38

CHAPTER 3.0 FUNCTIONAL ANALYSIS OF THE EFFECTS OF GP226 AND ITS

FRACTIONS ON DERMAL FIBROBLASTS

39

3.1 Introduction 39

3.2 Experimental Procedures 40



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3.2.5 WST‐1 Assay for Determination of Cell Viability 41


3.2.7 Analysis of Silicone and Protein Interaction 41



3.2.10 Statistical Analysis 42

3.3 Results 43

3.3.1 Analysis of Cell Viability and Proliferation following Treatment

with Silicone 43

3.3.2 Silicone and Protein Interaction 53

3.3.3 Further Investigation of HSF Morphology following Treatment

with Fraction IV 57

3.3.4 Investigating the Induction of HSF Apoptosis following

Treatment with Silicone. 57

3.4 Discussion 60

CHAPTER 4.0 FUNCTIONAL ANALYSIS OF THE EFFECTS OF SYNTHETIC

SILICONES, INCLUDING PDMS7‐g‐PEG7, ON DERMAL FIBROBLASTS

65

4.1 Introduction 65









4.3 Results 69

4.3.1 Analysis of Cell Proliferation following Treatment with the

Synthesised Silicones

69

4.3.2 Further Analysis of Cell Proliferation following Treatment with

PDMS7‐PEG7

71

4.3.3 Investigation of Cell Morphology following Treatment with

PDMS7‐PEG7

77

4.3.4 Investigation of the Induction of HSF Apoptosis following

treatment

82

4.4 Discussion 87

CHAPTER 5.0 INVESTIGATIONS INTO THE EFFECT OF AMPHIPHILIC SILICONE‐

PEG COPOLYMERS AT THE GENOMIC LEVEL

91

5.1 Introduction 91





5.2.4 RNA Extraction 92

5.2.5 Microarray 93

5.2.6 Gene Ontology, Canonical Pathway and Functional Network

Analysis 93

5.2.7 Super Arrays 93

5.2.8 Confirmation of Differential Gene Expression using Quantitative

RT‐PCR 93

5.2.9 Standard PCR Conditions 94

5.2.10 Primer Design 94

5.2.11 Reverse Transcription (RT) for qRT‐PCR 94

5.2.12 PCR and Amplicon Purification 94

5.2.13 qRT‐PCR 94


5.3 Results 95

5.3.1 Identification of Differential Gene Expression 95

5.3.2 Gene Ontology and Functional Analysis of Microarray 96

5.3.3 Differentially Expressed Genes as Depicted by Superarray 99

5.3.4 qRT‐PCR Validation of Differentially Expressed Genes 101

5.4 Discussion 112

CHAPTER 6.0 GENERAL DISCUSSION AND CONCLUSIONS 121

6.1 Limitations and Future Directions 132

6.2 Conclusion 137

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CHAPTER 7.0 APPENDICIES 137

Appendix 1 137

Appendix 2 139

Appendix 3 143

Appendix 4 143

CHAPTER 8.0 REFERENCES 145

LIST OF FIGURES

1.1 Hypertrophic and keloid scars 2

1.2 Histological comparison of a normal scar, hypertrophic scar and keloid

scar

4

1.1 Properties of GP226 18

3.1 Analysis of HSF viability following treatment with GP226 and its

fractions.

44

3.2 Analysis of nHSF viability following treatment with GP226 and its

fractions.

45

3.3 Analysis of KF viability following treatment with GP226 and its

fractions

46

3.4 Analysis of nKF viability following treatment with GP226 and its

fractions.

47

3.5 Analysis of HSF proliferation following treatment with GP226 and its

fractions.

48

3.6 Analysis of nHSF proliferation following treatment with GP226 and its

fractions.

49

3.7 Analysis of KF proliferation following treatment with GP226 and its

fractions.

50

3.8 Analysis of nKF proliferation following treatment with GP226 and its

fractions

51

3.9 Analysis of HK viability and proliferation following treatment with

GP226 and its fractions.

54

3.10 Investigation of HSF morphology following culture in media containing

different concentrations of FCS and treatment with GP226.

55

3.11 Analysis of protein aggregation in cell culture medium containing

silicone and varying concentrations of FCS

56

3.12 Analysis of HSF morphology following treatment with Fraction IV and

Tween 20.

58

3.13 Investigation of apoptosis in HSF following treatment with GP226,

Fraction III, Fraction IV and Tween 20.

59

4.1 Molecular weight distribution of the crude GP226 and active fraction,

Fraction IV.

66

4.2 Structure of Fraction IV 66

4.3 Analysis of HSF proliferation following treatment with the synthesised

silicones.

70

4.4 Analysis of nHSF proliferation following treatment with the

synthesised silicones.

71

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4.5 Analysis of HSF proliferation following treatment with GP226, PDMS7‐

g‐PEG7 and PDMS7.

72

4.6 Analysis of nHSF proliferation following treatment with GP226,

PDMS7‐g‐PEG7 and PDMS7.

73

4.7 Analysis of HK proliferation following treatment with GP226, PDMS7‐g‐

PEG7 and PDMS7.

75

4.8 Analysis of HK proliferation, grown without an i3T3 feeder layer,

following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7.

76

4.9 Analysis of HSF morphology following treatment with PDMS7‐g‐PEG7. 78

4.10 Analysis of nHSF morphology following treatment with PDMS7‐g‐PEG7. 79

4.11 Analysis of HK morphology following treatment with PDMS7‐g‐PEG7. 80

4.12 Analysis of HK morphology, grown without an i3T3 feeder layer,

following treatment with PDMS7‐g‐PEG7.

81

4.13 Investigation of apoptosis in HSF following treatment with PDMS7‐g‐

PEG7.

83

4.14 Investigation of apoptosis in nHSF following treatment with PDMS7‐g‐

PEG7.

84

4.15 Investigation of apoptosis in HK following treatment with PDMS7‐g‐

PEG7.

86

4.16 Schematic of the stratum corneum and possible permeation pathways 89

5.1 Validation of up‐regulated target genes in HSF and nHSF following

treatment with GP226 and Fraction IV.

104

5.2 Validation of down‐regulated target genes in HSF and nHSF following

treatment with GP226 and Fraction IV.

105

5.3 Validation of up‐regulated target genes in HSF and nHSF following

treatment with PDMS7‐g‐PEG7.

106

5.4 Validation of down‐regulated target genes in HSF and nHSF following

treatment with PDMS7‐g‐PEG7.

107

5.5 Validation of miscellaneous target genes in HSF and nHSF following

treatment with GP226, Fraction IV and PDMS7‐g‐PEG7.

108

5.6 Validation of up‐regulated, down‐regulated and miscellaneous target

genes in nHSF compared to HSF.

109

6.1 Predicted relationships between differentially regulated genes

identified in HSF and nHSF following treatment with the silicones.

128

6.2 Differential gene expression between HSF and nHSF and the effects on

cellular processes involved in myofibroblast differentiation.

131

LIST OF TABLES

1.1 Summary of aims and experimental approach 21

2.1 Summary of the commercially available polymers used as treatments 24

2.2 Patient data obtained from hypertrophic scar donors

2.3 Primers used for qRT‐PCR 35

4.1 Properties of the synthesised amphiphilic silicones used as treatments 67

5.1 Summary of differential gene expression obtained through microarray

analyses

96

5.2 Ontology of Differentially Expressed Genes in HSF in response to

GP226.

97

5.3 Focus genes identified through microarray analysis to be differentially

expressed in HSF exposed to GP226.

98

5.4 Expression of collagen genes identified through microarray analysis to

be differentially regulated in HSF exposed to GP226

99

5.5 Summary of differential gene expression obtained through superarray

analyses

100

5.6 Focus genes identified through superarray analysis to be differentially

expressed in HSF and nHSF treated with PDMS7‐g‐PEG7

101

5.7 Up‐regulated Focus Genes selected for validation by qRT‐PCR. 102

5.8 Down‐regulated Focus Genes selected for validation by qRT‐PCR. 102

5.9 Miscellaneous Focus Genes selected for validation by qRT‐PCR. 103

6.1 Summary of Results obtained for microarray, superarray and qRT‐PCR

analyses.

127

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LIST OF ABBREVIATIONS

2D 2‐dimensional

αSMA alpha – smooth muscle actin

AIFM2 Apoptosis‐inducing factor, mitochondrion‐associated, 2

bFGF basic Fibroblast growth factor

BSA Bovine serum albumin

cDNA Complementary deoxyribonucleic acid

CD70 Cluster of Differentiation 70

CFLAR Caspase 8‐like apoptosis regulator

COL1A1 Collagen type I, alpha I

COL3A1 Collagen type III, alpha I

ddH2O Double distilled water

dNTP Deoxyribonucleotide triphosphate

DAPI 4',6‐diamidino‐2‐phenylindole

DAPK1 Death‐associated protein kinase 1

DMEM Dulbecco’s modified eagle’s medium

DNA Deoxyribonucleic acid

DNase I Deoxyribonuclease I

DTT Dithiothreitol

ECM Extracellular matrix

FADD Fas‐associating protein with death domain

FAS TNF receptor superfamily, member 6

FCS Fetal calf serum

GP Genesee Polymers

HK Human skin keratinocytes

HLB Hydrophilic‐lipophilic balance

HPLC High performance liquid chromatography

HSF Hypertrophic scar‐derived human skin fibroblasts

i3T3 Irradiated murine feeder layer

IAP Inhibitor of apoptosis proteins

ID Identification

IGFII Insulin‐like growth factor II

IGF2R Insulin‐like growth factor 2 receptor

IL Interleukin

IPA Ingenuity pathway solutions

JNK Jun N‐terminal kinase

KF Keloid scar‐derived human skin fibroblasts

LTβR Lymphotoxin‐β receptor

LTA Lymphotoxin alpha

mRNA Messenger ribonucleic acid

M6P Mannose‐6‐phosphate

MALDI Matrix‐assisted laser desorption/ionization

MCL1 Myeloid cell leukaemia sequence 1

nHSF Normal human skin fibroblasts derived patients suffering from

hypertrophic scar

nKF Normal human skin fibroblasts derived patients suffering from keloid scar

NF‐κB Nuclear factor kappa‐light‐chain‐enhancer of activated B cells

NMR Nuclear magnetic resonance

NSMase Neutral sphingomyelinase

NSMAF Neutral sphingomyelinase activation associated factor

P Probability

PBS Phosphate‐buffered saline

PCR Polymerase chain reactions

PDMS Polydimethyl siloxane

PEG Polyethylene glycol

Primer‐BLAST Primer‐basic local alignment software

PSEC Preparative size exclusion chromatography

qRT‐PCR Quantitative reverse transcription‐polymerase chain reactions

QUT Queensland University of Technology

RNA Ribonucleic acid

rRNA Ribosomal ribonucleic acid

ROS Reactive oxygen species

SEM Standard error of the mean

SGS Silicone gel sheets

SMAD7 SMAD family member 7

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TGFβ Transforming growth factor – beta

THF Tetrahydrofuran

TNF Tumour necrosis factor alpha

TNFR Tumour necrosis factor receptor

TNFR1 Tumour necrosis factor receptor 1

TNFRSF10A Tumour necrosis factor receptor superfamily member 10A

TNFRSF10B Tumour necrosis factor receptor superfamily member 10B

TRADD Tumour necrosis factor receptor‐associated apoptotic signal transducer

TRAF2 Tumour necrosis factor receptor‐associated factor 2

TRAF3 Tumour necrosis factor receptor‐associated factor 3

Trypsin‐EDTA Trypsin‐ethylenediaminetetraacetic acid

WST-1 4‐[3‐(4‐Iodophenyl)‐2‐(4‐nitrophenyl)‐2H‐5‐tetrazolio]‐1,3‐benzene

disulfonate

LIST OF PUBLICATIONS AND PRESENTATIONS

PUBLICATIONS RELATING TO THIS THESIS

Lynam, E., Xie, Y., Loli, B., Dargaville, T. R., Leavesley, D., George, G. and Upton, Z. (2010)

The effect of amphiphilic siloxane oligomers on fibroblast and keratinocyte proliferation

and apoptosis. Journal of Biomedical Materials, Part A; 95(2):620‐31

Lynam, E., Farrugia, B., Keddie, D., Dargaville, T. R., Leavesley, D., George, G. and Upton, Z.

The effect of amphiphilic siloxane oligomers on fibroblast gene expression. Currently under

preparation for submission to Journal of Biomedical Materials, Part A.

OTHER PUBLICATIONS

Xie, Y., Rizzi, S.C., Dawson, R., Lynam, E., Richards, S., Leavesley, D.I. and Upton, Z. (2010)

Development of a Three‐Dimensional Human Skin Equivalent Wound Model for

Investigating Novel Wound Healing Therapies. Tissue Eng Part C Methods. 16(5):1111‐23

Lynam, E. And Upton, Z. (2011) Treatment Innovations in Hypertrophic and Keloid Scars.

Wounds International; 2(1); http://www.woundsinternational.com (Accessed 01/04/2011)

Farrugia, B., Keddie, D., George, G., Lynam, E., Brook, M., Upton, Z. And Dargaville, T. An

Investigation into the Effect of Amphiphilic Siloxane Oligomers on Dermal Fibroblasts.

Currently under review for the Journal of Biomedical Materials, Part A.

PRESENTATIONS

ASMR MRW QLD; May, 2009; Brisbane, Australia; Poster Presentation ‐ Lynam, E.,

McGrath, B., Dargaville, T., Leavesley, D., George, G. and Upton, Z.; “Development and Pre‐

Clinical Evaluation of a Silicone Dressing for Scar Remediation”

5th Joint Meeting of ETRS and WHS; August, 2009; Limoges, France; Poster Presentation ‐

Lynam, E., McGrath, B., Dargaville, T., Leavesley, D., George, G. and Upton, Z.;

“Development and Pre‐Clinical Evaluation of a Silicone Dressing for Scar Remediation”

AHTA ‐ National Hand Therapy Conference; October, 2009; Brisbane, Australia; Poster

Presentation – Lynam, E., McGrath, B., Dargaville, T., Leavesley, D., George, G. and Upton,

Z.; “Development and Pre‐Clinical Evaluation of a Silicone Dressing for Scar Remediation”

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IHBI Inspires; November 2009; Brisbane, Australia; Poster Presentation – Lynam, E., Keddie,

D., Dargaville, T., George, G. and Upton, Z.; “The Effect of Amphiphilic Siloxane Oligomers

on Fibroblast Proliferation and Apoptosis”

2nd Meeting of AWTRS; March, 2010; Perth, Australia; Oral Presentation ‐ Lynam, E.,

Keddie, D., Dargaville, T., George, G. and Upton, Z.; “The Effect of Amphiphilic Siloxane

Oligomers on Fibroblast Proliferation and Apoptosis”

IHBI Inspires; November, 2010; Gold Coast, Australia; Oral Presentation – Lynam, E.,

Keddie, D., Dargaville, T., George, G. and Upton, Z.; “Evaluation of a Novel Silicone for

Hypertrophic Scar Remediation: The Effect Observed at Gene Level”

L i t e r a t u r e R e v i e w

C h a p t e r 1 . 0 | 1

CHAPTER 1.0

LITERATURE REVIEW

1.1 SKIN HOMEOSTASIS

The primary functions of skin are to physically protect the body from the environment and

to mediate repair following wounding (Bouwstra and Ponec, 2006). Tissue growth and

repair occurs through a series of overlapping events involving many cell types, extracellular

matrix (ECM) components, cytokines and soluble mediators (Tuan and Nichter, 1998).

Specifically, wound healing is aimed at rapidly restoring barrier function (Martin, 1997).

While injuries in utero evoke embryonic processes that rapidly re‐establish natural

homeostasis to the skin without scarring, healing in adults is a much slower process and

involves robust inflammation, increased neovascularisation, as well as increased

extracellular matrix deposition (Mutsaers et al., 1997; Reish and Eriksson, 2008a). This

often does not restore the normal structure of tissue. Consequently, scarring often

manifests as macroscopic disfigurations to the skin following trauma, surgery, burns or

injury where the normal tissue structure is not remodelled perfectly after healing (Ferguson

and O'Kane, 2004; Ferguson et al., 1996).

Many processes occur in the skin during the formation and remediation of scars. Three

particular stages of repair, which are known as inflammation, tissue formation

(granulation) and remodelling arise as normal processes in the adult human body (Singer

and Clark, 1999). Of critical importance to scar resolution is the final remodelling stage,

involving the restoration of collagen equilibrium following previous over‐synthesis during

granulation. This stage of collagen turnover, involving collagenase activity and the

deposition of new and more organised collagen is vital in scar remediation and may take up

to 6‐24 months after the initial trauma to the skin (Mutsaers et al., 1997).

1.2 ABNORMAL SCARS

While in most cases scars recede as a natural part of wound healing, several fail to do so,

producing adverse effects such as undesirable appearance, itching, loss of function,

restriction of tissue movement or growth and psychological effects (Ferguson and O'Kane,

2004; Mutsaers et al., 1997). These scars can be classified as either keloid or hypertrophic

and are defined as benign hyperproliferative growths of dense fibrous tissue (Robles and


Berg, 2007). Clinically, keloid scars grow beyond the confines of the original wound and

invade surrounding tissue, generating a lesion that appears irregular and pendulous

(O'Brien and Pandit, 2006; Tuan and Nichter, 1998). Conversely, hypertrophic scars are

raised, yet remain within the original wound boundary, and have a tendency to appear and

regress spontaneously (Tuan and Nichter, 1998). Figure 1.1 illustrates an example of

hypertrophic and keloid scars.

A

B

Figure 1.1 – Hypertrophic and Keloid Scars (A) Hypertrophic scaring the neck region of a 12 year old female as a result of a chemical burn injury; (B) Ear lobe keloid in a 22 year old female as a result of ear piercing (Tuan and Nichter, 1998)

Hypertrophic scars tend to originate from surgery and thermal injuries such as burns

(Carney et al., 1994; Eisenbeiss et al., 1998; Shakespeare, 1993), while keloids often

develop after trivial injuries including ear piercing, insect bites and vaccination (O'Sullivan

et al., 1996). Interestingly, while hypertrophic scars are known to develop at any location,

keloid scars commonly affect chest, shoulder and ear lobe regions, which are areas under

low skin tension (Crockett, 1964; Marneros et al., 2001). It has been argued that skin or

wound tension is implicated in the formation and location of abnormal scars. Other

C h a p t e r 1 . 0 | 3

predisposing conditions of both hypertrophic and keloid scars appear to be related to

prolonged inflammation, such as repeated trauma, continued irradiation from foreign body

inclusions, excessive wound tension, infection or delayed reepithelialisation (Deitch et al.,

1983). Epidemiology is another factor involved in hypertrophic and keloid scar formation

but research relating to their relationship is limited. The data that is available suggests

differences among racial groups (Robles and Berg, 2007) and it has been reported that

higher rates of keloid scar formation occur in dark‐skinned individuals compared to those

who have a lighter skin colour (Brissett and Sherris, 2001). While there are no clearly

defined genetic loci linked to keloid scars, it is widely accepted that their etiology involves a

genetic factor (Robles and Berg, 2007). Additionally, a study by Marneros et al., (2001)

investigated the clinical and inheritance patterns of keloid scar development. Interestingly,

their data suggested that a child had a 50% chance of keloid scar development if one of

their parents had a keloid scar. No such studies have investigated this genetic association

with hypertrophic scar formation as yet (Brown and Bayat, 2009).

1.3 PATHOPHYSIOLOGY OF ABNORMAL SCARS

Both hypertrophic and keloid scars develop from an exaggerated fibroproliferative

response within the dermis, which creates an imbalance of matrix production and collagen

synthesis and results in an excessive accumulation of dermal collagen, fibronectin and

glycosaminoglycan content (Babu et al., 1989; Reish and Eriksson, 2008a). The most

prominent of these factors in scar formation is the excessive deposition of collagen types I

and III (Abergel et al., 1985; Rockwell et al., 1989). Even after many years, equilibration,

and hence elasticity, is not restored to the skin (Alster and Tanzi, 2003; Haukipuro et al.,

1991). One underlying reason is that wound fibroblasts undergo phenotypic change within

the granulation tissue to become more proliferative myofibroblasts. These cells up‐regulate

collagen production, resulting in scar development (Darby and Hewitson, 2007; Gabbiani et

al., 1971). Some data also suggests that reduced collagen degradation of newly synthesised

procollagen polypeptides may possibly contribute to the increased collagen deposition in

keloid and hypertrophic scars (Abergel et al., 1985; Arakawa et al., 1996). The alignment of

collagen is also considered to be different when comparing hypertrophic and keloid scars to

normal skin. Whorls of closely packed, thickened hyalinised collagen bundles classically

appear in keloid scars, while hypertrophic scars exhibit modular structures in which

fibroblastic cells, small vessels, and fine, randomly organised collagen fibres are present

(Ehrlich et al., 1994; Lee et al., 2004). The compacted accumulations of collagen as well as


the excess of myofibroblasts in hypertrophic and keloid scars compared to a normal scar

are depicted in Figure 1.2.

Figure 1.2 ‐ Histological comparison of a normal scar, hypertrophic scar and keloid scar. Histology of a (A) normal scar, (B) hypertrophic scar and (C) keloid scar, stained with haematoxylin and eosin. Collagen bundles in the dermis of normal scar tissue appear relaxed and arranged randomly. By contrast, collagen bundles in hypertrophic scars and keloid scars are thicker and are aligned in the same plane as the epidermis. Collagen bundles within keloid scars are also formed in acellular node‐like structures. The presence of cells within the epidermis and dermis of the hypertrophic scar and keloid scar is much more dense than when compared to the normal scar (Tuan and Nichter, 1998).

Recent studies of human fetal wound healing have further suggested that high levels of

inflammation, which occurs in adult but not in embryonic wound healing processes, may be

a major cause of abnormal scar formation (Reish and Eriksson, 2008a). Furthermore, an

increased number of macrophages, epidermal Langerhans’ cells and mast cells, all which

are involved in inflammatory processes, have been reported in hypertrophic and keloid

scars, although their contribution to scar formation is not known (Smith et al., 1987;

Tanaka et al., 2004). It is evident that further research in this area is required.

C h a p t e r 1 . 0 | 5

It is clear that cellular events must be altered in order for hypertrophic and keloid scars to

develop. Many have investigated this area and thus far it has been illustrated that

fibroblasts derived from abnormal scars not only overproduce collagen but express higher

levels of interleukin (IL)6, vascular endothelial growth factor, transforming growth factor

beta (TGFβ) and platelet derived growth‐factor‐α receptors as well (Ghazizadeh et al.,

2007; Marneros et al., 2001). Cytokines such as TGFβ1 and TGFβ2 are central to wound

repair as they are potent activators of extracellular matrix gene expression and stimulate

collagen and fibronectin synthesis by dermal fibroblasts (Beanes et al., 2003). Evidently, the

regulation of these cytokines is of major importance in the etiology of hypertrophic and

keloid scars (Bettinger et al., 1996; Ghahary et al., 1993).

1.3.1 Dermal Fibroblast Heterogeneity

It has been widely reported that the composition of connective tissue is altered in

hypertrophic and keloid scars. For instance, myofibroblast number is increased in

hypertrophic scars (Nedelec et al., 2001) and the differentiation of wound fibroblasts into

myofibroblasts represents a key element in wound healing and tissue repair (Darby and

Hewitson, 2007; Desmouliere et al., 2005; Ehrlich et al., 1994; Gabbiani et al., 1971; Hinz,

2007; Tiede et al., 2009). Myofibroblasts are characterized by the expression of alpha‐

smooth muscle actin (αSMA) and are thought to derive from fibroblast populations under

stimulation of profibrotic growth factors such as TGFβ1 (Desmouliere et al., 2005; Narine et

al., 2006; Wang et al., 2008). Tiede et al. (2009) demonstrated that basic fibroblast growth

factor (bFGF) efficiently inhibited the terminal differentiation of mesodermal progenitors

into myofibroblasts with a reduction in the level of αSMA positive cells being detected and

a subsequent decrease in αSMA expression at the ribonucleic acid (RNA) and protein level

following treatment. Furthermore, this inhibitory effect of bFGF was accompanied by an

inhibition of TGFβ receptor I and II expression. These results are significant as up until

recently, little had been reported regarding the influence of growth factors on excessive

myofibroblast formation and the associated development of hypertrophic scars, keloids

and fibrosis (Tiede et al., 2009).

It has also been demonstrated that normal adult human skin contains at least three distinct

subpopulations of fibroblasts that occupy unique niches in the dermis. Furthermore

fibroblasts from each of these niches exhibit distinctive differences when cultured

separately in vitro (Sorrell and Caplan, 2004; Wang et al., 2008). For example, it was


observed by Wang et al. (2008) that fibroblasts isolated from deeper layers of the dermis

were morphologically larger than those of the superficial layers. These deep layer

fibroblasts were also demonstrated to proliferate more slowly, produce less collagenase

but generate more collagen, TGFβ1 and αSMA. From these results, it was suggested that

fibroblasts isolated from deep layers in the dermis may play a critical role in the

development of hypertrophic scars (Wang et al., 2008). Another area of interest has been

uncovered in blood‐born fibroblast‐like cells called fibrocytes (Bucala et al., 1994).

Circulating fibrocytes migrate to wound sites during the inflammation phase of wounds and

are capable of producing proinflammatory cytokines, chemokines, growth factors and

extracellular matrix (Chesney et al., 1998; van der Veer et al., 2009). It is thought that

fibrocytes may be a precursor of myofibroblasts and several studies have suggested that

fibrocytes are implicated in wound healing and hypertrophic scar formation (Mori et al.,

2005; Yang et al., 2005).

1.3.2 Role of Epidermis in Wound Healing

As mentioned previously, tissue growth and repair occurs through a series of overlapping

events involving many cell types, ECM components, cytokines and soluble mediators (Tuan

and Nichter, 1998). As wound healing is a complex process, it is logical that close paracrine

relationships exist between the epidermis and dermis (Maas‐Szabowski et al., 1999; Tuan

and Nichter, 1998). It has been extensively reported that wound healing is aimed at rapidly

restoring barrier function through the formation of a functional epidermis (Martin, 1997).

However, it has also been reported that keratinocytes are dependent on signals received

from dermal fibroblasts to re‐establish the functional epidermal barrier required during the

reepithelialisation phase of wound healing (El Ghalbzouri et al., 2002; Ghahary and

Ghaffari, 2007). Additionally, it has recently been reported that delays of greater than 21

days in reepithelialisation during healing increases the likelihood of developing fibrotic

conditions, such as a hypertrophic or keloid scar, to 78% (El Ghalbzouri et al., 2002;

Ghahary and Ghaffari, 2007). Furthermore, keratinocyte‐derived growth factors have been

suggested to play a role in the formation of abnormal scars (Katz and Taichman, 1994;

Niessen et al., 2001) and a study by Funayama et al. (2003) demonstrated that a co‐culture

of normal fibroblasts with keloid keratinocytes resulted in collagen being produced in a

keloid‐like manner. It is clear that the cross‐talk of cells between the epidermis and dermis

plays an important role within wound healing.

C h a p t e r 1 . 0 | 7

1.3.3 Scarring and Apoptosis

Apoptosis is another process known to play a significant role in wound healing. The number

of apoptotic cells sharply increases as the wound closes, suggesting that apoptosis is the

mechanism responsible for the evolution of granulation tissue into a scar (Darby et al.,

1990; Desmouliere et al., 1995). Failure of apoptosis has often been postulated in the

pathogenesis of abnormal scars but the evidence presented in the literature is

controversial and often contradictory (Linge et al., 2005). For example, levels of apoptotic

cells were relatively low in tissue sections of normal and hypertrophic scars but were

significantly higher in keloid scar tissue sections (Akasaka et al., 2001). However, a study by

Messadi et al. (1999) demonstrated that fibroblasts derived from keloid scars showed

lower rates of apoptosis than normal fibroblasts.

Despite these controversial results, decreased apoptosis is postulated to cause the

hypercellularity and excess scar tissue formation of hypertrophic scars. Not surprisingly,

lower rates of apoptosis and a down‐regulation of apoptosis‐related genes have been

demonstrated by fibroblasts derived from hypertrophic, as well as keloid scars (Appleton et

al., 1996; Nedelec et al., 2001; Sayah et al., 1999). In fact, gene expression profiling of

normal skin derived from keloid‐prone and keloid‐resistant patients revealed differences in

levels of two caspases, 6 and 14 (Nassiri et al., 2009). It has been widely reported that

caspases, a family of cysteine proteases, play an important part in apoptosis and the

activation of these caspases is required for apoptosis to be executed (Lamkanfi et al., 2007;

Nassiri et al., 2009). It was suggested by Nassiri et al. (2009) that the distinct gene profile of

keloid‐prone patients could be important in their susceptibility as the familial clustering of

patients suffering from keloid scars has long been attributed to an underlying genetic

predisposition (Alhady and Sivanantharajah, 1969). However, the molecular mechanisms

behind this genetic involvement have not been well understood. Moreover, as caspase 14

is expressed by keratinocytes, but not fibroblasts, the hypothesis that dermal scarring may

be regulated by factors produced in the epidermis is strengthened (Nassiri et al., 2009).

A study by Linge et al., (2005) illustrated that reducing tissue transglutaminase activity in

collagen gels containing fibroblasts derived from hypertrophic scars allowed induction of

apoptosis on gel contraction. In contrast, increasing enzymatic activity of collagen gels

containing fibroblasts derived from normal skin completely abrogated collagen contraction‐

induced apoptosis. Newly formed extracellular matrix is stabilized by the enzyme, tissue


transglutaminase, which has been demonstrated to be over expressed by hypertrophic

scar‐derived fibroblasts in vivo and in vitro (Linge et al., 2005). It was suggested from these

results that hypertrophic scar‐derived fibroblasts may exhibit resistance to a specific form

of apoptosis that is dependent on the excess activity of cell surface tissue transglutaminase

(Linge et al., 2005). Furthermore, transglutaminase has been implicated in the activation of

TGF‐β1 via the cross‐linking of latent TGF‐β‐binding protein‐1 (Nunes et al., 1997; Verderio

et al., 1999). TGF‐β mediated apoptosis has been demonstrated during tissue formation

and remodelling in cutaneous wound healing (Crowe et al., 2000). It is evident that the

potential involvement of transglutaminase in activation of such an important profibrotic

protein as TGF‐β, combined with its overexpression in hypertrophic scars, has obvious

implications for the pathology of abnormal scars (Linge et al., 2005).

Understanding the relationships between scarring and apoptosis also extends to include

the investigation of mechanical stress as an in vitro model of granulation tissue. For

example Aarabi et al., (2007) demonstrated that mechanical stress applied to healing

wounds in p53‐null mice was sufficient to produce hypertrophic scars. The model resulted

in scars that were structurally similar to human hypertrophic scars, showing dramatic

increases in volume (20‐fold) and cellular density (20‐fold), which was accompanied by a 4‐

fold decrease in cellular apoptosis and increased activation of the prosurvival marker Akt.

Furthermore, scar hypertrophy and cell density was significantly reduced in proapoptotic

Bc/II‐nt null mice (Aarabi et al., 2007). In addition, it was illustrated by Grinnell et al. (1999)

that fibroblasts under mechanical tension showed little or no apoptosis. However, when

mechanical tension was released, an apoptotic response was triggered within 3‐6 hours.

1.4 MANAGEMENT OF ABNORMAL SCARS

As current aesthetic surgical techniques become more standardised and results more

predictable, a fine scar is becoming the demarcating line between acceptable and

unacceptable results (Widgerow et al., 2000). Consequently, current management of

hypertrophic and keloid scars include a wide range of techniques, from traditional invasive

methods to intralesional and topical application of agents designed to take effect on a

cellular level (Reish and Eriksson, 2008a). Some of these techniques, which have been

reported as being beneficial in the management of abnormal scars, include surgery,

corticosteroid injections (Darzi et al., 1992), pressure therapy (Staley and Richard, 1997),

C h a p t e r 1 . 0 | 9

radiotherapy (Ogawa et al., 2003), laser therapy (Goldman and Fitzpatrick, 1995),

cryotherapy (Zouboulis et al., 1993) and silicone gel therapy (Perkins et al., 1983).

Many researchers have also believed for some time that efforts at controlling scar outcome

should be initiated at the time of wounding, when the trigger for the sequence of events

essential for healing begins (Ono et al., 2007; Shah et al., 1994; Widgerow et al., 2009). It

has been widely reported that the prevention of hypertrophic and keloid scars is much

more effective than their later treatment, especially in patients predisposed to scarring

(Alster and Tanzi, 2003; Katz, 1995). However, until now, treatment is routinely

commenced when the wound has reepithelialised, rather than at the time of injury.

1.5 SILICONE GEL TREATMENTS

Silicone gel sheets (SGS) have been used as a topical treatment for hypertrophic and keloid

scars since 1983 (Perkins et al., 1983). These commercially available sheets consist of lightly

crosslinked polydimethyl siloxane (PDMS) (Van den Kerckhove et al., 2001) and reportedly

reduce the size, redness, pain and itching of scars (Berman and Flores, 1999; Eishi et al.,

2003), as well as promote increased skin elasticity (Ahn et al., 1989), softness (Quinn et al.,

1985) and even prevent scar formation (Gold et al., 2001). The flexible ‐Si‐O‐Si‐ backbone

structure, along with surrounding hydrophobic methyl groups that is attributed to PDMS

polymers, confer the strong and flexible properties of SGS as well as high oxygen

permeability, low surface energy and resistance to hydrolytic scission (LeVier et al., 1993).

Furthermore, PDMS oligomers of lower chain length are also trapped into the crosslinked

network (LeVier et al., 1993).

The use of SGS is well documented in the literature and an early uncontrolled study by

Quinn (1987) reported an improvement of scars in 81% of the 125 treated patients.

Another uncontrolled analysis was performed by Katz (1995), who evaluated the use of SGS

on both newly formed and chronic hypertrophic and keloid scars, and reported

improvement rates of 79% and 56% respectively. Subsequent controlled studies by Ahn et

al. (1991), Sproat et al. (1992), Carney et al. (1994), Gold (1994), Fulton (1995) and Lee et

al. (1996) have yielded similar results. More recently, Momeni et al. (2009) demonstrated

that silicone gel was an effective treatment for hypertrophic burn scars. The significance

behind this is that the surface of burn wounds differs from other wounds in that they are

characterised by coagulation of the superficial blood vessels and usually do not tend to


bleed excessively, whereas normal wounds do (van der Veer et al., 2009). This finding was

important as the majority of reported literature has investigated the effect of SGS on

hypertrophic scars resulting from injuries other than burns.

1.5.1 Mechanism of Silicone Gel Sheet Action

Despite considerable research effort worldwide, the mechanism of SGS action is yet to be

completely elucidated (Borgognoni, 2002). The early study by Quinn (1987) showed that

the outcomes of SGS were not due to pressure, temperature or oxygen tension. This has

led other investigators to hypothesise that SGS action results from increased temperature

(Krieger et al., 1993), improved hydration, higher mechanical and bacterial protection

(Chang et al., 1995), electrostatic changes (Hirshowitz et al., 1998) and the movement of

chemical species into the skin (Shigeki et al., 1999).

In reviewing SGS literature it has been stated that the occlusion and hydration of

hypertrophic and keloid scars is therapeutic and that these effects can be obtained with

both silicone and non‐silicone dressings (De Oliveira et al., 2001; Ricketts et al., 1996;

Sawada and Sone, 1992; Wittenberg et al., 1999). For example, it has been illustrated that

the exposure of keratinocytes to hydrated conditions causes significant inhibition of

fibroblast proliferation and their production of collagen and glycosaminoglycans, while

exposure of keratinocytes to silicone oil or liquid paraffin does not influence fibroblast

behaviour (Chang et al., 1995). Furthermore, Sawada and Sone (1992) successfully treated

hypertrophic scars and keloids with an occlusive dressing technique using cream which did

not contain silicone oil. Conversely, Suetake et al. (2000) demonstrated that the magnitude

of SGS hydration was less than that of plastic film occlusion, with SGS producing a more

favourable skin condition similar to the natural stratum corneum state and also providing

mechanical protection to the area. A subsequent study using a rabbit model of

hypertrophic scarring illustrated that a polyurethane dressing, of similar occlusive nature to

SGS, or Tegaderm (3M Healthcare), a dressing approximately five times less occlusive, did

not have the same beneficial effects as SGS on abnormal scarring (Saulis et al., 2002). It was

concluded from this study that while occlusion and hydration were essential to minimise

scarring, they were not sufficient to explain SGS action.

Quinn et al. (1985) reported that the presence of silicone oil, continually released from

SGS, was the important factor, rather than hydration, in its mechanism of action. However,

C h a p t e r 1 . 0 | 11

Sawada and Sone (1990) found that a cream containing 70% silicone oil was statistically

successful in treating 82% of hypertrophic and keloid scars only when used in conjunction

with an occlusive dressing. This suggested that the dressing itself was still important in the

healing process. The same study also showed that earlier treatments produced better

outcomes, suggesting that silicone materials have a much less dramatic effect on mature

scars (Niessen et al., 1998; Sawada and Sone, 1990).

Another interesting potential mechanism by which SGS could reduce scars is through the

regulation of fibrogenic cytokines (Reish and Eriksson, 2008b). Kuhn et al. (2001)

demonstrated that hypertrophic scar fibroblasts in a populated collagen lattice showed

decreased contraction and TGF‐β2 expression when exposed to silicone sheeting than

when compared to an unexposed control. Furthermore, an earlier study examining dermal

cytokine messenger RNA (mRNA) levels in silicone gel‐treated hypertrophic scars found

that treatment of hypertrophic scars with silicone or occlusive dressings resulted in

increased mean levels of bFGF, IL8 and granulocyte‐macrophage colony‐stimulating factor,

as well as decreased levels of TGF‐β and fibronectin mRNA (Ricketts et al., 1996). The

isoform of TGF‐β analysed in this study was not specifically identified by the authors. This

result was confirmed by Hanasono et al. (2004), who also illustrated that bFGF was

increased in fibroblast cell cultures of normal, keloid and fetal skin after exposure to

silicone gel. While Hanasono et al. (2004) did not compare their results with a non‐silicone

treatment, the findings by Ricketts et al. (1996) agree with other literature stating that

silicone is not a necessary component of occlusive scar dressings. Nevertheless, as

Hanasono et al. (2004) hypothesized, it is quite possible that substances favourably

influencing wound healing do so by correcting a deficiency, or over‐abundance, of the

growth factors that orchestrate tissue repair processes.

Additional studies have also proposed that static electricity generated by friction‐activated

silicone sheeting could be the reason for the effects of SGS (Amicucci et al., 2005;

Hirshowitz et al., 1998). Berman and Flores (1999) investigated this hypothesis but no

difference between hypertrophic and keloid scar improvement was demonstrated when

treated with a silicone gel‐filled cushion of high static electricity and traditional SGS.

Nevertheless, analysis of reports reveals in all cases poor methodology with no placebo or

control treatments, scar categorization or randomization evident. Consequently, there is no


current evidence supporting the concept that static electricity causes higher rates of scar

involution.

Increased perfusion and temperature are other mechanisms being investigated as having

important roles in the treatment of abnormal scars with silicone gel. Musgrave et al.

(2002) reported that hypertrophic scars displayed higher levels of perfusion than normal

skin but no change was found after treatment with silicone gel. Despite this, the same

analysis demonstrated that the application of SGS significantly increases the mean baseline

surface temperature of hypertrophic scars from 29 ± 0.8 oC to 30.7 ± 0.6 oC (Musgrave et

al., 2002). While the magnitude of this result seems minimal, temperature differences of

less than 1 oC have been reported to cause significant effects on collagenase kinetics, which

in turn may alter scarring (Krieger et al., 1993; Lyle, 2001). It should be noted, however,

that Musgrave et al. (2002) did not include any control scar treatments in the study and

whether an occlusive dressing containing no silicone could have the same effect was not

investigated. No further research has been reported examining the role of temperature on

abnormal scarring and thus, it has not yet been proven to be the primary mechanism of

SGS action.

1.5.2 The Role of Silicone in Scar Prevention

As stated previously, preventing hypertrophic and keloid scar formation is a much more

effective strategy than their later treatment (Alster and Tanzi, 2003; Katz, 1995).

Consequently, a large body of research has been directed at investigating the use of

silicone in scar prophylaxis. For example, it was claimed by Fulton (1995) that young scars

of less than three months duration did not develop abnormally when treated with silicone

as soon as reepithelialisation had occurred. Another report demonstrated that SGS applied

to wounds 2 weeks post‐operatively for 12 hours per day resulted in significantly decreased

scar volume after 2 months, compared to controls treated without silicone; this result was

confirmed with a 6 month follow‐up study (Cruz‐Korchin, 1996). However, this investigation

included only 20 patients, and a larger but similar controlled study involving 96 patients,

who were divided into groups of low and high risk, was performed by Gold et al. (2001).

This latter study showed no difference between treatments in the low risk groups, but

larger differences between treatments in the high risk groups and even more significant

improvements in those undergoing scar revision surgeries. In contrast, an additional study

of 129 breast reduction patients demonstrated no improvement in scar prevention with

C h a p t e r 1 . 0 | 13

the use of SGS at a 1 year assessment (Niessen et al., 1998). Reasons for these results were

suggested by the authors as due to early initiation of SGS treatment, with therapy

commencing immediately after surgery before the wound had reepithelialised.

1.5.3 Adverse Effects of Silicone Treatment

Although the safety, effectiveness and ease of SGS use are generally accepted (Shigeki et

al., 1999), it is a treatment with many limitations. For example, SGS require consistent and

hygienic application for at least twelve hours a day for up to twelve consecutive months,

beginning as soon as reepithelialisation has occurred (Ahn et al., 1991). In addition, some

parts of the body, such as joints or large areas suffering from abnormal scarring, are not

suitable for SGS use. Furthermore, taping is often required to secure SGS to the skin and

patients may be reluctant to use the treatment on unclothed areas during the day (Mustoe,

2008).

Many reports have demonstrated adverse effects of utilising SGS on scars. It was reported

by Nikkonen et al. (2001) that high frequency problems were associated with SGS when

used to remediate scars in Saudi Arabia, an area of characteristically high temperatures and

aridity. Patients in this study suffered persistent pruritus, skin breakdown, skin rash, skin

maceration and even foul smells arising from the gel. Nevertheless, all patients except one

were able to continue treatment with temporary cessation or improved hygiene measures.

Other reports have also documented rash, skin breakdown, problems with application and

poor gel durability, especially when SGS is used in paediatric care (Gibbons et al., 1994). It

is estimated that the complication rate is higher than reported in the literature and the

reasons behind these adverse effects are numerous, including poor patient hygiene and

often inappropriate application, especially when involving children (Rayatt et al., 2006).

1.5.4 Type of Silicone Gel Treatment

Numerous silicone treatments are currently available for scar remediation but there is little

information in the literature comparing their efficacy. One study of interest was performed

by Carney et al. (1994), who compared the efficacy and safety of two commonly used SGS

treatments, Cica‐Care (Smith and Nephew) and Silastic Gel Sheeting (Dow Corning). It was

demonstrated that there was no difference in scar improvement between the two

treatments, although it was suggested that patients tolerated Cica‐Care better because of

its improved comfort and adhesiveness. Furthermore, Lee et al. (1996) compared Sil‐K


(Degania Silicone Ltd) and Epiderm (Biodermis) in the treatment of abnormal scars and

reported that both performed equally, with each having different optimal factors, including

cost, durability, conformity and hygiene, that should be taken into consideration when

choosing treatment.

Until the adverse effects of SGS began to be publicised, the physical structure of SGS had

rarely been investigated and preliminary studies into the use of cushions, spreadable and

self‐drying products, as well as controlled‐release, silicone‐based emulsions have only

recently begun to be published (Bott et al., 2007; Hirshowitz et al., 1998; Signorini and

Clementoni, 2006). A recent evaluation of a new spreadable, self‐drying silicone gel was

performed by Signorini and Clementoni (2006) and it was reported that fresh surgical scars

treated with a spreadable, self‐drying silicone gel fared better than those which were

untreated in a study of 160 patients. It was noted that patient compliance was improved

because, unlike traditional SGS, the silicone gel was transparent and did not need fixation.

These are two advantageous factors when scars are located on exposed areas such as the

face. Chan et al. (2005) performed a similar study comparing the use of Dermatix (Valeant

Pharmaceuticals International), a commercially available spreadable silicone gel, to a

placebo gel. The results from this analysis revealed that the scars developed during silicone

gel treatment, 3 months after surgery, were significantly flatter, more pliable, as well as

less itchy and painful than the control scars. Furthermore, another study evaluated the use

of Dermatix and SGS in scar remediation, with comparable results being reported between

the two treatments and both showing higher scar resolution compared to non‐treated

control scars. Once again, patients rated Dermatix as easier to use than the conventional

SGS (Chernoff et al., 2007).

1.5.5 Comparing other Therapies to Silicone

As noted previously, many therapies other than SGS are available for hypertrophic and

keloid scar remediation. Indeed, many reports exist comparing the use of SGS to these

other treatments. One such study by Tan et al. (1999) compared SGS to intralesional

injections of triamcinolone acetonide in the treatment of keloid scars and reported that

SGS did not perform as well. The study, however, only investigated keloid scars that were

more than 2 years old. Early therapy of scars, especially with SGS, produces more optimal

results compared to scars which are treated with SGS later. Furthermore, Sproat et al.

(1992) compared the same treatments in hypertrophic scars and demonstrated that SGS

C h a p t e r 1 . 0 | 15

provided earlier symptomatic relief and more aesthetic scars, compared to scars treated

with intralesional injections only. Comparisons have also been made between SGS and

laser therapy for the treatment of abnormal scars (Paquet et al., 2001; Wittenberg et al.,

1999). Although Wittenburg et al. (1999) investigated hypertrophic scars and Paquet et al.

(2001) evaluated keloid scars, neither study found any significant improvement after

treatment with either SGS or 585 nm flashlamp‐pumped pulsed dye lasers compared to the

control scars.

The idea of combining therapies has also been postulated by clinicians. One such analysis

demonstrated that the addition of vitamin E to silicone gel treatments improved scar

remediation to a greater extent than SGS itself in a simple‐blinded study (Palmieri et al.,

1995). Moreover, Li‐Tsang et al. (2006) found that SGS along with deep massage

significantly reduced the thickness and improved the pliability of hypertrophic scars

compared with massage alone for six months.

1.5.6 Limitations of Research

While a great deal of evidence concerning the efficacy of SGS is available, no specific factor

has been proven to be the major contributor or inducer of the optimal conditions

attributable to SGS. Although the mode of SGS action is most likely a combination of

chemical and mechanical effects, this is a significant knowledge gap in this field of research.

Moreover, differences among the studies are apparent and the various brands of SGS,

subjective analysis methods used, along with the lack of acceptable controls are some

factors requiring further consideration. One report admitted that regardless of the

inclusion of controlled groups, it is still difficult to evaluate scar treatment results because

non‐objective methods for measurements, such as scar colour and induration, both of

which are subjective classifications, are used (De Oliveira et al., 2001). Furthermore, a

recent review performed by O’Brien and Pandit (2006) found that only two studies,

performed by Sproat et al. (1992) and Wittenburg et al. (1999), out of all published

controlled silicone gel trials, met four or more of the seven quality criteria of methodology.

An example of the quality criteria included: was the assessor of the primary outcome

masked to treatment allocation; were data from all randomised participants included in the

analysis; and was there a valid assessment of comparability of the study groups at baseline

(O'Brien and Pandit, 2006)? It was also made clear by O’Brien and Pandit (2006) that only

three studies (De Oliveira et al., 2001; Gold, 1994; Niessen et al., 1998) distinctly classified


patients suffering from hypertrophic or keloid scars and hence subsequently discussed the

results separately. The literature examined in this review has not revealed any other

controlled trials that have met these criteria.

Another aspect is that the follow‐up of silicone gel trials should ideally continue for at least

one year after treatment because the processes of wound healing and scarring are long‐

term events (Shaffer et al., 2002). The current literature shows that six studies (Ahn et al.,

1989; Cruz‐Korchin, 1996; Gold et al., 2001; Palmieri et al., 1995; Sproat et al., 1992; Tan et

al., 1999) have followed patients for less than six months post treatment and only two

(Carney et al., 1994; Niessen et al., 1998) have reported follow‐up examinations twelve

months post treatment. Furthermore, due to the large variety of commercial silicone gel

treatments available for scar remediation, there is a lack of uniformity between the trials

performed. Taking these facts into account, it is clearly evident that there remains a great

need for additional clinical studies using well‐designed, double‐blinded, placebo‐controlled,

multicenter randomized trials with objective and standardized evaluation measures to

assess SGS efficacy (Reish and Eriksson, 2008a).

With respect to studies investigating the role of apoptosis in wound healing only,

discrepant findings have been discussed as resulting from differences in experimental

procedures, such as whether cultured cells or tissue were examined, the maturity of scar

tissue and methods used to induce apoptosis, some of which bear no relation to the

physiological processes within the maturing wound (Linge et al., 2005). These differences

are also evident in other areas, such as SGS treatment of abnormal scars.

Progress to minimise the limitations of clinical scar assessments are being made, however.

For example, elements of different scales were combined by Widgerow et al. (2009).

Specifically, they investigated a multimodal approach to scar management aimed at

controlling scar tension, hydration, collagen maturation and inflammation, and

incorporated these aspects into the assessment parameters of their study. Thus elements

of the Vancouver Scar Scale (Baryza and Baryza, 1995) and the Scale of Morphologic

Features (Signorini and Clementoni, 2006) were combined with additional patient and

observer assessments to give a more complete measurement of outcomes (Widgerow et

al., 2009).

C h a p t e r 1 . 0 | 17

1.6 RETURNING TO THE BEGINNING

The volume of inconsistent information indicates that current studies should perhaps

return to hypotheses made early in SGS history, specifically Quinn’s claim in 1987 that

SGS’s action must involve a chemical factor. It was determined at that time that further

research was essential to investigate whether silicone species could penetrate the stratum

corneum, interact with the dermal fibroblasts present and critically alter the chemical

composition of scars (Quinn, 1987). A study performed soon after this hypothesis by Ahn et

al. (1989) found no histological evidence of silicone in biopsies of SGS‐treated scars. This

result was strengthened by Fulton who published in 1995 that while SGS (Epi‐derm) did

improve 85% of hypertrophic and keloid scar cases, no silicone was found at the wound

site. However, an investigation by Shigeki et al. (1999) measured the amount of silicone

distributed from SGS (Cica‐Care) into a phosphate buffer solution and found that

concentration increased with time, depending on the pH of the medium. In addition,

silicone was detected in all skin samples after application of SGS (Cica‐Care) to excised rat

skin, human axilla skin and hypertrophic scars in vitro. It was concluded from this that

pharmacological effects could possibly result from silicone migration, affecting fibroblasts

and subsequent scar tissue formation (Shigeki et al., 1999). In spite of this, it has never

actually been proven that silicones have a functional influence on scar development,

further illustrating the lack of knowledge in the area.

1.6.1 Research undertaken at QUT

Research undertaken by our laboratory has re‐investigated the chemical role of SGS on

scarring. It has been demonstrated that small amounts of linear, oligomeric silicones with

low molecular weight have the ability to migrate from commercially available SGS into the

human stratum corneum (Sanchez et al., 2005). It was hypothesized from these data that

the slow rate of silicone diffusion from SGS into the skin may explain why such long

treatment times are required to achieve the desired results on scars using the products

currently available commercially. In view of this, our research team screened a large

number of commercially available oligomeric silicones (sourced from Genesee Polymers)

for their ability to attenuate collagen synthesis in human skin fibroblasts derived from

hypertrophic scars (HSF). The most effective species were found to be amphiphilic, low

molecular weight polyether surfactants of approximately 3000 Daltons, encompassing a

rake PDMS structure with side branches of polyethylene glycol (PEG). The product codes of

these are Genesee Polymers (GP)218 and GP226 (Figure 1.3A), as well as a silicone carbinol,


GP507. Interestingly, these species were not found in the original SGS analysed (Sanchez et

al., 2003).

A

Si

Me

Me

Me

O Si

Me

Me

O Si

MeO Si

Me

Me

MeCH2

CH2

CH2

(OCH2CH2)

x

y

ORn

B

Figure 1.3 ‐ Properties of GP226. (A) Chemical structure of GP226 showing the rake structure comprised of a silicone backbone and PEG sidechains. (B) Molecular weight distribution of crude GP226 obtained by PSEC and the chromatographs of four of the separate fractions. Note that fractions III and IV were further separated into two fractions (Radi, 2010).

Analyses of collagen synthesis, as measured by [3H]‐proline incorporation, and total

collagen, as determined by Sirius Red staining, revealed that only GP226 decreased

collagen synthesis in HSF, while both GP226 and GP218 reduced the total amount of

collagen present (McGrath, 2006). It is hypothesised that the combination of hydrophobic

and hydrophilic domains, being the PDMS and PEG structures incorporated respectively,

contributes to the action of these silicones. Furthermore, the low molecular weight of the

species is also thought to be another factor adding to their novel abilities (Hirshowitz et al.,

1993). Taken together, these data has led us to propose that once penetrating the stratum

corneum, these silicones are able to interact with dermal fibroblast cells and affect key

C h a p t e r 1 . 0 | 19

cellular mechanisms pertinent to scarring, such as collagen formation. However, there

were several limitations to these early studies, including the fact that only hypertrophic

skin‐derived cells were investigated and commercial GP226 samples used, in which purity

and identity was never explored. This is of importance as the use of poorly characterised

samples with unknown structure‐property characteristics can be the source of erroneous

and often conflicting results due to sample inconsistencies (Moss et al., 2002). Moreover,

as is illustrated in Figure 1.3B, analysis via preparative size exclusion chromatography

(PSEC) illustrated that GP226 was a crude polymer mixture containing different fractions of

varying molecular weights (Radi, 2010). Of note, each fraction represented 40% (I), 35% (II),

9% (III), 5% (IV) and 8% (V) of the total crude GP226 respectively (Gardoni, M. pers. comm.).

In view of this, the potential of each fraction for scar remediation was assessed in a range

of assays (2007). This included characterising the effect of the fractions on both HSF and

keloid scar‐derived human skin fibroblasts (KF), as well as non‐scar fibroblasts derived from

the same patients (nHSF and nKF respectively). Investigating cells derived from keloid scars

was particularly significant as GP226 had only ever previously been tested on cells derived

from hypertrophic scars. It was revealed that fraction III, further resolved into fractions III

and IV, was the most active fraction of GP226, demonstrating dose‐dependent effects

similar to the crude silicone mixture in cell viability, collagen synthesis and total collagen

assays. In particular, GP226 was found to reduce the total number of viable cells present,

thereby reducing collagen production and total collagen. It was also demonstrated that the

effect of GP226 was similar in both HSF and KF. Moreover, additional studies undertaken

by our laboratory have demonstrated fractions III and IV of GP226 to be capable of

penetrating the stratum corneum (Gardoni, 2007).

1.6.2 Silicone and Apoptosis?

Our data indicating significant decreases in cell viability following exposure to GP226 and

its fractions make sense in many respects as tissue homeostasis, which is maintained

through a balance between cell proliferation and death, is a key element in hypertrophic

and keloid scar formation (Bellemare et al., 2005; Moulin et al., 2004). Interestingly,

decreased apoptosis of fibroblasts has been reported as a major factor in the

etiopathogenesis of hypertrophic and keloid scars (Appleton et al., 1996; Saray and Gulec,

2005; Sayah et al., 1999). Thus, our preliminary results demonstrate that the development

of a therapy containing active fractions (eg. III and IV) of GP226 has the potential to remove


the tissue bulk associated with abnormal scars and eliminate excess collagen present by

removing the cells that produce it.

Interestingly, the role of silicone on dermal fibroblast apoptosis has not yet been

investigated. It is evident that the cell viability assays performed in our laboratory only

determined the viability of dermal fibroblasts but not the mode of death. Therefore,

further investigations into this issue are pertinent to determine whether GP226, and

particularly the active fractions (eg. III and IV), cause death of dermal fibroblasts by

apoptosis or necrosis. It has also become evident through the literature that a treatment

time of 48 hours with silicone, as was performed in our studies, may not be enough.

Hanasono et al. (2004) found many differences in cell growth curves between days 2 and 5

with application of silicone to dermal fibroblasts. From this, it has become clear that our

cells treated with silicone need to be analysed at different time points of up to one or two

weeks. This is another knowledge gap that is required to be investigated through this

doctoral degree.

1.6.3 In Vitro to In Vivo

Clearly the complete data were generated in vitro, which may not truly represent the in

vivo situation. Firstly, only fibroblasts were investigated in the preliminary analyses. As

there is a close paracrine relationship between the dermis and epidermis within the skin,

responses of keratinocytes as well as fibroblasts to GP226 need to be assessed (Werner et

al., 2007). Secondly, differences in fibroblast behaviour following exposure to silicone in

vitro have been reported by McCauley et al. (1990) and Sank et al. (1993), but Chang et al.

(1995) claimed that the same outcomes would not likely be observed in vivo. This was

based on the presumption that silicone oil has difficulty penetrating the stratum corneum

and gaining access to dermal fibroblasts when applied topically to skin surfaces because it

consists of hydrophobic inert macromolecules (Chang et al., 1995) which are based

primarily on lower molecular weight PDMS oligomers (Van den Kerckhove et al., 2001). In

spite of this, our laboratory (Sanchez et al., 2005) and others (Shigeki et al., 1999) have

successfully demonstrated that the combination of hydrophilic and hydrophobic qualities,

as well as the low molecular weight of various silicone species, like GP226, can facilitate

this permeation. Interestingly, of all fractions investigated within the preliminary studies,

the most active, III and IV, were those of low molecular weight. While fractions III and IV

were not the smallest fractions tested, the results obtained indicate that they are the most

C h a p t e r 1 . 0 | 21

effective and thereby have the greatest potential to modulate dermal fibroblasts activity in

vivo. Further investigation of active fractions (e.g. III and IV) is therefore clearly warranted.

1.7 PROJECT HYPOTHESIS AND AIMS

Although SGS are widely used, there clearly exists a need for further research within the

area. Thus, the research conducted for this doctoral degree was designed to investigate the

knowledge gaps present in current literature and extend on our preliminary findings.

1.7.1 Hypothesis

The underlying hypothesis of this project was that low molecular weight amphiphilic

silicones, such as GP226, fraction III and IV, decrease cell viability by regulating apoptosis in

dermal fibroblasts within hypertrophic and keloid scars.

1.7.2 Aims

A summary of the following aims and experimental approach are described in Table 1.1.

Aim 1 Aim 2 Aim 3

Cell Types HSF, nHSF, KF, nKF, HK HSF, nHSF, HK HSF, nHSF

Treatments GP226, Fractions I, II, III, IV and

IV

GP226 Synthetic silicones PDMS7‐g‐PEG7

GP226 Fraction IV

PDMS7‐g‐PEG7 Experiments

Cell Viability Cell Proliferation Cell Morphology

Induction of Apoptosis

Cell Proliferation Cell Morphology

Induction of Apoptosis

Analysis of Gene Expression

Replicates Each assay was repeated with cells from three different patients.

Each variable was tested in triplicate

Each assay was repeated with cells from three different patients.


Each assay was repeated with cells from three different patients.


Table 1.1 – Summary of aims and experimental approach

Aim 1 ‐ To determine the function of commercially available low molecular weight silicones

by investigating their effect on viability, proliferation, apoptosis and necrosis in

hypertrophic scar‐derived human skin fibroblasts (HSF), keloid scar‐derived human skin

fibroblasts (KF), and matched normal human skin fibroblasts derived from the same


patients (nHSF and nKF, respectively) as well as keratinocytes isolated from normal human

skin (HK).

Aim 2 ‐ To investigate and determine the function of fully synthetic low molecular weight

silicones by investigating their effect on proliferation and apoptosis of fibroblasts derived

from hypertrophic scars (HSF) and the unaffected skin of the same patients (nHSF), as well

as keratinocytes isolated from normal skin (HK).

Aim 3 ‐ To investigate the mechanisms underpinning the functional responses induced by

the low molecular weight silicones.

C h a p t e r 2 . 0 | 23

CHAPTER 2.0

MATERIALS AND METHODS

2.1 INTRODUCTION

All materials and methods referred to in the following results chapters are outlined in this

methods chapter. Any variations to these protocols are described in the individual results

chapters. All general reagents were supplied from various commercial suppliers and were

of the highest laboratory grade. Suppliers of method specific reagents are detailed in each

section.

The majority of this experimental work, including all GP226 separation, cell culture, silicone

treatment, cell viability, cell proliferation, silicone and protein interaction, cell morphology,

apoptosis detection, RNA extraction, cDNA synthesis, primer design and qRT‐PCR

experiments, was performed by the author. PDMS‐PEG oligomers were synthesized by

Daniel Keddie, Marilla Dickfos, Brooke Farrugia and Tim Dargaville, Queensland University

of Technology (QUT, Kelvin Grove, QLD, Aus), the HumanHT‐12 v3 Expression BeadChip

microarray was performed by Katie Nones at the Special Research Facility Microarray

Service within the Institute of Molecular Biosciences, University of Queensland (St Lucia,

QLD, Australia) and microarray data analysed by Daniel Haustead, University of Western

Australia (Perth, WA, Aus), and Jacqui McGovern, QUT.

2.2 SILICONE PREPARATION

2.2.1 Fractionation of GP226

An analogue of the silicone oligomers present in Silicone Gel Sheets (SGS), the commercial

amphiphilic polydimethylsiloxane, GP226 (Genesee Polymers, Burton, MI, USA), and its

purified fractions, forms the basis of research in this doctoral degree. Preparative scale Size

Exclusion Chromatography (PSEC) and High Performance Liquid Chromatography (HPLC),

using an Agilent 1200 preparative High Performance Liquid Chromatograph (Agilent

Technologies, Santa Clara, CA, USA) with a SecurityGuard Preparative Cartridge C18

15x21.2 mm (Phenomenex, Lane Cove, NSW, Aus), a SecurityGuard Preparative Cartridge

Holder Kit 21.2 mm (Phenomenex) and a Luna 5u C18(2) 100A AXIA packed 100 x 21.2 mm

column (Phenomenex), were used to separate and obtain pure samples of fractions from

GP226. Tetrahydrofuran (THF; Sigma‐Aldrich, Castle Hill, NSW, Aus) was used at 5 mL/min

M a t e r i a l s a n d M e t h o d s

as a solvent for GP226 (700μL injection volume) during the separation process. Following

separation, excess THF was evaporated via rotary evaporation (Rotavapor, Buchi, Flawil, St.

Gall, Switzerland). The crude, unpurified GP226 and negative controls were prepared at

concentrations ranging from 0.01% to 1% in cell culture medium and used as treatments

applied to cultured cells. In experiments with fractionated GP226, amounts of each fraction

were used at concentrations equivalent to what were originally present in the

unfractionated GP226 mixture. Hence, proportionate dilutions as a w/w % of the active

fractions, compared to the unpurified GP226, were prepared as treatments (Table 2.1). A

silicone polymer previously shown as a major non‐migratory component of SGS and the

backbone of GP226, PDMS (Sigma‐Aldrich), together with PEG (2 kDa; Sigma‐Aldrich), a

representative of the sidearms of GP226, were used as negative controls in all experiments

(Sanchez et al., 2005).

Treatment

Density Proportion 0.01%

(ng/mL)

0.03%

(ng/mL)

0.1%

(ng/mL)

0.3%

(ng/mL)

1%

(ng/mL)

PEG 1.20 100% 105 360 1200 3600 12000

PDMS 0.98 100% 100 295 1000 2950 10000

GP226 1.04 100% 105 310 1050 3100 10500

Fraction I * 25% 26.3 77.5 263 775 2630

Fraction II * 38% 39.8 117 398 1170 3980

Fraction III * 10% 10.5 31 105 310 1050

Fraction IV * 3% 3.5 10 35 100 350

Fraction V * 24% 25 75 250 750 2500

Table 2.1 – Summary of the commercially available polymers used as treatments. Properties (where known) of PEG, PDMS, GP226 and fractions are presented with their tested concentrations. The densities of GP226 fractions (*) are not available but considered to be equivalent to GP226.

2.2.2 Synthesis of PDMS‐PEG oligomers

PDMS‐PEG oligomers were synthesized by Daniel Keddie, Marilla Dickfos, Brooke Farrugia

and Tim Dargaville (QUT) as previously described (Dickfos, 2008; Keddie et al., 2011).

Methods explaining the synthesis of the silicones used are described in Appendix 1. The

synthetic silicones, including PDMS15.2‐PEG8, PDMS10.5‐PEG8, PDMS15.2‐PEG4, and PDMS7‐g‐

PEG7 were prepared at concentrations ranging from 0.03% to 0.3% in cell culture medium

for experiments screening all synthetic silicones. For all other experiments, PDMS7‐g‐PEG7

C h a p t e r 2 . 0 | 25

was prepared at concentrations ranging from 0.0001% to 1.0% in cell culture medium. In

the experiments examining cell proliferation, the PDMS backbone used for synthesis,

PDMS7, was included as a negative control.

2.3 CELL CULTURE

2.3.1 Fibroblast Cell Culture

Hypertrophic scar‐derived human skin fibroblasts (HSF), keloid scar‐derived human skin

fibroblasts (KF), and matched normal human skin fibroblasts derived from the same

patients (nHSF and nKF, respectively) were purchased from Cell Research Corporation

(Singapore). The fibroblasts were isolated using the explant method by Cell Research

Corporation, and cultures contained fibroblasts that originated from both dorsal and

ventral orientations of dermis. To perform biologically relevant replicates, cells from three

patients suffering from hypertrophic scars were purchased (Table 2.2). The cells were

routinely cultured independently in Dulbecco’s Modified Eagle’s Medium (DMEM;

Invitrogen, Mulgrave, VIC, Aus) containing 10% fetal calf serum (FCS; Hyclone, Thermo

Fisher Scientific, Scoresby, VIC, Aus), penicillin (100 U/mL; Invitrogen), streptomycin (100

U/mL; Invitrogen) and L‐Glutamine (2 x 10‐3 M; Invitrogen). Furthermore, the cells were

maintained in T175cm2 cell culture flasks (Nunc, Thermo Fisher Scientific) within a

humidified atmosphere of 5% CO2 at 37°C with media being replaced every 2‐3 days and

used at low passage number (p5‐10). All cultures were passaged every 5‐7 days by 0.05%

Trypsin‐ethylenediaminetetraacetic acid (Trypsin‐EDTA; Invitrogen) detachment.

Patient

Age Sex Scar

Location

Ethnicity

105 28 Male Hand Chinese

106 51 Male Forearm Chinese

107 28 Female Forearm Chinese

Table 2.2 – Patient data obtained from hypertrophic scar donors.

Patient information for the donors of hypertrophic scar‐derived human skin fibroblasts and matched

normal human skin fibroblasts as provided by Cell Research Corporation.

2.3.2 Keratinocyte Cell Culture

Keratinocytes (HK) were isolated as previously described (Dawson et al., 2006) from human

skin of consenting patients undergoing elective abdominoplasty or mammoplasty


procedures. Ethics approval to use this tissue was obtained from the Queensland University

of Technology Human Research Ethics Committee (ID:3673H). An irradiated murine feeder

layer (i3T3; ATCC, Manassas, VA, USA) was used to expand the freshly isolated HK. In brief,

the 3T3 cells were maintained in DMEM supplemented with FCS (5%), penicillin (100

U/mL), streptomycin (100 U/mL) and L‐Glutamine (2 x 10‐3 M) before being irradiated (50

Gy) at the Australian Red Cross Blood Service (Brisbane, QLD, Australia). The i3T3 cells were

then seeded into cell culture flasks (1 x 106 cells/T75cm2 cell culture flask; Nunc) 2 hours

before the HK were added (2 x 106 cells/T75cm2 cell culture flask).

The HK were routinely cultured in Full Green’s Medium, consisting of DMEM and Ham's F12

medium (Invitrogen) in a 3:1 ratio, supplemented with FCS (10%), insulin (1 µg/mL; Sigma‐

Aldrich), human recombinant epidermal growth factor (10 ng/mL; Invitrogen), cholera toxin

(0.1 µg/mL; Sigma‐Aldrich), nonessential amino acids solutions (0.01% (v/v); Invitrogen),

hydrocortisone (0.4 µg/mL; Sigma‐Aldrich), adenine (180 µM; Sigma‐Aldrich), transferrin (5

µg/mL; Sigma‐Aldrich), triiodothyronine (0.2 µM; Sigma‐Aldrich), L‐glutamine (2 mM),

penicillin (100 U/mL) and streptomycin (100 U/mL), and were maintained in the standard

conditions of 37°C and 5% CO2. All cultures were passaged every 5‐7 days by 0.05% Trypsin‐

EDTA detachment.

2.4 FUNCTIONAL ASSAYS

2.4.1 WST‐1 Assay for Determination of Cell Viability

Cell viability was measured using the WST‐1 (Roche Applied Science, Castle Hill, NSW, Aus)

Assay Kit. In brief, WST‐1 is a modified tetrazolium salt (4‐[3‐(4‐Iodophenyl)‐2‐(4‐

nitrophenyl)‐2H‐5‐tetrazolio]‐1,3‐benzene disulfonate), which when in contact with active

coenzymes, such as NADH and NADPH, present on the plasma membrane, has its

tetrazolium ring cleaved to form a water‐soluble formazan product, with a corresponding

increase in absorbance at 440nm (Berridge et al., 1996).

The response of HSF, KF, nHSF, nKF and HK to silicone treatments after different periods of

exposure were evaluated using this method. Following culture, HSF, KF, nHSF, nKF and HK

were seeded into 96 well plates (Nunc) for treatment with silicones. Cultures of HSF, KF,

nHSF and nKF cells were seeded at a density of 3000 cells/200 µL into each well and were

treated with silicone solutions (as described in sections 2.2.1 and 2.2.2) diluted in cell

culture medium (200μL /well) 24 hours later. Cultures of HK cells were seeded (1.2 x 104

C h a p t e r 2 . 0 | 27

cells/well) onto irradiated 3T3 cells (1 x 104 cells/well) in 96 well plates (Nunc) and silicone

solutions (as described in sections 2.2.1 and 2.2.2) diluted in Full Green’s Medium were

added 24 hours later (200 µL/well). Controls included: a blank control, containing medium

only; an untreated control, containing cells treated with medium but no silicone; and

negative controls, including cells treated with the silicone controls described in sections

2.2.1 and 2.2.2. During all incubations, the cells were maintained in the standard conditions

of 37°C and 5% CO2.

Following incubation with silicone treatments for 24, 48, 72 or 168 hours (7 days), cell

culture medium was removed by gently inverting the 96‐well plate, followed by blotting on

paper towels to remove excess medium. In HK cultures, the 3T3 layer was removed by a

quick wash with 0.05% Trypsin‐EDTA and visualization to confirm removal was obtained by

light microscopy (Nikon Eclipse TS100). The working WST‐1 reagent for each 96‐well plate

was prepared by combining 500 µL WST‐1 reagent with 4.5 mL cell culture medium. The

prepared WST‐1/cell culture medium solution was added to each sample well (50µL) before

the plates were stored at 37°C for 1.5 hours incubation. The absorbances of the samples in

the plates were then quantitatively measured using a plate reader (Microplate Manager

Version 5.2; Bio‐Rad Laboratories, Gladesville, NSW, Aus) at 440 nm, using 620 nm as a

reference.

2.4.2 Cyquant Assay for Determination of Cell Proliferation

Cell proliferation was measured using the Cyquant (Invitrogen) Assay Kit. Cyquant directly

measures cellular deoxyribonucleic acid (DNA) content using a fluorescent DNA binding

dye. Because cellular DNA content is highly regulated, it is closely proportional to cell

number (Myers, 1998).

The response of HSF, KF, nHSF, nKF and HK to silicone treatments after different periods of

exposure were evaluated using this method. Following culture, HSF, KF, nHSF, nKF and HK

were seeded into 96 well plates (Nunc) for treatment with silicones. Cultures of HSF, KF,

nHSF and nKF cells were seeded at a density of 3000 cells/200 µL into each well and were

treated with silicone solutions (as described in sections 2.2.1 and 2.2.2) diluted in cell

culture medium (200 μL/well) 24 hours later. Cultures of HK cells were seeded (1.2 x

104cells/well) onto irradiated i3T3 cells (Section 2.3.2; 1 x 104 cells/well) in 96 well plates

and silicone solutions (as described in sections 2.2.1 and 2.2.2) diluted in Full Green’s


Medium were added 24 hours later (200 µL/well). In some experiments, HK were seeded

into the 96 well plates (1.2 x 104 cells/well) without an i3T3 feeder layer. Controls included:

a blank control, containing medium only; an untreated control, containing cells treated

with medium but no silicone; and negative controls, including cells treated with the silicone

controls described previously in sections 2.2.1 and 2.2.2. During all incubations the cells

were maintained in the standard conditions of 37°C and 5% CO2.

Following incubation with silicone treatments for 24, 48, 72 or 168 hours (7 days), cell

culture medium was removed by gently inverting the 96‐well plate, followed by blotting on

paper towels to remove excess medium. In cultures of HK grown with an i3T3 feeder layer,

the i3T3 layer was removed by a quick wash with 0.05% Trypsin‐EDTA and confirmation

that the cells had been removed was confirmed by visualization using light microscopy

(Nikon Eclipse TS100; Nikon, Lidcombe, NSW, Aus). The plates were then sealed in parafilm

and frozen in a ‐80°C freezer overnight. The Cyquant assay was performed on the following

day. For each 96‐well plate tested, the Cyquant solution was prepared by diluting 1 mL of

20 X concentrated cell‐lysis buffer stock solution in 19 mL nuclease‐free distilled water and

adding 50 µL Cyquant stock solution. The frozen 96‐well plates were thawed at room

temperature before 200 µL of Cyquant dye/cell lysis solution was added to each sample

well. The plates were incubated in darkness for 2‐5 minutes at room temperature.

Following this, fluorescence was measured with excitation at 485 nm and emission at 530

nm using the POLARstar OPTIMA Plate Reader (BMG Labtech, Mornington, VIC, Aus) and

v1.30‐0 OPTIMA software.

2.4.3 Analysis of Silicone and Protein Interaction

The morphology of HSF following culture in varying concentrations of FBS, with and

without GP226, was investigated to observe if GP226 was causing an aggregation of

proteins within the cell culture medium. HSF (2x104 cells/well) were seeded into 24 well

plates (Nunc) and cultured in 0%, 5%, 10% or 20% FCS/DMEM. GP226 (0.1%) diluted in the

respective cell culture media was used to treat the cells 24 hours later (1 mL/well).

Following 48 hours treatment with silicone, HSF were stained with Sirius Red (0.1% (w/v)

Direct Red 80 powder; Sigma Aldrich/double distilled water (ddH2O)) for 1.5 hrs before

being washed twice with ddH2O. The cells were stained with Sirius Red for ease of cell

morphology analysis and to localize the collagen content within the cultures. Cell

C h a p t e r 2 . 0 | 29

morphology was documented using digital photography (Nikon Coolpix 4500, Nikon) and

light microscopy (Nikon Eclipse TS100, Nikon).

Aggregation of proteins in cell culture medium was also assessed via gel electrophoresis.

Protein concentrations in DMEM containing 0%, 5%, 10% or 20% FCS, with or without 0.1%

GP226 were determined using the bicinchoninic acid protein assay (Thermo Scientific)

according to the manufacturer’s directions. Equal amounts of protein were then diluted to

7.5 µL in ddH2O and combined with 2.5 µL of 4 x NuPAGE lithium dodecyl sulphate sample

buffer (Invitrogen). Samples were then loaded into 14‐well pre‐cast 4‐12% Bis‐Tris gels

(1mm; Invitrogen), with each gel including one lane of 2 µL protein molecular weight

standard (Precision Plus Protein Standards, Bio‐Rad Laboratories, Hercules, CA, USA).

Electrophoresis was then performed at 200V for 35 minutes using NuPAGE 2‐(N‐

morpholino)ethanesulfonic acid buffer (Invitrogen). Gels were fixed in 2 washes of 10%

ethanol / 30% acetic acid for 15 minutes, each with gentle shaking at room temperature.

The Pierce Silver Stain kit (Thermo Scientific) was then used to stain the proteins present

within the gels, using the protocol supplied by the manufacturer. Following development,

the silver stain reaction was stopped using 10% acetic acid for 10 minutes with gentle

shaking at room temperature. An image of the gels was documented using a flat‐bed

scanner (CanoScan 8600F, Canon, Ohta‐ku, Tokyo, Japan).

2.4.4 Analysis of Cell Morphology via Real‐Time Microscopy

Real‐time microscopy was used to visualize the response of HSF, nHSF and HK to silicone

treatment over 48 hours. In brief, HSF (2 x 104 cells/well) and nHSF (2 x 104 cells/well) were

plated into 24 well plates and treated with 24 hours later GP226 (0.1%), Fraction IV (0.1%),

Tween 20 (0.1%) and PDMS7‐g‐PEG7 (0.01 % and 0.03%) diluted in cell culture medium (1

mL/well) being added 24 hours later. Cultures of HK cells were seeded (6 x 104 cells/well)

onto irradiated 3T3 cells (5 x 104 cells/well) in the 24 well plates and silicone solutions

diluted in Full Green’s Medium were added 24 hours later (1 mL/well). In some

experiments, HK were seeded into the 96 well plates (6 x 104 cells/well) without an i3T3

feeder cell layer. Treatments included GP226, Fraction IV, Tween 20 (a commonly used

surfactant) and PDMS7‐g‐PEG7 at concentrations ranging from 0.01% to 1%. Real‐time

microscopy was then performed with a Leica AF6000 Widefield Microscope (Leica

Microsystems, North Ryde, NSW, Aus) and images were taken every 15 minutes for 48

hours. Still images and movies were prepared using Leica Application Suite software.


2.4.5 Tunel Assay for the Detection of Apoptosis

The induction of apoptosis following exposure of HSF, nHSF, and HK to the silicone

treatments was measured using the Tunel assay (Roche Applied Science). In brief,

apoptosis causes the cleavage of genomic DNA into double stranded, low molecular weight

DNA fragments, as well as causes single strand breaks in high molecular weight DNA. These

DNA breaks are able to be fluorescently‐labelled using reactions between terminal

deoxynucleotidyl transferase, the nicks in the DNA and fluorescein‐labeled nucleotides,

which taken together is known as Tunel technology (Gavrieli et al., 1992).

Cell cultures of HSF and nHSF (4 x 103 cells/400 µL) were prepared in glass 8‐well chamber

slides (BD Biosciences, North Ryde, NSW, Aus) and treated with GP226, Fraction III, Fraction

IV, Tween 20, PEG and PDMS7‐g‐PEG7 diluted in cell culture medium 24 hours later (400

µL/well). Concentrations tested included 0.1% for GP226, Fraction III, Fraction IV, Tween 20

and PEG, as well as a range of concentrations (0.003% to 0.03%) for PDMS7‐g‐PEG7.

Cultures of HK cells were seeded (2.5 x 104 cells/400 µL) onto irradiated 3T3 cells (2 x 104

cells/400 µL) in glass 8‐well chamber slides and silicone solutions, diluted in Full Green’s

Medium, were added 24 hours later (400 µL/well). An untreated control, containing cells

treated with media only was included.

After 48 hrs incubation the cells were fixed in 4% paraformaldehyde (200 µL; Chem Supply,

Gillman, SA, Aus) for 15 minutes, washed once in 200 µL phosphate‐buffered saline (PBS;

Invitrogen) and were permeabilised in 0.2% Triton X‐100 (Sigma‐Aldrich)/PBS for 15

minutes. Following further washes in PBS, the cells were incubated with 50 µL Tunel

reaction mixture for 60 minutes at 37 °C. A positive control, consisting of untreated cells

incubated with recombinant Deoxyribonuclease I (DNase I; Sigma‐Aldrich) for 10 minutes at

room temperature following permeabilisation, as well as a negative control, containing

untreated cells incubated with labelling solution only without Tunel enzyme, were also

included. After incubation with the Tunel reaction mixture, the samples were washed with

PBS and mounted in SlowFade Gold antifade reagent (Invitrogen) with added 4',6‐

diamidino‐2‐phenylindole (DAPI), a nuclear stain (Invitrogen), and were then sealed with

glass coverslips (Labtek, Brendale, QLD, Aus) and clear nail polish. A Nikon Eclipse TE2000‐U

fluorescence microscope (Nikon) was used to visualise and photograph the cells.

C h a p t e r 2 . 0 | 31

2.5 GENE ANALYSIS

2.5.1 Extraction of RNA from Silicone‐Treated Fibroblasts

Total RNA was extracted from the treated cell cultures using Qiagen RNeasy Mini kit

(Qiagen, Doncaster, VIC, Aus). Cell cultures of HSF and nHSF were prepared in T25 cm2 cell

culture flasks (Nunc) at a density of 2 x 105 cells/5 mL cell culture medium. Following 24

hours incubation in normal tissue culture conditions, cell culture medium containing GP226

(0.1%), Fraction IV (0.1%) and PDMS7‐g‐PEG7 (0.01% and 0.03%) were added. Cell cultures

treated with GP226 and Fraction IV were incubated for 48 hours while cell cultures with

PDMS7‐g‐PEG7 were incubated for either 6 hours or 48 hours. An untreated control, a cell

culture treated with cell culture medium containing no silicone for 6 or 48 hours was also

prepared. Following treatment, the cell culture medium was removed by aspiration and the

cells were lysed directly in the T25 cm2 cell culture flask. To lyse the cells, 600 µL of Buffer

RLT Plus, combined with 6 µL β‐mercaptoethanol (Sigma‐Aldrich), was added into the cell

culture flasks and the cells were dislodged from the surface with a cell scraper. Cell lysis

solutions were then stored at ‐80°C.

To extract RNA using the Qiagen RNeasy Mini kit, the cell lysates were thawed at room

temperature, transferred to a gDNA eliminator spin column placed in a 2 mL collection tube

and centrifuged for 30 seconds at ≥8000 x g. The column was discarded and 600 µL ethanol

(70%) added. The samples (600 µL at a time) were then transferred to an RNeasy spin

column placed in a 2ml collection tube and centrifuged for 15 seconds at ≥ 8000 x g, with

the flow‐through being discarded after each centrifugation. The RNeasy columns were then

washed once with 700 µL Buffer RW1 and twice with 500 µL Buffer RPE, each with

centrifugations at ≥8000 x g and the flow‐through being discarded. The RNeasy columns

were then placed in new 2 mL collection tubes and centrifuged at full speed for 2 minutes

to remove any excess washing buffers. Following this, the RNeasy columns were placed

into new 1.5 mL collection tubes and RNA eluted with 30 µL RNase‐free water in addition to

centrifugation at ≥ 8000 x g for 1 minute. RNA was quantified at 260nm using a Nanodrop

spectrophotometer (Thermo Fisher Scientific). RNA samples were stored at ‐80°C.

2.5.2 HumanHT‐12 v3 Expression BeadChip Microarray

The HumanHT‐12 v3 Expression BeadChip (Illumina, San Diego, CA, USA) microarray was

used to determine differentially expressed genes in cultures of HSF and nHSF following

treatment with and without 0.1% GP226 and 0.1% Fraction IV. The HumanHT‐12 Expression


BeadChip was chosen as it targets more than 25 000 annotated genes with more than 48

000 probes and provides genome‐wide transcriptional coverage of well characterized

genes, gene candidates and splice variants. More specifically, the probes were selected

from reputable RefSeq (Build 36.2, Ref 22) and UniGene (Build 199) databases.

Furthermore, the HumanHT‐12 BeadChip encompassed a higher throughput system,

containing the same probes present on the larger HumanWG‐6 BeadChip, but allowing 12

samples to be processed simultaneously. The HumanHT‐12 v3 Expression BeadChip

microarray was performed with total RNA extracted from treated and untreated cell

cultures by Katie Nones at the Special Research Facility Microarray Service within the

Institute of Molecular Biosciences, University of Queensland (St Lucia, QLD, Australia).

2.5.3 Microarray Data Analysis using GeneSpring GX 10.0

Following microarray, the data were returned for analysis, and kindly analysed by Daniel

Haustead, University of Western Australia (Perth, WA, Aus), and Jacqui McGovern, QUT.

Biological replicates of samples were unable to be performed within the microarray,

meaning that standard microarray analysis could not be carried out. In brief, data analysis

was performed in GeneSpring GX 10.0 (Agilent Technologies) and quantile normalization

used. Data were pre‐processed using the baseline to median baseline transformation tool

with standard filtering options. A flag cell check of each data point was then performed,

with data signals being labelled ‘present’, ‘marginal’ and ‘absent’ and all ‘absent’ signals

being removed. Data points with ‘present’ or ‘marginal’ tags were used for final analysis

between samples of treated and untreated cell cultures. Further analysis or false discovery

testing was not performed as no biological replicates were included in the microarray.

Changes occurring between sample pairs were manually investigated by fold‐change on a

gene‐to‐gene basis. Specifically, only genes with fold‐change values greater than ± 2 were

included. Resulting lists of genes and their fold‐change values between sample pairs

comparing treated and untreated cell culture samples were then used to select genes for

validation via quantitative reverse transcription‐polymerase chain reactions (qRT‐PCR).

Results were manually sorted and genes relating to apoptosis and collagen production

pathways were selected for validation.

C h a p t e r 2 . 0 | 33

2.5.4 Gene Ontology and Canonical Pathway Analysis using Ingenuity Pathway Analysis

tools

Gene ontology, canonical pathway and functional network analyses were undertaken using

Ingenuity Pathway Analysis (IPA) tools (Ingenuity Systems, www.ingenuity.com; Based in

Redwood City, CA, USA). IPA is a database consisting of gene‐based results and includes

functions and interactions between genes/gene products mined from the peer‐reviewed

literature. Data sets containing gene identifiers and corresponding expression values of

probe sets determined to be significantly expressed by at least ± 2‐fold in response to

GP226 and Fraction IV compared to untreated samples, were uploaded into the

application. Genes which map to the IPA knowledge base, called focus genes, were then

used to generate biological networks and identify biological functions that were most

significant to each uploaded data set. IPA computed a score for each network according to

the fit within the original set of significant genes submitted. The resultant score reflected

the algorithm of probability (P) that indicated the likelihood of the focus genes in a network

being found together by random chance. The functional analysis was also used to identify

genes that were most significantly involved in the affected biological functions for each

data set.

2.5.5 Apoptosis Superarrays

In addition to the microarrays performed above, quantitative reverse transcription‐

polymerase chain reaction (qRT‐PCR) superarrays (SA Biosciences, Qiagen) were also

performed on HSF and nHSF cultures exposed to either 0.01% or 0.03% PDMS7‐g‐PEG7.

These arrays provide a sensitive gene expression profiling technique, which allows for

panels of genes related to different biological pathways to be screened for deregulation.

More specifically, arrays analysing biological processes relevant to this project, including

the human apoptosis pathway, were utilized. Furthermore, the 384 well format of the

superarray was chosen, which meant that more data could be obtained from one plate.

Firstly, total RNA of HSF and nHSF cell cultures treated with and without PDMS7‐g‐PEG7

(0.01% and 0.03%) was extracted using the protocol described in section 2.5.1 and first

strand complementary DNA (cDNA) subsequently synthesised using the SA Biosciences First

Strand Synthesis Kit (SA Biosciences). In brief, genomic DNA was removed by combining 1

μg of total RNA from each sample with 2 μL 5X gDNA Elimination buffer and diluted to 10

μL with nuclease‐free H2O. Each RNA sample was then incubated at 42°C for 5 minutes


before being chilled on ice for at least 1 minute. First strand synthesis of total RNA was

then performed by adding 4 μL 5X RT Buffer 3, 1 μL Primer and External Control Mix, 2 μL

RT Enzyme Mix 3 and 3 μL nuclease‐free H2O to each sample. When large numbers of

samples needed to be prepared at once, a RT mastermix was prepared with the described

reagents and once combined, 10 μL added to each total RNA sample. The volume of each

total RNA sample was then 20 μL. The RNA samples were incubated at 42 °C for 15 minutes

and the reaction stopped by heating at 95 °C for 5 minutes. Following this, each 20 μL cDNA

synthesis reaction was diluted with 91 μL of nuclease‐free H2O.

Following the synthesis of cDNA, the arrays were performed as per the manufacturer’s

instructions. Briefly, an experimental master mix was prepared for each sample in which

102 μL of the diluted first strand cDNA synthesis reaction was combined with 550 μL of 2 X

SABiosciences RT2 qPCR Mastermix (SYBR Green; SA Biosciences) and 448 μL of nuclease‐

free H2O. Following this, the samples were pipetted into a 384‐well format apoptosis PCR

array (10 μL/well) using 384 EZLoad Covers (SA Biosciences). Using the 384 array, 4 samples

were able to load into one array, though in these experiments, 2 samples (untreated vs

treated) were loaded in duplicate.

The array process was completed in an ABI 7900HT Thermal Cycler (Applied Biosystems,

Life Technologies, Mulgrave, VIC, Aus). The SYBR Green Dye detection system was used

with the following temperature cycling conditions: 94°C for an initial 10 minutes, then 40

cycles of 94°C for 15 seconds and 60°C for 60 seconds. Data generated were analysed using

data analysis provided by SA Biosciences, a program designed to analyse qRT‐PCR data

obtained through superarrays. During these analyses, automatic baseline and threshold

cycles were used. Resulting lists of genes and their fold‐change values between treated and

untreated cell culture samples were then used to select genes for validation via qRT‐PCR.

The results were manually sorted and genes with fold‐change values of greater than ± 1.8

in three or more comparisons were selected for validation. Statistical examination was

unable to be performed as biological replicates were not included in the superarrays.

2.6 CONFIRMATION OF DIFFERENTIAL GENE EXPRESSION USING QUANTITATIVE RT‐PCR

2.6.1 Standard PCR Conditions

PCR reactions were performed using the Platinum Taq PCR kit (Invitrogen). The standard

reaction conditions included: 1 U Platinum Taq, 1.5 mM MgCl2, 0.1 mM each of

C h a p t e r 2 . 0 | 35

deoxyribonucleotide triphosphate (dNTP), 1.25 μM forward and reverse primers and 1 μL

of cDNA template in a total volume of 20 μL. PCR reactions were then run on a MJ Research

Thermocycler (Geneworks, Hindmarsh, SA, Aus) with the following temperature cycling

conditions: 94°C for an initial 5 minutes, then 35 cycles of 94°C for 45 seconds, 60°C for 60

seconds and 72°C for 90 seconds, followed by a final stage of 72°C for 10 minutes. PCR

amplicon size was then confirmed using agarose gel electrophoresis (1.5% agarose) and

ethidium bromide (Bio‐Rad Laboratories)/UV visualization (G Box; Syngene, Frederick, MD,

USA) using a 0.07 ‐ 12.2 kbp molecular weight marker (Roche Applied Science) as a marker

for size.

2.6.2 Primer Design

Primers for qRT‐PCR were designed using Primer‐basic local alignment software (Primer‐

BLAST) software, a web‐based primer designing tool available through the National Center

for Biotechnology Information (www.ncbi.nlm.nih.gov/tools.primer‐blast/) The settings

that were used to define acceptable primer sets included: minimum primer melting

temperature (Tm) of 57°C, maximum of 63°C and optimal Tm of 60°C, with no more than

3°C difference in Tm between primers; a preference for primers to span an exon‐exon

junction and amplicon length between 70 and 150 base pairs. Once designed, PCR primers

(25 nmoles; desalted purity) were purchased from Invitrogen. Table 2.3 depicts the primers

that were designed using this process.


Gene Symbol Sequence – 5’ to 3’ GenBank

Accession #

Amplicon

Position

αSMA F‐ CTGCTGAGCGTGAGATTGTC

R‐ CTCAAGGGAGGATGAGGATG

NM_001613.2 10‐113

AIFM2 F‐ CCTACCGCAAAGCGTTTGAGAGCA

R‐ TGGCGTAGACGTTGCTGTGGC

NM_032797.4 966‐1061

APAF1 F‐ CCTGTTGTCTCTTCTTCCAGTGTAAGG

R‐ AAACAACTGGCCTCTGTGGTACTCCA

NM_001160.2 851‐920

BIK F‐ GCATGGAGGGCAGTGACGCA

R‐ ACCTCGGAGAGCTGGGCCAG

NM_001197.3 209‐308

CD70 F‐ TGGTCGCGGGCTTGGTGATC

R‐ GGTCCTGCTGAGGTCCTGTGTGA

NM_001252.3 227‐361

CFLAR F‐ CAGGACGAACTCCCCCACT

R‐ CAGGACGAACTCCCCCACT

NM_003879 9‐108

CIDEA F‐ ACAGACCTCCAGGCCCGCTAG

R‐ AAATGTCAGGGGCCTGATGAGGG

NM_001279.3 33‐116

COL1A1 F‐ ACGAAGACATCCCACCAATC

R‐ AGATCACGTCATCGCACAAC

NM_000088.3 11‐139

COL3A1 F‐ GCCTCCCGGAAGTCAAGGAGAAAG

R‐ CTTTAGGACCGGGGAAGCCCATG

NM_000090.3 1773‐1870

DAPK1 F‐ GACATCGTGGAGTGTCTGGCCG

R‐ GGGCAATGTGTCCGTCCTTGTCG

NM_004938.2 1948‐2017

FAS F‐ CCTCCTACCTCTGGTTCTTACG

R‐ CAGTCTTCCTCAATTCCAATCC

NM_000043 4‐107

HRK F‐ TACTGGCCTTGGCTGTGCGC

R‐ CACCAACCTGTTGCTCGCTCC

NM_003806.1 322‐459

IL8 F‐TGTGGAGAAGTTTTTGAAGAGGGCTG

R‐ CACTGGCATCTTCACTGATTCTTGGA

NM_000584.2 365‐448

IGF2R F‐AGCTCAAGAATTGGAAGCCAGCAAGG

R‐ GCGGTACCTTTGGCAGAGAGAGG

NM_000876.2 3132‐3246

MCL1 F‐ ACGAGACGGCCTTCCAAGGCAT

R‐ TCCTGCCCCAGTTTGTTACGCC

NM_021960.3 820‐920

NSMAF F‐ GCTGCAGCTCTACTCCAAGGAGA

R‐ TGATTTTCCTTTCATGGTGACTGCCC

NM_003580.3 250‐371

SMAD7 F‐ ACCAACTGCAGACTGTCCAGATGCT

R‐ TCCCAGTATGCCACCACGCAC

NM_005904.3 947‐1090

TGFβ1

F‐ TGTCACCGGAGTTGTGCGGC

R‐ CGGCCGGTAGTGAACCCGTTG

NM_000660.4 1479‐1610

TNF F‐ TCTGGCCCAGGCAGTCAGATC

R‐ TTGGCCAGGAGGGCATTGGC

NM_000594.2 385‐513

TNFRSF9 F‐ AGAGCCAGGACACTCTCCGCAG

R‐ CGGAGCGTGAGGAAGAACAGCA

NM_001561.4 665‐754

TNFRSF10B F‐ TCCACCTGGACACCATATCTC

R‐ AGCAGAAAAGGAGGTCATTCC

NM_003842 6‐103

TRAF2 F‐ ACCGTTGGGGCTTTGTTC

R‐ CCAGGAGGGTCTTGGAGAAG

NM_021138 9‐117

TRAF3

18S‐ rRNA

F‐ CGCGAGAACTCCTCTTTCC

R‐ CGGTCAGTGTGCAGCTTTAG

F‐TTCGGAACTGAGGCCATGAT

R‐ CGAACCTCCGACTTTCGTTC

NM_003300

NR_003286.2

12‐108

898‐1048

Table 2.3 ‐ Primers used for qRT‐PCR

C h a p t e r 2 . 0 | 37

2.6.3 Reverse Transcription (RT) for qRT‐PCR

First strand cDNA synthesis was performed using SuperscriptTM III Reverse Transcriptase

(Invitrogen). Total RNA (1 μg) of HSF and nHSF untreated or treated with GP226 (0.1%),

Fraction IV (0.1%) and PDMS7‐g‐PEG7 (0.03%) was combined with 250 ng of random primers

(Invitrogen) and 0.5 mM dNTP mix (Invitrogen) and diluted to 13 μL with nuclease‐free

water. The samples were incubated at 65°C for 5 minutes and then cooled rapidly on ice for

at least 1 minute. First strand synthesis was then performed by adding 200 U SuperscriptTM

III Reverse Transcriptase, 5 mM dithiothreitol (DTT; Invitrogen), 1 x first strand buffer

(Invitrogen) and 40 U RNaseOUT Recombinant RNase Inhibitor (Invitrogen) to each RNA

sample to a final volume of 20 μL. Each reaction was then incubated at 25°C for 5 minutes,

50°C for 60 minutes and then 70°C for 15 minutes to inactivate the Superscript III enzyme.

The resulting cDNA samples were either stored at ‐80°C or quantified by

spectrophotometry (Thermo Fisher Scientific) and diluted to 20 ng/μL with nuclease‐free

water for qRT‐PCR analysis.

2.6.4 PCR and Amplicon purification

PCR was performed as described previously (Section 2.6.1) using Platinum Taq (Invitrogen)

and sequence specific primers (Table 2.3) for each transcript to be analysed by qRT‐PCR.

PCR amplicons were electrophoresed on 1.5% agarose gels (Sigma‐Aldrich) with ethidium

bromide and visualized using UV illumination (G Box). The resulting band of interest was

excised from the agarose gel using a sterile scalpel blade (Labtek). PCR amplicons were

then purified from the agarose using the MinElute gel extraction kit (Qiagen) according to

the manufacturers’ instructions. PCR products were quantified by spectrophotometry

(Nanodrop spectrophotometer) at 260 nm and the yields converted to absolute cDNA

transcript copy numbers per μL of sample based on 1 DNA bp having a molecular mass of

660 grams/mole.

2.6.5 qRT‐PCR

Microarray and superarray data were validated by using qRT‐PCR to measure absolute

expression levels of the selected genes of interest in cDNA samples of HSF and nHSF

untreated or treated with GP226 (0.1%), Fraction IV (0.1%) and PDMS7‐g‐PEG7 (0.03%). The

primers used in qRT‐PCR were designed using Primer‐BLAST

(www.ncbi.nlm.nih.gov/tools/primer‐blast/), including the 18S ribosomal RNA (rRNA)

normalizing control, and are outlined in Table 2.2. Standard curves, generated by 10‐fold


serial dilutions of purified PCR target amplicons and covering at least 7 logs of amplicon

copy number, were used to achieve absolute quantitation. The reactions were performed

in 20 μL volumes in 96‐well format in triplicate using SYBR green and an ABI Prism 7000

Sequence Detection System (Applied Biosystems). Reactions contained 1 x SYBR‐green PCR

mastermix (Applied Biosystems), 1 μM of each forward and reverse primers and 100 ng of

cDNA. PCR amplification followed a two step cycling protocol with an initial denaturation

for 10 min at 95°C, with 40 cycles of 95°C for 15 seconds and 60°C for 1 minute. All

reactions included a post‐amplification melt curve analysis to determine the melting

temperature (Tm) of the amplified PCR product, indicating amplification of the correct

sequence. Analysis was performed using the ABI Sequence Detection System software

version 1.2.3 (Applied Biosystems) using the automatic options for baseline and threshold

values. The software determines the PCR cycle at which each reaction reached its log‐linear

phase and is directly proportional to the amount of starting cDNA transcript. The cDNA

copy number for each reaction was then calculated by direct comparison to the known

standards for each gene, which are run concurrently. Expression of target genes for each

treated sample was normalized to 18S rRNA and compared to normalized untreated

samples.

2.7 STATISTICAL ANALYSIS

To obtain replicate biological samples, the assays were repeated three times using HSF and

nHSF cells from three different patients. Within each experiment, each variable was tested

in triplicate. Data were expressed as a mean ± standard error of the mean (SEM) and each

treatment group declared significantly different to the untreated group with analysis of

variance followed by a pairwise t‐test post hoc. P values < 0.05 were considered significant.

C h a p t e r 3 . 0 | 39

CHAPTER 3.0

FUNCTIONAL ANALYSIS OF THE EFFECTS OF GP226 AND ITS

FRACTIONS ON DERMAL FIBROBLASTS

3.1 INTRODUCTION

Despite considerable research, the mechanism of SGS action has not yet been completely

elucidated (Borgognoni, 2002). Research undertaken by our laboratory has re‐investigated

the chemical role of SGS on scarring. It has been demonstrated that small amounts of

amphiphilic, linear, oligomeric silicones with low molecular weight have the ability to

migrate from commercially available SGS into the human stratum corneum (Sanchez et al.,

2005). In a preliminary study Sanchez (2006) screened a large number of commercial

amphiphilic oligomeric silicones to investigate their effect in down‐regulating collagen

production and found that the silicone GP226 appeared to be the most effective. Further, it

was also determined that there is a fraction of GP226 which contained the most active

component(s) (Charters, 2007). Fraction IV, defined as the fourth peak eluted in PSEC and

representing 8% of the total GP226 mixture, demonstrated dose‐dependent effects similar

to the crude GP226 silicone mixture in terms of effects on cell viability, collagen synthesis

and total collagen assays conducted in HSF, nHSF, KF and nKF cells. Thus, GP226, including

its active fraction, reduced the total number of viable cells present, resulting in reduced

collagen production and total collagen. It was also demonstrated that the effect of GP226

was similar in fibroblasts isolated from both hypertrophic (HSF) and keloid (KF) scars

(Charters, 2007).

While fibroblasts were investigated in these earlier studies, the other prominent cell type in

skin, keratinocytes, was not studied. Since there is: 1) a close paracrine relationship

between the dermis and epidermis within skin; and 2) wound healing involves restoring

barrier function through the formation of a functional epidermis (Martin, 1997), responses

of keratinocytes as well as fibroblasts to GP226 required assessment. Furthermore,

although significant decreases in cell viability following exposure of fibroblasts to GP226 for

48 hours were demonstrated, Hanasono et al. (2004) reported that differences in cell

growth curves of dermal fibroblasts occurred between days 2 and 5 of treatment with

silicone. This demonstrated a need to look at the effect of the silicones at multiple time

points, rather than just at 48 hours. Finally, the previous analyses only measured cell

F u n c t i o n a l A n a l y s i s o f G P 2 2 6

viability, a method that doesn’t directly measure cell numbers but instead measures the

amount of metabolically active cells present. Thus, further investigation of GP226 and

Fraction IV is required. In particular, examination of GP226 and its fractions not only on

fibroblasts derived from hypertrophic scars but also from the normal skin of the same

patients is necessary. In addition, investigations to elucidate the optimal treatment time

and concentration of GP226 and Fraction IV was also required to facilitate future cell

culture studies directed at examining the mode of action of these silicones.

3.2 EXPERIMENTAL PROCEDURES

Full specifications of materials and methods used in experimental procedures for this

chapter are described in Chapter 2. The following is a summary of the procedures used to

generate data in sections 3.3.1 to 3.3.4.


Samples of fractions from GP226 was separated and obtained using PSEC and HPLC. Please

refer to section 2.2.1 for further details relating to the fractionation process. In

experiments with fractionated GP226, amounts of each fraction were used at

concentrations equivalent to what were originally present in the unfractionated GP226

mixture. Hence, proportionate dilutions as a w/w % of the Fractions, compared to the

unpurified GP226, were prepared as treatments. For further details relating to the

dilutions, please see Table 2.1. The crude, unpurified GP226 and negative controls,

including PDMS and PEG, were prepared at concentrations ranging from 0.01% to 1% in cell

culture medium and used as treatments.


HSF, KF, nHSF and nKF were independently and routinely cultured for use in the

experiments performed in this chapter. Please refer to section 2.3.1 for specific details.


HK were isolated as previously described (Dawson et al., 2006) from human skin of

consenting patients undergoing elective abdominoplasty or mammoplasty procedures for

use within experiments required for this chapter. Ethics approval to use this tissue was

obtained from the Queensland University of Technology Human Research Ethics

C h a p t e r 3 . 0 | 41

Committee (ID:3673H). For more specific details relevant to HK culture, please refer to

section 2.3.2.

3.2.5 WST‐1 Assay for Determination of Cell Viability

Following culture, HSF, KF, nHSF, nKF and HK were seeded into 96 well plates (Nunc) for

treatment with silicones. Cultures of HSF and nHSF cells were seeded into each well and

were then treated 24 hours later with silicone solutions diluted in cell culture medium, as

described in section 3.2.1. Cultures of HK cells were seeded onto i3T3 cells in 96 well plates

and then treated 24 hours later with silicone solutions diluted in Full Green’s Medium, as

described in section 3.2.1. Controls included: a blank control, containing medium only; an

untreated control, containing cells treated with medium but no silicone; and negative

controls, including cells treated with the silicone controls described in section 3.2.1. During

all incubations, the cells were maintained in the standard conditions of 37°C and 5% CO2.

Following 48 hours silicone treatment, cell viability was measured using the WST‐1 Assay.

Further details describing the WST‐1 Assay methodology can be found in section 2.4.1.


Following culture, HSF, KF, nHSF, nKF and HK were seeded into 96 well plates (Nunc) for

treatment with silicones. Cultures of HSF or nHSF cells were seeded into each well and

were then treated 24 hours later with silicone solutions diluted in cell culture medium, as

described in section 3.2.1. Cultures of HK cells were seeded onto i3T3 cells in 96 well plates

and then treated 24 hours later with silicone solutions diluted in Full Green’s Medium, as

described in section 3.2.1. Controls included: a blank control, containing medium only; an


controls, including cells treated with the silicone controls described previously in section

3.2.1. During all incubations, the cells were maintained in the standard conditions of 37°C

and 5% CO2. Following 48 hours treatment with the silicones, cell proliferation was

measured using the Cyquant (Invitrogen) Assay Kit. Further details describing the Cyquant

Assay methodology can be found in section 2.4.2.

3.2.7 Analysis of Silicone and Protein Interaction

The morphology of HSF following culture in varying concentrations of FCS, with and

without silicone, was investigated to observe if GP226 was causing an aggregation of

proteins within the cell culture medium. HSF were plated into 24 well plates and cultured in


0%, 5%, 10% or 20% FCS/DMEM. GP226 (0.1%), diluted in the respective cell culture media,

was used to treat the cells 24 hours later. Following 48 hours treatment with the silicones,

HSF were stained with Sirius Red to facilitate cell morphology analysis and to localise the

collagen content within the cultures. Cell morphology was documented using digital

photography and light microscopy. Specific details for this experiment are described in

section 2.4.3. Aggregation of proteins in cell culture media was also assessed by gel

electrophoresis, with the distribution and molecular weight of proteins present being

stained by the Pierce Silver Stain kit. The gel was documented using a flat‐bed scanner.

Specific details for this experiment can be found in section 2.4.3.


Real‐time microscopy was used to visualize the response of HSF to silicone treatment over

48 hours. In brief, HSF cells were plated into 24 well plates and treated with GP226,

Fraction IV and Tween 20 (0.1%) 24 hours later. Real‐time microscopy was then performed

with a Leica AF6000 Widefield Microscope and images taken every 15 minutes for 48 hours.

Still images and movies were prepared using Leica Application Suite software. Specific

details can be found in section 2.4.4.


The induction of apoptosis following treatment of HSF to GP226, Fraction III, Fraction IV,

Tween 20 and PEG was measured using the Tunel assay. Cell cultures of HSF were prepared

in glass 8‐well chamber slides and treated 24 hours later with GP226, Fraction III, Fraction

IV, Tween 20 and PEG (0.1%) diluted in cell culture medium. An untreated control,

containing cells treated with media only, was included. After 48 hours incubation, the Tunel

assay was performed. Following this, a Nikon Eclipse TE2000‐U fluorescence microscope

was used to visualise and photograph the cells. For more specific details describing the

Tunel assay, please see section 2.4.5.

3.2.10 Statistical Analysis

All cell viability and proliferation experiments were performed three times, each using HSF

and nHSF cells from three different patients. Within each experiment, each variable was

tested in triplicate. Data were expressed as a mean ± SEM and assessed for significance

with analysis of variance followed by a pairwise t‐test post hoc. P values < 0.05 were

considered significant.

C h a p t e r 3 . 0 | 43

3.3 RESULTS

3.3.1 Analysis of Cell Viability and Proliferation following Treatment with Silicone

Significant decreases in cell viability occur following exposure of fibroblasts to GP226 have

been previously demonstrated (Charters, 2007). However, many aspects of these studies

required further investigation, including the optimal treatment time and methods of

measuring the functional activity of cells. It was therefore decided to measure cell viability

and proliferation of HSF, nHSF, KF and nKF following treatment with silicone for 24, 48, 72

and 168 hours (7 days). Cell viability and proliferation were measured using WST‐1, an

assay that measures cell metabolic activity, as well as Cyquant, an assay that measures DNA

content. The viability of HSF, nHSF, KF and nKF cells over 7 days are depicted in Figures 3.1‐

3.4 whereas the proliferation of HSF, nHSF, KF and nKF cells over 7 days are illustrated in

Figures 3.5‐3.8. The numerical values from the assays assessing viability and proliferation of

HSF following silicone treatment for 24, 48, 72 and 168 hours (7 days) are depicted in

Tables A 2.1 – A2.8 within Appendix 2.

These analyses reveal that from 48 hours onwards, GP226 significantly and dose

dependently decreased the viability of HSF, nHSF, KF and nKF compared to the untreated

controls. The proliferation data also revealed significant and dose dependant decreases

following treatment of GP226 from 48 hours onwards for HSF, nHSF, KF and nKF, compared

to the untreated controls.

It was observed that neither Fraction I nor II significantly decreased the viability of HSF,

nHSF, KF or nKF at 24, 48 and 72 hours until the highest concentration, 1%, was tested. At

168 hours (7 days), however, viability was dose dependently decreased when HSF, nHSF, KF

and nKF were treated with Fraction I and II. It was also observed on day 7 that the lower

concentrations, 0.01% and 0.03%, of Fractions I and II actually significantly increased nHSF

and nKF viability compared to the untreated controls. This trend was not observed in

assays assessing HSF and KF viability. Unlike the cell viability results, Fraction I and II dose

dependently decreased proliferation from 24 hours onwards in HSF, nHSF, KF and nKF. The

proliferation results obtained with Fraction I or II, however, were not as significantly

decreased as was found with GP226. As was observed in the cell viability data, treatment of

nHSF or nKF with Fraction I or II also did not increase proliferation.


Figure 3.1 – Analysis of HSF viability following treatment with GP226 and its fractions. Graphs of HSF viability, as measured by the WST‐1 assay, following treatment with silicone for (A) 24, (B) 48, (C) 72 and (D) 168 hours. Data are expressed as an average of the absorbance values (440 nm, ref = 620 nm) pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.

C h a p t e r 3 . 0 | 45

Figure 3.2 – Analysis of nHSF viability following treatment with GP226 and its fractions. Graphs of nHSF viability, as measured by the WST‐1 assay, following treatment with silicone for (A) 24, (B) 48, (C) 72 and (D) 168 hours. Data are expressed as an average of the absorbance values (440 nm, ref = 620 nm) pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.


Figure 3.3 – Analysis of KF viability following treatment with GP226 and its fractions. Graphs of KF viability, as measured by the WST‐1 assay, following treatment with silicone for (A) 24, (B) 48, (C) 72 and (D) 168 hours. Data are expressed as an average of the absorbance values (440 nm, ref = 620 nm) pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.

C h a p t e r 3 . 0 | 47

Figure 3.4 – Analysis of nKF viability following treatment with GP226 and its fractions. Graphs of nKF viability, as measured by the WST‐1 assay, following treatment with silicone for (A) 24, (B) 48, (C) 72 and (D) 168 hours. Data are expressed as an average of the absorbance values (440 nm, ref = 620 nm) pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.


Figure 3.5 – Analysis of HSF proliferation following treatment with GP226 and its fractions. Graphs of HSF proliferation, as measured by the Cyquant assay, following treatment with silicone for (A) 24, (B) 48, (C) 72 and (D) 168 hours. Data are expressed as an average of the fluorescence values (ex = 480 nm, em = 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.

C h a p t e r 3 . 0 | 49

Figure 3.6 – Analysis of nHSF proliferation following treatment with GP226 and its fractions. Graphs of nHSF proliferation, as measured by the Cyquant assay, following treatment with silicone for (A) 24, (B) 48, (C) 72 and (D) 168 hours. Data are expressed as an average of the fluorescence values (ex = 480 nm, em = 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.


Figure 3.7 – Analysis of KF proliferation following treatment with GP226 and its fractions. Graphs of KF proliferation, as measured by the Cyquant assay, following treatment with silicone for (A) 24, (B) 48, (C) 72 and (D) 168 hours. Data are expressed as an average of the fluorescence values (ex = 480 nm, em = 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.

C h a p t e r 3 . 0 | 51

Figure 3.8 – Analysis of nKF proliferation following treatment with GP226 and its fractions. Graphs of nKF proliferation, as measured by the Cyquant assay, following treatment with silicone for (A) 24, (B) 48, (C) 72 and (D) 168 hours. Data are expressed as an average of the fluorescence values (ex = 480 nm, em = 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.


Cell viability was significantly and dose dependently decreased to levels below the

untreated controls when Fractions III and IV were applied to HSF, as depicted in Tables 3.4

and 3.5 respectively. A similar decrease was found in KF cells from 48 hours onwards.

Viability was dose dependently, but not significantly, decreased when Fraction III was

applied to nHSF and nKF from 48 hours onwards. In fact, the viability of nHSF and nKF was

not decreased to the levels observed with GP226 following treatment with Fraction III over

the four timepoints investigated. Overall, the results obtained with Fraction IV were the

most similar to those observed with GP226 in terms of raw values obtained, dose

dependency and significance. Interestingly, Fraction IV did not significantly decrease nHSF

and nKF viability until day 7. The results obtained for the proliferation studies indicate that

both Fraction III and IV dose dependently decrease HSF, nHSF, KF and proliferation from 24

hours onwards. Once again, it was observed that Fraction IV decreased cell proliferation of

all cell types to levels most similar to those obtained for GP226 in terms of the raw values

obtained, dose dependency and significance.

Fraction V, at a concentration of 0.1% and greater, significantly decreased HSF, nHSF, KF

and nKF viability to levels well below those obtained for GP226. The numerical values for

HSF viability following treatment with Fraction V are depicted in Table 3.6. These results

demonstrating significantly reduced cell viability from 24 hours onwards suggest that

Fraction V was toxic. Interestingly, while the viability of HSF, nHSF, KF and nKF was virtually

zero when treated when the two highest concentrations, 0.3% and 1%, of Fraction V, the

proliferation assays (Figures 3.5‐3.8) revealed that a significant amount of DNA content was

still present compared to the untreated control.

The PEG and PDMS controls did not dose dependently reduce the viability and proliferation

of any cell type to the same extent as found with GP226. The numerical values for HSF

viability and proliferation following treatment with PEG and PDMS are depicted in Tables

3.7 and 3.8, respectively. In some cases, PEG and PDMS significantly increased the viability

and proliferation of nHSF and nKF compared to the untreated controls of nHSF and nKF.

These increases in viability and proliferation were particularly evident in the results

obtained at 168 hours (7 days). In terms of cell viability and proliferation of the untreated

controls, all cell types appeared to reach peak viability and DNA content at either 48 hours

or 72 hours. Therefore, 48 hours was selected as the best timepoint for further

experiments.

C h a p t e r 3 . 0 | 53

In view of the cell proliferation and viability results obtained following application of GP226

to fibroblasts, it was decided to investigate the effect of GP226 and its fractions on HK at 48

hours only (Figure 3.9). Analysis of the data obtained for HK revealed that PEG did not

significantly (p<0.05) decrease the viability or proliferation of HK compared to the

untreated controls. Fraction IV significantly (p<0.05) reduced both HK viability (0.82 ± 0.06)

and proliferation to the same extent as observed for GP226. While proliferation of HK in

response to Fraction V was significantly (p<0.05) reduced, and to a similar extent as GP226,

HK viability was found to be even further decreased, supporting the concept that Fraction V

is toxic. Taken together, these assays demonstrate that Fraction IV was most similar to the

crude GP226 when applied to HSF, nHSF, KF, nKF and HK.

3.3.2 Silicone and Protein Interaction

Silicone oil has previously been implicated in the induction of protein aggregation

(Bernstein, 1987). It was therefore hypothesized that the silicone being used to treat the

cells was lowering the availability of proteins through their aggregation. This would in turn

affect the cells indirectly by starving the cell cultures of proteins that are required for

normal function and growth. An indirect investigation of this hypothesis was undertaken

whereby the cells were cultured with cell culture medium containing varying amounts of

FCS, up to 20%, a concentration that provides double the standard concentration of FCS

used to culture the cells. Figure 3.10 illustrates the morphology of HSF cells when cultured

in 0%, 5%, 10% or 20% FCS and treated with GP226 (0.1%) at the same time. Following

treatment, the samples were treated with Sirius Red to stain collagen. Figure 3.10

demonstrates that the cell density of HSF is dramatically reduced after 48 hours of silicone

treatment in all cell culture conditions. Furthermore, the distribution of collagen present

within the cells was not altered in any of the different culture conditions. This suggests that

the results obtained with GP226 are not related to the amount of FCS present.


C

Figure 3.9 – Analysis of HK viability and proliferation following treatment with GP226 and its fractions. Graphs of (A) HK viability, as measured by WST‐1, and (B) HK proliferation, as measured by the Cyquant assay following treatment with silicone for 48 hours. (C) Numerical values of HK viability and proliferation following treatment with silicone for 48 hours. Data are expressed as an average of the absorbance (440 nm, ref = 620 nm) and fluorescence (ex = 480 nm, em = 520 nm) values obtained from three replicate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.

Cell Viability Cell Proliferation

Untreated 1.40 ± 0.08 12353 ± 1116

i3T3 0.17 ± 0.04 1367 ± 50.0

PEG ‐ 0.1% 1.45 ± 0.11 12477 ± 1019

GP226 ‐ 0.1% 0.77 ± 0.04 8138 ± 504

Fraction I ‐ 0.1% 1.36 ± 0.09 11143 ± 534

Fraction II ‐ 0.1% 0.05 ± 0.02 1582 ± 402

Fraction III ‐ 0.1% 1.40 ± 0.08 9491 ± 329

Fraction IV ‐ 0.1% 0.82 ± 0.06 9505 ± 291

Fraction V ‐ 0.1% 0.36 ± 0.03 8120 ± 283

C h a p t e r 3 . 0 | 55

Figure 3.10 – Investigation of HSF morphology following culture in media containing different concentrations of FCS and treatment with GP226. Images of HSF morphology obtained via light microscopy following culture in media containing 0%, 5%, 10% or 20% FCS and following treatment with GP226 for 48 hours. The untreated control is HSF treated with medium containing no silicone. Cells were stained with Sirius Red and representative images are depicted from three replicate experiments (n = 3). Scale bar represents 100 µm.


The phenomenon of protein aggregation following interaction with silicone was also

investigated via gel electrophoresis. Figure 3.11 illustrates the molecular weight and

distribution of proteins present within cell culture medium containing 0%, 5%, 10% or 20%

FCS, with and without GP226. It is observed that the distribution of proteins do not differ

between the cell culture media containing no silicone and the cell culture media containing

GP226. Specifically, no high molecular weight bands indicative of aggregated protein

appear in cell culture media containing GP226. Taken together, GP226 does not appear to

be exerting its action via the induction of protein aggregation in cell culture medium.

Figure 3.11 – Analysis of protein aggregation in cell culture medium containing silicone and varying concentrations of FCS Electrophoresed gel of proteins present in cell culture media with and without 0.1% GP226 and containing 0%, 5%, 10% or 20% FCS. Proteins present were silver stained for determination of distribution and molecular weight. Lane M indicates the protein molecular weight standard. Cell culture media containing no silicone and varying concentrations of FCS are depicted in lanes 1 (0% FCS), 2 (5% FCS), 3 (10% FCS) and 4 (20% FCS). Cell culture media containing 0.1% GP226 and varying concentrations of FCS are depicted in lanes 5 (0% FCS), 6 (5% FCS), 7 (10% FCS) and 8 (20% FCS).

C h a p t e r 3 . 0 | 57

3.3.3 Further Investigation of HSF Morphology following Treatment with Fraction IV

In view of the cell viability and proliferation data, real time microscopy was used to observe

the direct response of HSF to GP226, Fraction IV and Tween 20 at concentrations between

0.01% and 1%. Tween 20 was included as it is a surfactant like GP226 and Fraction IV.

Figure 3.12 illustrates the morphology of HSF when treated with Fraction IV and Tween 20

at a concentration of 0.1% for 48 hours. It can be observed that HSF density and

morphology were dramatically affected following treatment with Fraction IV and Tween 20.

Furthermore, HSF undergoing apoptosis exhibited characteristics such as membrane

blebbing and cytoplasm swelling after 24 and 48 hours of treatment. Similar results were

obtained when HSF were treated with GP226 (data not shown). When HSF were treated

with Tween 20, membrane swelling and movement, resulting in fuzzy images, was evident

immediately. This suggests that the HSF treated with Tween 20 were undergoing a form of

apoptosis called anoikis, which is induced when anchorage‐dependent cells detach from

the surrounding ECM (Frisch and Screaton, 2001). Furthermore, this phenomenon of the

immediate membrane swelling and detachment from the cell culture surface was not

observed when HSF were treated with GP226 or Fraction IV, suggesting that anoikis was

not occurring in these samples.

3.3.4 Investigating the Induction of HSF Apoptosis following Treatment with Silicone.

The observations recorded with real‐time microscopy prompted us to examine the

induction of apoptosis following treatment of fibroblasts with the amphiphilic silicones.

This has not been reported in the literature previously. Immunofluorescent microscopy

analysis, using the Tunel assay, of apoptotic HSF in response to GP226, Fraction III and

Fraction IV (0.1%), is depicted in Figure 3.13. This reveals that the positive control, including

untreated cells incubated with DNAse I, exhibited positive immunofluorescence for the

assay. Furthermore, apoptotic cells are evident in the HSF cultured with GP226, Fraction III

and Fraction IV. Although not observed in any other samples, the small size of positive

immunofluorescent particles in the HSF sample treated with Fraction III suggests that the

positive staining may well be background immunofluorescence. Nevertheless, background

immunofluorescence was not detected in the negative control. Furthermore, no apoptotic

cells were detected in the untreated HSF or the HSF cultures treated with PEG or Tween 20.


Figure 3.12 – Analysis of HSF morphology following treatment with Fraction IV and Tween 20. Images of HSF morphology obtained through real‐time microscopy following treatment with Fraction IV and Tween 20 for 48 hours. The untreated control includes HSF treated with medium containing no silicone. Representative images are depicted from three replicate experiments, in which each treatment was performed in triplicate (n = 9). Examples of apoptotic cells are indicated by the arrows. Scale bar represents 50 µm.

C h a p t e r 3 . 0 | 59

Figure 3.13 – Investigation of apoptosis in HSF following treatment with GP226, Fraction III, Fraction IV and Tween 20. Immunofluorescent images of Tunel positive HSF following treatment with silicone and Tween 20 for 48 hours. Green indicates Tunel positive cells, while blue localises the cell nucleus. Controls displayed include the untreated control (HSF treated with cell culture medium containing no silicone), positive control (including fixed and permeabilised untreated HSF treated with DNAse I) and negative control (including untreated HSF stained with the labelling solution only without enzyme). Representative images are depicted from three replicate experiments, in which each treatment was performed in triplicate (n = 9). Scale bar represents 100 µm.


3.4 DISCUSSION

The primary aim of the experiments in this chapter was to investigate the potential scar

remediating ability of a commercially available silicone species, GP226, that is structurally

similar to those found previously to traverse the stratum corneum (Sanchez et al., 2005). It

is reported herein that GP226, fractioned into five species of differing molecular weights,

reduces cell viability and proliferation of HSF, nHSF, KF and nKF. Interestingly, differing

effects on cell viability and proliferation were observed when the cells were treated with

the five fractions. Of note, application of Fraction IV to dermal fibroblasts in vitro, was

found to induce effects similar to GP226. Furthermore, it was demonstrated that GP226

and its active fraction, IV, induces apoptosis in dermal fibroblasts derived from

hypertrophic scars.

These investigations were motivated by a need to validate the earlier findings from our

laboratory indicating that exposure of dermal fibroblasts to GP226 and its fractions

decreased cell viability. While investigating cell viability provides a measure of

metabolically viable cells present, it doesn’t, however, indicate the total number of cells

present. Therefore, it was evident that not only cell viability but cell proliferation, which

measures the amount of DNA present, needed to be evaluated. While prior experiments

demonstrated that significant decreases in cell viability occur following exposure of

fibroblasts to GP226 for 48 hours, Hanasono et al. (2004) reported that differences in the

cell growth curves of dermal fibroblasts occurred between days 2 and 5 following

treatment with silicone. This prompted the examination of cells in our functional studies at

a range of different timepoints. Cell viability and proliferation of HSF, nHSF, KF and nKF

following treatment with silicone was therefore measured at 24, 48, 72 and 168 hours (7

days).

The data generated through these studies confirmed the previously documented results,

revealing that not only viability, but also proliferation, of HSF, nHSF, KF and nKF was

affected by GP226 and its fractions over the four time points investigated. It was also

observed that Fractions III, IV and V are the most active components of GP226 and

decreased the viability, as well as proliferation, of all cell types in a dose‐dependent

manner (Figure 3.1‐3.8). It was clear that Fraction IV and V induced effects most similar to

those found with GP226 in HSF, nHSF, KF and nKF. However, Fraction V was found to

produce inconsistent results in other experiments performed within our laboratory,

C h a p t e r 3 . 0 | 61

especially as it contained contaminants of the GP226 manufaction process (Radi, B. pers.

comm.). Unlike the data generated with the preliminary assays (Charters, 2007), Fractions I

and II dose dependently reduced the viability and proliferation of all cell types, albeit not to

the same extent as observed with GP226. Unlike Hanasono et al., (2004), increases in

viability or proliferation of all cell types following treatment with the silicones was not

observed after 48 hours of treatment with silicone. In view of this, 48 hours was selected as

the best timepoint for further experiments as this was the timepoint that led to the

greatest proliferation in the most of the control treatments. With respect to these, the

effect of PEG and PDMS actually significantly increased the viability and proliferation of the

dermal fibroblasts, particularly at 168 hours (7 days), to levels well above the control

(Figures 3.1‐3.8). These results were significant as they demonstrated that the decreases in

cell viability and proliferation observed in fibroblasts exposed to GP226 and its fractions

were due to the unique structure of PDMS‐PEG copolymers, but not the PDMS or PEG

structures individually.

There are two main cell types, keratinocytes and fibroblasts, within the epidermis and dermis of

skin (Werner et al., 2007). Responses of keratinocytes (HK) as well as fibroblasts to GP226 and

its fractions were therefore assessed (Figure 3.9). Interestingly, in terms of cell viability and

proliferation, the HK responses were different to the fibroblasts. For example, 262.5 ng/mL of

Fraction II decreased HK viability and proliferation significantly, a concentration that did not

affect fibroblast viability or proliferation. This difference, however, was not explored further in

these doctoral studies as the focus of the experiments was on fibroblasts. Furthermore, while

the control treatments increased fibroblast viability and proliferation, the application of PEG to

HK did not significantly affect cell viability or proliferation. Taken together, these data suggest

that different cell types within skin tissue have different responses to the silicones, illustrating

the importance of analysing the effect of GP226 and its fractions on both fibroblasts and

keratinocytes. Nevertheless, it was still clear that Fraction IV exhibited the most similar results

to those that were observed with GP226.

Another important aspect of the analyses conducted here is that the effects of both HSF

and KF were examined. This is especially important the effect of GP226 on keloid‐derived

tissues has not been reported in the literature. It has previously been reported that

silicone‐related effects are not as evident in keloid‐derived fibroblasts than when

compared to effects observed in hypertrophic‐derived fibroblasts (Hanasono et al., 2004).


Our analyses of cell viability and proliferation, however, demonstrated no major

differences between HSF and KF or nHSF and nKF when compared to the controls. Keloid

and hypertrophic scars are histologically and morphologically different (Bayat et al., 2003)

and the fibroblasts derived from keloid scars appear and behave differently to those

derived from hypertrophic scars (Hanasono et al., 2004). Furthermore, while dermal

fibroblasts have been derived from keloid scars and used in vitro (Bayat et al., 2003) for a

variety of wound repair studies, it is not known if these differences in KF behaviour are

typical of their in vivo phenotype. Unlike others, no differences in KF behaviour were

observed, but the fact that cells of passage 4‐7 were used, rather than cells derived directly

from tissues, may explain the similarities observed. Nevertheless, in view of the lack of

pronounced differences between HSF and KF in our studies, the future analyses described

in this thesis investigated only HSF and nHSF.

The decreases in cell viability and proliferation observed following application of GP226

and its fractions to HSF, nHSF, KF and nKF caused us to consider what mechanisms

underpinned the effects of the silicone on cells. One hypothesis considered was that the

silicone being used to treat the cells was inducing aggregation of FCS proteins within the

cell culture medium and was therefore ‘starving’ the cells of their required nutrients.

Silicone oil has previously been implicated in the induction of protein aggregation

(Bernstein, 1987). Jones et al. (2005) reported that silicone oil‐induced aggregation of

proteins occurred dramatically with bovine serum albumin (BSA), possibly causing it to

undergo gross conformational changes and rendering it non‐functional. However, when

HSF were cultured with medium containing FCS of varying concentrations (up to 20%) and

simultaneously treated the cells with silicone, cell density was still dramatically decreased

(Figure 3.10). Similar responses were obtained regardless of what concentrations of FCS

were present. Furthermore, gel electrophoresis of cell culture media containing silicone

and different concentrations of FCS did not indicate any higher molecular weight,

aggregated protein products compared to cell culture media containing FCS but no silicone

(Figure 3.11).

Following on from this, cell morphology following treatment with GP226, Fraction IV and

Tween 20 was investigated. Tween 20 was included as it is a surfactant like GP226 and

Fraction IV. It became evident through real‐time microscopy that HSF exhibited

morphology indicative of apoptosis. This included cell shrinkage, membrane blebbing and

C h a p t e r 3 . 0 | 63

nuclear fragmentation following treatment with GP226 and Fraction IV (Figure 3.12). This

morphology was not observed when Tween 20 was used as a treatment, demonstrating

that not all surfactants have the same effect upon exposure to dermal fibroblasts. While

the cell viability and proliferation assays used in this study only determined the presence

and viability of cells, the mode of death is not evident. Therefore, the presence of HSF

undergoing apoptosis following treatment with GP226 and Fraction IV was confirmed using

the Tunel assay (Figure 3.13). Decreased apoptosis of fibroblasts has been reported as a

major factor in the etiopathogenesis of hypertrophic and keloid scars (Appleton et al.,

1996; Saray and Gulec, 2005; Sayah et al., 1999). The regulation of ECM deposition and

remodelling is a key element in hypertrophic scar formation (Eckes et al., 2000), but tissue

homeostasis, which is maintained through a balance between cell proliferation and death,

is equally important (Bellemare et al., 2005; Moulin et al., 2004). Moreover, it is

increasingly apparent that subtle moderations of scar microenvironments, other than

reduced collagen production, can influence the disappearance and maintenance of wound

granulation tissue (Moulin et al., 2004). The development of a therapy containing the active

fraction (IV) of GP226 therefore has potential as a scar treatment as it may remove the

tissue bulk associated with abnormal scars, as well as eliminate any excess collagen

production through the removal of cells that are producing it.

Cytotoxic agents are in fact currently being used clinically to reduce the extraneous tissue

associated with hypertrophic scar formations (Espana et al., 2001; Meier and Nanney,

2006). One such example is bleomycin, an apoptotic agent that is also used in

chemotherapy (Yamamoto, 2006). Although this therapy has shown encouraging results in

remediating scars in many reports (Espana et al., 2001; Saray and Gulec, 2005), the

injection of bleomycin into the skin of healthy subjects has been found through histology to

cause an increase in inflammatory infiltrate and higher rates of keratinocyte necrosis

(Templeton et al., 1994). Furthermore, its use requires application by injection, an invasive

procedure with increased associated complications. The use of bleomycin also has the

potential to cause major side‐effects of systemic toxicity, as well as pulmonary, renal and

cutaneous fibrosis in the longer term (Crooke and Bradner, 1976; Shastri et al., 1971).

Clearly, a non‐invasive topical silicone treatment that stimulates an equivalent apoptotic

effect with minimal side‐effects would be advantageous.


An implication of our findings, however, involves the issue of whether the amphiphilic

silicone products investigated in our study are truly inert within the human body. It is

generally accepted that silicone is an inert substance (Chan et al., 2005) and numerous

clinical studies on silicone gel‐filled breast implants have failed to find any association with

increased rates of systemic disease (Lewin and Miller, 1997). However, silicone‐PEG

copolymers, consisting of PDMS and PEG structures similar to that in GP226, have also

been reported to be physically inert. In fact, silicone‐PEG copolymers are used as surface

tension depressants, wetting agents and emulsifiers in cosmetics, often being included at

concentration ranges of 0.1% ‐ 10% (Sage, 1982). Despite this, our own findings suggest

that silicone‐PEG copolymers, such as GP226 and Fraction IV, are not inert and induce

decreases in cell viability, proliferation and an increase in apoptosis. This demonstrates that

not all silicone‐PEG copolymers are inert within the human body. It is hypothesised that the

low molecular weight of GP226 and Fraction IV contributes to their unique action following

exposure to dermal fibroblasts. In summary, the results reported here suggest that GP226,

and particularly Fraction IV, are novel silicone species with potential for clinical application

in scar remediation.

C h a p t e r 4 . 0 | 65

CHAPTER 4.0

FUNCTIONAL ANALYSIS OF THE EFFECTS OF SYNTHETIC

SILICONES, INCLUDING PDMS7‐g‐PEG7, ON DERMAL

FIBROBLASTS

4.1 INTRODUCTION

The results described in Chapter 3.0 demonstrated that GP226 and its active fraction, IV,

reduced cell viability, as well as proliferation, and induced apoptosis in fibroblasts derived

from hypertrophic scars. These results were significant as GP226 was a crude polymer

mixture containing different fractions of varying molecular weight and it was unknown if

the individual fractions had similar biological effects. By separating GP226 via PSEC and

HPLC and analysing the different components through various cell‐based functional assays,

it was found that only a small proportion of the polymer mixture, namely Fraction IV, was

responsible for the effects elicited by GP226. The fractionation approach, however, yielded

only 8% of Fraction IV, meaning that a large amount of GP226 was required in order to

obtain small amounts of Fraction IV (Gardoni, M. pers. comm.). Furthermore, the

fractionation methodology was labour‐intensive and time inefficient.

In view of this, methods that could be used to more efficiently isolate and purify Fraction IV

from GP226 were examined (Dickfos, 2008). Reverse‐phase HPLC was one such method

that was further investigated and Figure 4.1 depicts the resultant chromatograms obtained

for GP226 and Fraction IV. It is evident in Figure 4.1 that Fraction IV, depicted as the ‘active’

fraction, was found to be a mixture of compounds with differing chemical properties. These

differences most likely arise from variations in the silicone (PDMS) backbone and PEG side

chain lengths within Fraction IV (Dickfos, 2008).

It was demonstrated through these studies that the HPLC method could not be optimised

to become more efficient at fractionating Fraction IV. It therefore became apparent that

the silicones would need to be synthesised to be used as treatments in our laboratory in

order to achieve the level of purity and quantities of silicones required for experimentation.

The silicones to be synthesised were based on the structure of Fraction IV. While Fraction

IV was not one single species of specific structure (Figure 4.1), it was the silicone treatment

that had demonstrated the most consistent and dose‐dependant results in our in vitro

F u n c t i o n a l A n a l y s i s o f t h e S y n t h e t i c S i l i c o n e s

assays of cell viability and proliferation. Therefore, the structure of Fraction IV was fully

characterised by matrix‐assisted laser desorption/ionization (MALDI) and nuclear magnetic

resonance (NMR) analysis, with the resulting structure depicted in Figure 4.2 (Dickfos,

2008). It can be observed in Figure 4.2 that Fraction IV was found to be a rake amphiphilic

silicone consisting of a PDMS backbone and PEG side chains. Based on this structure, a

range of amphiphilic silicones were subsequently synthesised.

Figure 4.1 – Molecular weight distribution of the crude GP226 and active fraction, Fraction IV. Chromatogram of GP226 and Fraction IV as determined by HPLC (Dickfos, 2008).

Figure 4.2 – Structure of Fraction IV Chemical structure of Fraction IV showing the rake structure comprised of a silicone backbone and PEG sidechains (Dickfos, 2008).

C h a p t e r 4 . 0 | 67

The synthesis of amphiphilic silicones, also known as silicone surfactants, is well reported,

with applications ranging from cosmetics, detergents, toiletries, emulsifying agents and in

agriculture (Hill, 1999, 2002; Schlachter and Feldmann‐Krane, 1998). Synthesis of a series of

amphiphilic silicones, including both rake‐ and block‐ substructures, were performed by

chemists within our laboratory (Dickfos, 2008; Keddie et al., 2010). A two‐step approach

was undertaken whereby a range of PDMS backbones and PEG sidechains with differing

oligomeric length were separately synthesised before being coupled together. Different

combinations of PDMS and PEG were coupled together to generate a small library of

amphiphilic silicone oligomers with low molecular weight and a broad range of properties

(Table 4.1). Importantly, the amphiphilicity of the synthesised silicones, as indicated by

their hydrophilic‐lipophilic balance (HLB) factor, varied greatly. Of note, the equation for

HLB is 20 x (MH / (MH + ML) (Griffin, 1954). The HLB scale operates on a scale of 1‐20,

whereby lower HLB values indicate more hydrophobic compounds, such as PDMS15.2‐PEG8

and PDMS15.2‐PEG4, while higher values indicate more hydrophilic compounds, including

PDMS10.5‐PEG8 and PDMS7‐g‐PEG7 (Robbers and Bhatia, 1961).

Oligomer ID No. of Silicone

repeat units

No. of PEG

repeat units

Type of

Structure

Molecular

weight (Da)

HLB

factor

PDMS15.2‐PEG8 15.2 8 Block 2036 8.36

PDMS10.5‐PEG8 10.5 8 Block 1687 10.08

PDMS15.2‐PEG4 15.2 4 Block 1684 5.92

PDMS7‐g‐PEG7 7 7 Rake 1265 12.06

Table 4.1 – Properties of the synthesised amphiphilic silicones used as treatments. Properties of PDMS15.2‐PEG8, PDMS10.5‐PEG8, PDMS15.2‐PEG4 and PDMS7‐g‐PEG7 are presented.

The purpose of the studies reported within this chapter is to evaluate the scar remediating

potential of these various amphiphilic silicone species. To investigate this, cell proliferation

and the induction of apoptosis in HSF, nHSF and HK was evaluated following treatment

with these new synthetic silicones.



Full specifications of materials and methods used in experimental procedures for this

chapter are described in Chapter 2. Following on is a summary of the procedures used to

generate data in sections 4.3.1 to 4.3.4.


For full specifications of the synthesis of PDMS‐PEG oligomers used in this chapter, please

see section 2.2.2 as well as Appendix 1. The synthetic silicones were prepared at

concentrations ranging from 0.03% to 0.3% for screening experiments, and from 0.0001%

to 1.0% for other experiments, in cell culture medium and used as treatments applied to

cultured cells. In the experiment examining cell proliferation, the PDMS backbone, PDMS7,

was included as a negative control.


HSF and nHSF were independently and routinely cultured for use in the experiments

performed in this chapter. Please refer to section 2.3.1 for more specific culture details.


HK were isolated as previously described (Dawson et al., 2006) from human skin of

consenting patients undergoing elective abdominoplasty or mammoplasty procedures for

use within experiments required for this chapter. Ethics approval to use this tissue was

obtained from the Queensland University of Technology Human Research Ethics

Committee (ID:3673H). For more specific details relevant to HK culture, please refer to

section 2.3.2.


Following culture, HSF, nHSF and HK were seeded into 96 well plates (Nunc) for treatment

with silicones. Cultures of HSF and nHSF cells were seeded into each well and were then

treated 24 hours later with silicone solutions, as described in section 4.2.1, diluted in cell

culture medium. Cultures of HK cells were seeded onto i3T3 cells in 96 well plates and then

24 hours later silicone solutions, as described in section 4.2.1, diluted in Full Green’s

Medium were added. In some experiments, HK were seeded into the 96 well plates without

an i3T3 feeder layer. Controls included: a blank control, containing medium only; an


C h a p t e r 4 . 0 | 69

control, including cells treated with the PDMS backbone, PDMS7, . During all incubations,

the cells were maintained in the standard conditions of 37°C and 5% CO2. Following 48

hours treatment with silicone, cell proliferation was measured using the Cyquant Assay Kit.

Further details describing the Cyquant Assay methodology can be found in section 2.4.2.


Real‐time microscopy was used to visualize the response of HSF, nHSF and HK to silicone

treatment over 48 hours. In brief, HSF, nHSF and HK cells were plated into 24 well plates

and treated with PDMS7‐g‐PEG7 (0.01 % and 0.03%) 24 hours later. Real‐time microscopy

was then performed with a Leica AF6000 Widefield Microscope and images taken every 15

minutes for 48 hours. Still images and movies were prepared using Leica Application Suite

software. Please refer to section 2.4.4 for more specific details.


The induction of apoptosis following treatment of HSF, nHSF and HK to PDMS7‐g‐PEG7 was

measured using the Tunel assay. Cell cultures of HSF, nHSF and HK were prepared in glass

8‐well chamber slides and treated with PDMS7‐g‐PEG7 (0.003% to 0.03%) diluted in cell

culture medium 24 hours later. An untreated control, containing cells treated with media

only, was included. After 48 hrs incubation, the Tunel assay was performed. Following this,

a Nikon Eclipse TE2000‐U fluorescence microscope was used to visualise and photograph

the cells. For more specific details describing the Tunel assay, please see section 2.4.5.


All experiments were performed three times with each variable tested in triplicate. For

each replicate experiment performed, cells isolated from a different patient were used. For

all proliferation assays and the quantification of apoptotic cells, data was expressed as a

mean ± SEM and significance determined via analysis of variance followed by a pairwise t‐

test post hoc. P values < 0.05 were considered significant.

4.3 RESULTS

4.3.1 Analysis of Cell Proliferation following Treatment with the Synthesised Silicones

It was previously established that GP226 and its active Fraction, IV, significantly decreased

cell viability and proliferation when exposed to fibroblasts and keratinocytes. It was also

identified that a treatment time of 48 hours in vitro was optimal. Here, cell proliferation of


HSF, nHSF and HK following treatment with the synthesized silicones prepared within our

laboratory was measured. As our previous results showed similarities between HSF and KF

as well as nHSF and nKF, it was decided to only study HSF and nHSF from hereon. In

addition, as no major differences in our previous proliferation and viability data, only

proliferation of HSF, nHSF and HK following exposure to the synthetic silicones were

investigated.

The response of HSF and nHSF following 48 hours treatment with the synthetic silicones are

illustrated in Figure 4.3 and Figure 4.4, respectively. The silicones each had differing effects

upon application to HSF and nHSF, despite the fact that the various silicones were of similar

molecular weight and structure. For example, PDMS15.2‐PEG8 and PDMS15.2‐PEG4,

significantly and dose‐dependently decreased HSF and nHSF proliferation to levels similar

to those observed for GP226, compared to the untreated controls. PDMS10.5‐PEG8 also

significantly decreased both HSF and nHSF proliferation, however, the results were not

dose dependant. The most significant and dose dependant decreases in cell proliferation

were observed when HSF and nHSF were treated with PDMS7‐g‐PEG7, whereby

proliferation was decreased beyond that obtained with GP226. For this reason, PDMS7‐g‐

PEG7 was chosen for further examination.

Figure 4.3 ‐ Analysis of HSF proliferation following treatment with the synthesised silicones. Graph of HSF proliferation, as measured by the Cyquant assay, following treatment with the synthesised silicones for 48 hours. Data are expressed as an average of the true values pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Each replicate experiment used fibroblasts from a different patient. Significant differences (*) are illustrated where P<0.05 relative to the untreated control. Error bars indicate SEM.

C h a p t e r 4 . 0 | 71

Figure 4.4 ‐ Analysis of nHSF proliferation following treatment with the synthesised silicones. Graph of nHSF proliferation, as measured by the Cyquant assay, following treatment with the synthesised silicones for 48 hours. Data are expressed as an average of the true values pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Each replicate experiment used fibroblasts from a different patient. Significant differences (*) are illustrated where P<0.05 relative to the untreated control. Error bars indicate SEM.

4.3.2 Further Analysis of Cell Proliferation following Treatment PDMS7‐PEG7

In view of the cell proliferation data obtained when screening the synthesised silicones,

PDMS7‐g‐PEG7 was further investigated by analysing its effect on HSF and nHSF proliferation

over a larger range of concentrations. In addition, the silicone backbone of PDMS7‐PEG7,

PDMS7 was also analysed. The response of HSF and nHSF to 48 hours treatment of GP226,

PDMS7‐g‐PEG7 and PDMS7 are depicted in Figure 4.5 and 4.6, respectively. Firstly, it is

observed that 48 hour treatment with GP226 and PDMS7‐g‐PEG7 dose dependently and

significantly decreased HSF and nHSF proliferation, compared to the untreated control. It

was evident that PDMS7‐g‐PEG7 exerted its effects on HSF and nHSF at a much lower

concentration that what was required with GP226. For example, a concentration of 0.003%

PDMS7‐g‐PEG7 induced similar effects on HSF and nHSF to those obtained with 0.1% GP226.

With regards to the silicone backbone control, PDMS7 did not significantly or dose

dependently decrease HSF proliferation. However, at some concentrations, PDMS7

significantly decreased nHSF proliferation, albeit these decreases were not dose dependent

nor consistent.


A

B

Figure 4.5 ‐ Analysis of HSF proliferation following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7. (A) Graph of HSF proliferation, as measured by the Cyquant assay, following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7 for 48 hours. (B) Numerical values of HSF proliferation following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7 for 48 hours. Data are expressed as an average of the true values pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Each replicate experiment used fibroblasts from a different patient. Significant differences (*) are illustrated where P<0.05 relative to the untreated control. Error bars indicate SEM.

Untreated 4413 ± 319.9

Treatment GP226 PDMS7‐g‐PEG7 PDMS7

0.0001% 4338 ± 359.4 4408 ± 453.2 3793 ± 238.3

0.0003% 4063 ± 319.2 3974 ± 363.7 3863 ± 280.0

0.001% 3747 ± 206.4 3670 ± 446.4 4016 ± 412.4

0.003% 3764 ± 354.6 2688 ± 381.6 4654 ± 512.4

0.01% 3293 ± 367.9 789 ± 133 4387 ± 541.3

0.03% 2915 ± 403.2 536 ± 53.9 4113 ± 479.6

0.1% 2510 ± 243.1 115 ± 16.0 4436 ± 557.2

0.3% 1715 ± 274.0 172 ± 20.8 4625 ± 649.8

1.0% 551 ± 73.8 356 ± 63.6 4324 ± 570.4

C h a p t e r 4 . 0 | 73

A

B

Figure 4.6 ‐ Analysis of nHSF proliferation following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7. (A) Graph of nHSF proliferation, as measured by the Cyquant assay, following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7 for 48 hours. (B) Numerical values of nHSF proliferation following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7 for 48 hours. Data are expressed as an average of the true values pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Each replicate experiment used fibroblasts from a different patient. Significant differences (*) are illustrated where P<0.05 relative to the untreated control. Error bars indicate SEM.



0.0001% 2981 ± 387.6 3648 ± 381.9 3348 ± 301.6

0.0003% 3131 ± 252.8 3137 ± 308.8 2963 ± 327.8

0.001% 2865 ± 203.8 2817 ± 226.8 3229 ± 256.7

0.003% 2795 ± 171.9 2542 ± 305.0 3249 ± 313.6

0.01% 2432 ± 290.5 710 ± 89.0 3356 ± 191.4

0.03% 2049 ± 212.4 450 ± 31.1 3083 ± 231.1

0.1% 1887 ± 282.4 82.0 ±14.2 3048 ± 291.2

0.3% 1534 ± 257.4 146 ± 27.4 3317 ± 318.2

1.0% 516 ± 97.2 329 ± 78.5 3547 ± 305.4


In view of these results, the effect of GP226, PDMS7‐g‐PEG7 and PDMS7 on HK were

investigated as well (Figure 4.7). These studies reveal that similar decreases, to what was

found in HSF proliferation assays, occur in HK following 48 hours treatment with GP226,

PDMS7‐g‐PEG7 and PDMS7. For example, lower concentrations of PDMS7‐g‐PEG7 than GP226

PEG7 exerted a similar effect on HK as found with 0.01% GP226. In addition, PDMS7 did not

dose dependently decrease HK proliferation, even through some concentrations elicited

significant decreases in HK proliferation. Furthermore, the i3T3 control showed essentially

no proliferation compared to the untreated control. This indicates that the feeder layer was

successfully removed before HK proliferation was assessed.

It is evident that PDMS7‐g‐PEG7 affects both fibroblasts and keratinocytes. However, the

results of these first HK studies, involving HK grown on a fibroblast i3T3 feeder layer,

caused us to question whether the silicone species were actually affecting the

keratinocytes or the fibroblast layer beneath it. Therefore, HK were cultured without the

i3T3 feeder layer and once again measured proliferation following 48 hours treatment with

GP226, PDMS7‐g‐PEG7 and PDMS7 (Figure 4.8). This revealed that some concentrations of

GP226, PDMS7‐g‐PEG7 and PDMS7 significantly decreased HK proliferation compared to the

untreated control. These decreases in HK proliferation were, however, not dose

dependent. Furthermore, the untreated control (Figure 4.8) led to much less proliferation

compared to the untreated control grown on an i3T3 layer (Figure 4.7).

C h a p t e r 4 . 0 | 75

A

B

Figure 4.7 ‐ Analysis of HK proliferation following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7. (A) Graph of HK proliferation, as measured by the Cyquant assay, following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7 for 48 hours. (B) Numerical values of HK proliferation following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7 for 48 hours. Data are expressed as an average of the true values pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Each replicate experiment used fibroblasts from a different patient. Significant differences (*) are illustrated where P<0.05 relative to the untreated control. Error bars indicate SEM.

Untreated 18142 ± 1048

i3T3 603 ± 141


0.0001% 14888 ± 1978 15141 ± 1727 13646 ± 1664

0.0003% 13347 ± 1587 13951 ± 1446 15552 ± 2045

0.001% 12754 ± 1771 11782 ± 1727 15431 ± 1995

0.003% 11820 ± 1673 11022 ± 1162 16258 ± 1577

0.01% 10377 ± 1591 6324 ± 887 14263 ± 1248

0.03% 9237 ± 1025 100 ± 34.0 13218 ± 1508

0.1% 7721 ± 964 112 ± 37.0 14263 ± 1248

0.3% 3717 ± 929 315 ± 117 13218 ± 1508

1.0% 358 ± 64 337 ± 112 14061 ± 1835


A

B

Figure 4.8 ‐ Analysis of HK proliferation, grown without an i3T3 feeder layer, following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7. (A) Graph of HK proliferation, as measured by the Cyquant assay, following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7 for 48 hours. HK were cultured without an i3T3 feeder layer. (B) Numerical values of HK proliferation, grown without an i3T3 feeder layer, following treatment with GP226, PDMS7‐g‐PEG7 and PDMS7 for 48 hours. Data are expressed as an average of the true values pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Each replicate experiment used fibroblasts from a different patient. Significant differences (*) are illustrated where P<0.05 relative to the untreated control. Error bars indicate SEM.



0.0001% 2203 ± 164.8 2110 ± 155.0 1777 ± 52.69

0.0003% 2121 ± 135.6 2194 ± 343.3 1892 ± 76.73

0.001% 1757 ± 162.7 2254 ± 399.7 2146 ± 194.2

0.003% 1922 ± 142.1 1963 ± 66.06 2272 ± 156.4

0.01% 2050 ± 97.18 2266 ± 271.2 2230 ± 264.6

0.03% 2178 ± 270.9 1998 ± 255.8 1978 ± 87.97

0.1% 2030 ± 287.8 1649 ± 238.8 1900 ± 58.21

0.3% 1820 ± 224.5 1583 ± 202.2 1858 ± 74.17

1.0% 1684 ± 348.1 1680 ± 257.5 1967 ± 88.85

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4.3.3 Investigation of Cell Morphology following Treatment with PDMS7‐PEG7

Real time microscopy was used to observe the direct response of fibroblasts to PDMS7‐

PEG7. Figure 4.9 and 4.10 illustrate the effect of PDMS7‐PEG7, at concentrations of 0.01%

and 0.03%, on HSF and nHSF, respectively. The full sequence of images taken over 48 hours

for these cultures via real time microscopy are provided in Appendix 3.1 to 3.6. It can be

observed that HSF and nHSF displayed different effects depending on the concentration of

PDMS7‐g‐PEG7 used. HSF or nHSF displaying apoptotic morphology were not observed

during the 48 hours treatment with 0.01% PDMS7‐g‐PEG7. Despite this, the density of HSF

and nHSF following 48 hours treatment with 0.01% PDMS7‐g‐PEG7, was evidently decreased

compared to the untreated control. Following application of 0.03% PDMS7‐g‐PEG7 to HSF

and nHSF, apoptosis was evident by membrane blebbing and by the swelling of the

cytoplasms of cells from 12 hours onwards. Furthermore, micelles of PDMS7‐g‐PEG7, as

depicted by the large round bubbles in Figures 4.9 and 4.10, are observed in both HSF and

nHSF cultures treated with 0.03% PDMS7‐g‐PEG7 from 12 hours onwards. Smaller micelles

are also evident in HSF and nHSF cultures treated with 0.01% PDMS7‐g‐PEG7 from 12 hours

onwards.

The response of HK, grown with and without an i3T3 feeder layer, was also documented by

real time microscopy and is illustrated in Figures 4.11 and 4.12 respectively. The full

sequence of images taken over 48 hours for these cultures via real time microscopy is

provided in Appendix 3.7 to 3.12. The untreated HK control, grown with an i3T3 feeder

layer, exhibited proliferative growth and colony formation at 48 hours. However, following

treatment of HK grown with an i3T3 layer with 0.01% PDMS7‐g‐PEG7, the HK appear to

begin differentiating from 12 hours onwards. Terminally differentiated HK are depicted by

swollen and flattened HK cytoplasm and loss of nucleus (Rheinwald, 1979). When HK grown

with an i3T3 feeder layer were treated with 0.03% PDMS7‐PEG7, cell death occurs from 12

hours but the presence of HK with apoptotic morphology is not evident.


Figure 4.9 – Analysis of HSF morphology following treatment with PDMS7‐g‐PEG7. Images of HSF morphology obtained through real‐time microscopy following treatment with 0.01 and 0.03% PDMS7‐g‐PEG7 for 48 hours. The untreated control includes HSF treated with medium containing no silicone. Representative images are depicted from three replicate experiments, in which each treatment was performed in triplicate (n = 9). Examples of apoptotic cells are indicated by the arrows. Scale bar represents 50 µm.

C h a p t e r 4 . 0 | 79

Figure 4.10 – Analysis of nHSF morphology following treatment with PDMS7‐g‐PEG7. Images of nHSF morphology obtained through real‐time microscopy following treatment with 0.01 and 0.03% PDMS7‐g‐PEG7 for 48 hours. The untreated control includes nHSF treated with medium containing no silicone. Representative images are depicted from three replicate experiments, in which each treatment was performed in triplicate (n = 9). Examples of apoptotic cells are indicated by the arrows. Scale bar represents 50 µm.


Figure 4.11 – Analysis of HK morphology following treatment with PDMS7‐g‐PEG7. Images of HK morphology obtained through real‐time microscopy following treatment with 0.01 and 0.03% PDMS7‐g‐PEG7 for 48 hours. HK were cultured with an i3T3 feeder layer. The untreated control includes HK treated with medium containing no silicone. Representative images are depicted from three replicate experiments, in which each treatment was performed in triplicate (n = 9). Scale bar represents 50 µm.

C h a p t e r 4 . 0 | 81

Figure 4.12 – Analysis of HK morphology, grown without an i3T3 feeder layer, following treatment with PDMS7‐g‐PEG7. Images of HK morphology obtained through real‐time microscopy following treatment with 0.01 and 0.03% PDMS7‐g‐PEG7 for 48 hours. HK were cultured without an i3T3 feeder layer. The untreated control includes HK treated with medium containing no silicone. Examples of apoptotic cells are indicated by the arrows. Representative images are depicted from three replicate experiments, in which each treatment was performed in triplicate (n = 9). Scale bar represents 50 µm.


Following treatment of PDMS7‐g‐PEG7 to HK grown without an i3T3 layer, different effects

compared to those which occurred to HK grown with an i3T3 layer were evident. In

comparing the untreated controls only, it is evident that HK grown without an i3T3 layer

are not proliferative. A large proportion of the HK in these cultures exhibit flattened

cytoplasm and appear more motile, searching for improved growth conditions.

Furthermore, the colonies formed in cultures of HK grown without an i3T3 feeder layer are

not uniform and lacked the usual HK ‘cobblestone’ formation. In addition, contaminating

i3T3 feeder cells, which are present from previous passages, are evident but the effects of

these within the HK cultures is unknown. It can be observed that HK differentiated

following application of 0.01% PDMS7‐g‐PEG7 for 48 hours when the HK were grown

without an i3T3 layer. Moreover, when higher concentrations of PDMS7‐g‐PEG7 were added

(0.03%), HK undergoing apoptosis are clearly evident from 12 hours onwards.

4.3.4 Investigation of the Induction of HSF Apoptosis following Treatment with PDMS7‐g‐

PEG7

The results described in Chapter 3 indicated that treatment of HSF with GP226 and Fraction

IV led to the induction of apoptosis. Since HSF, nHSF and HK also exhibited decreased

proliferation and apoptotic morphology following treatment with PDMS7‐g‐PEG7, further

studies using the Tunel assay were also conducted. Analysis of apoptotic HSF and nHSF

were performed by immunofluorescent microscopy following the Tunel assay and are

depicted in Figure 4.13 and Figure 4.14 respectively. The HSF and nHSF positive controls

exhibit positive immunofluorescence in 100 ± 0% of the cells. Furthermore, no positive

immunofluorescence or background fluorescence is apparent in the negative controls (HSF,

0 ± 0%; nHSF, 0 ± 0%). Similarly, untreated HSF and nHSF demonstrated minimal positive

immunofluorescence (HSF, 0.65 ± 0.42%; nHSF, 0.30 ± 0.30%), indicating that the

population of apoptotic cells in treated cultures was not present before exposure to the

silicone treatments. It is clearly evident by the positive immunofluorescence in Figures 4.13

and 4.14 that apoptotic cells are present in HSF and nHSF cultures treated with PDMS7‐g‐

PEG7 (0.03% PDMS7‐g‐PEG7, HSF, 92.22 ± 3.72%; nHSF, 100 ± 0%). Not surprisingly, the

number of viable cells present following treatment, as illustrated by the blue DAPI

immunofluorescence, decreases as concentration of PDMS7‐g‐PEG7 increases. Additionally,

the proportion of apoptotic cells present following treatment increases as PDMS7‐g‐PEG7

C h a p t e r 4 . 0 | 83

concentration is increased (lowest to highest concentration; HSF, 0 ± 0%, 32.1 ± 9.62%, 92.2

± 3.72%; nHSF, 7.09 ± 2.76%, 44.8 ± 5.48%, 100 ± 0%).

Figure 4.13 ‐ Investigation of apoptosis in HSF following treatment with PDMS7‐g‐PEG7. (A) Immunofluorescent images of Tunel positive HSF following treatment with PDMS7‐g‐PEG7 for 48 hours. Green indicates Tunel positive cells, while blue localises the cell nucleus. Controls displayed include the untreated control (HSF treated with cell culture medium containing no silicone), positive control (including fixed and permeabilised untreated HSF treated with DNAse I) and negative control (including untreated HSF stained with the labelling solution only without enzyme). Represented images are depicted from three replicate experiments (n = 3). Scale bar represents 100 μm. (B) Graph of percentage Tunel positive HSF following treatment with PDMS7‐g‐PEG7. Apoptotic cells were quantified by counting the percentage of positive immunofluorescent cells in three


representative images for each treatment and for each replicate experiment performed (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.

Figure 4.14 ‐ Investigation of apoptosis in nHSF following treatment with PDMS7‐g‐PEG7. (A) Immunofluorescent images of Tunel positive nHSF following treatment with PDMS7‐g‐PEG7 for 48 hours. Green indicates Tunel positive cells, while blue localises the cell nucleus. Controls displayed include the untreated control (HSF treated with cell culture medium containing no silicone), positive control (including fixed and permeabilised untreated HSF treated with DNAse I) and negative control (including untreated HSF stained with the labelling solution only without enzyme). Represented images are depicted from three replicate experiments (n = 3). Scale bar represents 100 μm. (B) Graph of percentage Tunel positive nHSF following treatment with PDMS7‐g‐PEG7. Apoptotic cells were quantified by counting the percentage of positive immunofluorescent cells in three representative images for each treatment and for each replicate experiment performed (n = 9).

C h a p t e r 4 . 0 | 85

Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.

The effect of PDMS7‐g‐PEG7 on HK apoptosis was also investigated using the TUNEL assay

(Figure 4.15). In view of our cell proliferation and real time microscopy results, HK were

cultured with an i3T3 feeder layer for this experiment. Additionally, light microscope

images of the HK cultures were also included for ease of differentiation between HK from

the i3T3 feeder layer cells. Figure 4.15 demonstrates that HK density is markedly

decreased and apoptosis is induced following treatment with 0.01% (11.9 ± 0.82%) and

0.03% (71.2 ± 2.76%) PDMS7‐g‐PEG7. The positive control exhibited positive

immunofluorescence for the assay. However, only 86.6 ± 1.75 % HK in the positive control

demonstrated positive immunofluorescence. This most likely occurred due to stratified

nature of HK proliferation in vitro, along with the inability of the DNase 1 or Tunel enzyme

to reach all layers of cells. Despite this, no positive immunofluorescence for apoptosis was

detected in the negative control (0 ± 0%) and minimal immunofluorescence was present in

the untreated control (2.40 ± 0.62%).


Figure 4.15 ‐ Investigation of apoptosis in HK following treatment with PDMS7‐g‐PEG7. (A) Immunofluorescent images of Tunel positive HK, cultured with an i3T3 feeder layer, following treatment with PDMS7‐g‐PEG7 for 48 hours. Green indicates Tunel positive cells, while blue localises the cell nucleus. Controls displayed include the untreated control (HSF treated with cell culture medium containing no silicone), positive control (including fixed and permeabilised untreated HSF treated with DNAse I) and negative control (including untreated HSF stained with the labelling solution only without enzyme). Represented images are depicted from three replicate experiments (n = 3). Scale bar represents 100 μm. (B) Graph of percentage Tunel positive HK following treatment with PDMS7‐g‐PEG7. Apoptotic cells were quantified by counting the percentage of positive immunofluorescent cells in three representative images for each treatment and for each replicate experiment performed (n = 9). Significant differences (*) are illustrated where P < 0.05 relative to the untreated control. Error bars indicate SEM.

C h a p t e r 4 . 0 | 87

4.4 DISCUSSION

The purpose of the studies reported within this chapter was to evaluate the scar

remediating potential of a variety of fully synthetic silicone species prepared by chemists

within our laboratory. The need for synthesised silicones became apparent when the

amount of pure Fraction IV required for our experiments could not be isolated efficiently

from GP226. The synthesised silicones were of low molecular weight but with a broad

range of properties, including varied amphiphilicity as demonstrated by HLB factor and

structure. Our aim was to find a species of silicone that had either equal or greater effects

on HSF, nHSF and HK than GP226 and Fraction IV, in terms of cell proliferation and

apoptosis. Fraction IV was unable to be included in our experiments investigating the effect

of the synthesised amphiphilic silicones as the amount required for these assays exceeded

that available from fractionation of GP226.

Our experiments indicated that each of the synthesised silicones had differing effects on

HSF and nHSF. However, PDMS7‐g‐PEG7 was the only silicone to decrease both HSF and

nHSF proliferation to levels below those obtained for GP226. Interestingly, PDMS7‐g‐PEG7

had the lowest molecular weight, was the most hydrophilic (as indicated by its high HLB

factor), and was the only silicone oligomer synthesised that consisted of a rake structure.

Although a range of silicones with differing characteristics were evaluated, the exact

structural characteristic of PDMS7‐g‐PEG7 responsible for its biological effects would require

a wide range of rake silicone copolymers with similar molecular weight and HLB range to be

studied to provide further insight.

Upon examination of PDMS7‐g‐PEG7 applied at a wider range of concentrations to HSF and

nHSF, a dose dependant effect leading to decreased proliferation was evident.

Interestingly, dose dependant decreases in cell proliferation were also demonstrated when

PDMS7‐g‐PEG7 was applied to HK, but these decreases were not observed when the

experiment was repeated using HK grown without an i3T3 feeder layer. It was initially

unclear whether the decreases in keratinocyte proliferation observed following PDMS7‐g‐

PEG7 treatment were an actual measure of decreased keratinocyte proliferation or an

effect arising from the fibroblast feeder layer beneath the HK cells. It has been widely

reported that 3T3 fibroblasts have a protective role in HK culture (Rheinwald and Green,

1975). Furthermore, their presence, along with the growth factors they produce, are

essential for the proliferation and subcultivation of HK (Green, 1991; Green et al., 1977;


Rheinwald and Green, 1975). Indeed, comparisons of the untreated controls in the Cyquant

experiments revealed that HK grown without a feeder layer exhibited much lower levels of

proliferation than when HK were cultured with a feeder layer. Our data suggests that the

HK grown without an i3T3 feeder layer were not actively proliferating at the beginning of

the experiments. Therefore, assessing PDMS7‐g‐PEG7 on these cells in this situation

indicated results that do not represent the true effects of the silicone.

The decreases in cell proliferation observed following application of PDMS7‐g‐PEG7 to HSF,

nHSF and HK led us to hypothesise that apoptosis was occurring during treatment with

these silicones. The induction of apoptosis in cultures of HSF, nHSF and HK following

treatment with PDMS7‐g‐PEG7 was confirmed through analyses of cell morphology and the

Tunel assay. Although apoptosis in HSF, nHSF and HK following exposure to PDMS7‐g‐PEG7

was not always evident in our studies of cell morphology, our Tunel experiments

demonstrated a dose dependant rise in the proportion of apoptotic cells as the

concentration of PDMS7‐g‐PEG7 increased. Interestingly, HK undergoing apoptosis could not

be distinguished in our cell morphology studies when HK were cultured with an i3T3 feeder

layer while treated with PDMS7‐g‐PEG7. However, Tunel assays assessing these same

culture conditions revealed the presence of HK undergoing apoptosis. Why apoptosis was

observed in our cell morphology studies with PDMS7‐g‐PEG7 applied to HK cultures grown

without a feeder layer but not in HK cultures grown with a feeder layer is not immediately

apparent. Nevertheless, as mentioned above, 3T3s have a protective role in HK culture and

this may explain why apoptotic HK were not visually apparent following treatment with

PDMS7‐g‐PEG7 in HK cultures grown with an i3T3 feeder layer (Green et al., 1977;

Rheinwald and Green, 1975).

Apoptosis as a cellular function has been widely investigated and is marked by cellular

shrinking, condensation and margination of the chromatin, as well as ruffling of the plasma

membrane with eventual breakup of the cell in apoptotic bodies (Taylor et al., 2008). Our

investigations of real‐time and immunofluorescent microscopy indicate that PDMS7‐g‐PEG7

induces apoptosis in cultures of HSF, nHSF and HK. Despite these results, it is extremely

important to be aware of artifacts that can occur in tissue culture models when attempting

to define mechanisms and events leading to cell death (Bonfoco et al., 1995). The

reasoning behind this is that the distinction between apoptosis and necrosis in vitro can be

C h a p t e r 4 . 0 | 89

confused because there is a lack of scavenging cells, thus the phagocytic step after

apoptosis may not occur. Instead, the apoptotic cell can eventually undergo secondary

necrosis and rupture its contents into the surrounding medium (Duke and Cohen, 1986).

This form of cell death was observed in our real‐time microscopy results with the fibroblast

cultures treated with 0.03% PDMS7‐g‐PEG7. That is, the cell contents were ruptured

between the 24 and 48 hour assessment time points, following the observation of

apoptosis at 12 hours.

It is important to reiterate that the experiments performed in this doctoral study have

utilised 2‐dimensional (2D) in vitro cell cultures and may not truly represent effects that

would occur in vivo. Furthermore, as noted earlier in the literature review (Chapter 1.0),

the in vitro effects of silicone are not likely to be observed in vivo because silicone oils,

which are usually hydrophobic, have difficulty penetrating the stratum corneum (Chang et

al., 1995). The barrier properties of skin are determined by the stratum corneum, the

outermost lipidic layer that interphases with the aqueous medium beneath it (Mitragotri,

2003). The transport of lipophilic chemicals, such as hydrophobic silicone oils, through the

stratum corneum is difficult as these compounds must travel through either an intercellular

route, moving in between the cells, or transfer through the cells, a trancellular route, and

directly into an aqueous medium (Figure 4.16; (Junginger et al., 1994). However, the

silicones investigated in our studies, including GP226, Fraction IV and PDMS7‐PEG7, can

actually facilitate this permeation as they are different to most silicone oils. In particular,

they are amphiphilic, containing both hydrophobic and hydrophilic qualities.

Figure 4.16 – Schematic of the stratum corneum and possible permeation pathways. (Junginger et al., 1994)


It has been demonstrated previously that silicone species with low molecular weight and

amphiphilic properties can move through the stratum corneum (Sanchez et al., 2005;

Shigeki et al., 1999). Indeed, in other studies performed in our laboratory, it has been

shown that GP226 traverses the stratum corneum at approximately 14 ± 2 hours at 32°C

(Dickfos, 2008; Gardoni, 2007). The maximum diffusion coefficient for GP226 was found to

be 6.9 ± 0.25 x 10‐9 cm2s‐1 at 32°C. Furthermore, the breakthrough time for PDMS7‐g‐PEG7

occurred at 4 ± 0.5 hours with the diffusion coefficient through the stratum corneum at

32°C being 1.7 ± 0.2 x 10‐8 cm2s‐1 at 32°C. Taken together, this suggests that the unique

combination of amphiphilic and low molecular weight properties of GP226, Fraction IV and

PDMS7‐g‐PEG7 facilitate permeation through the stratum corneum. The mode of transport

in which these silicones traverse the stratum corneum, including the localisation and

movement of silicones within the cells as would occur in trancellular transport, has also

been investigated. The methods investigated involved the incorporation of a FITC tag onto

the silicone structure as well as encasing the FITC molecule into a silicone micelle; these

two methods proved troublesome, however, especially as the FITC molecule altered the

molecular weight of the silicone treatments, therefore modifying its effect on the cells

(Keddie, D. and Farrugia, B. pers. comm.).

In conclusion, the experiments described in this chapter were directed at identifying a

silicone species that had an equivalent or greater effect on dermal fibroblasts and

keratinocytes than the previously reported results observed with GP226 and Fraction IV. It

is evident through the investigations reported here that PDMS7‐g‐PEG7 is a suitable

substitute for GP266 and Fraction IV as it elicited significant and dose‐dependent decreases

in cell proliferation, as well as an induction of apoptosis, following application to cells in

culture.

C h a p t e r 5 . 0 | 91

CHAPTER 5.0

INVESTIGATIONS INTO THE EFFECT OF AMPHIPHILIC

SILICONE‐PEG COPOLYMERS AT THE GENOMIC LEVEL

5.1 INTRODUCTION

As noted earlier, the use of silicone gel sheets for hypertrophic scarring has been well

documented in the literature. Despite their widespread use and the considerable amount

of research worldwide, knowledge regarding their mechanism of action still remains

limited. It was determined by Quinn in 1987 that the action of silicone gel sheets did not

result from pressure, temperature or oxygen tension but must involve a chemical factor.

The results in Chapters 3.0 and 4.0 of this thesis demonstrated that low molecular weight

and amphiphilic silicones, such as GP226, Fraction IV and PDMS7‐g‐PEG7, significantly affect

cellular functions in fibroblasts and keratinocytes. Indeed, the data presented in Chapters

3.0 and 4.0 have illustrated that cell proliferation and viability is significantly and dose‐

dependently reduced by these chemical species. Further, this is likely due to apoptosis

being induced in response to treatment with the silicones. In addition, other studies

undertaken within our laboratory have demonstrated that these low molecular weight and

amphiphilic silicone species are capable of penetrating the stratum corneum (Dickfos,

2008; Gardoni, 2007). Taken together, these results have led us to propose that the silicone

species are able to interact with dermal fibroblast cells and affect key cellular mechanisms

pertinent to scarring, such as cell proliferation and apoptosis, once they have penetrated

the stratum corneum. Nevertheless, the mechanisms occurring in fibroblasts undergoing

apoptosis during treatment with the silicones requires investigation.

It is well recognised that changes in gene expression play major roles in cellular biology

(Risch and Merikangas, 1996). Therefore, changes in gene expression following exposure of

cells to the silicone treatments was pursued; this may increase our understanding of the

mechanisms responsible for the induction of apoptosis following treatment and also

provide insights into changes in other cellular processes that are affected by silicone

treatment. While Paddock et al., (2003) and Wu et al., (2004) have performed gene

microarray studies comparing the differences between hypertrophic scars, normal scars

and uninjured skin, global gene expression studies examining differences between

abnormal scars treated with and without silicone have not been reported in the literature.

I n v e s t i g a t i o n s a t t h e G e n o m i c l e v e l

In this chapter, the mechanism of silicone action following application of GP226, Fraction IV

and PDMS7‐g‐PEG7 to dermal fibroblasts was investigated through differential gene

expression studies using gene microarray, apoptosis superarray and qRT‐PCR approaches.

Although the results in Chapters 3.0 and 4.0 demonstrated that GP226, Fraction IV and

PDMS7‐g‐PEG7 affect proliferation and induce apoptosis in both fibroblasts and

keratinocytes, the studies reported in this chapter focus only on differential gene

expression in dermal fibroblasts following treatment with the silicones.



Pure samples of Fraction IV from GP226 were separated and obtained using PSEC and

HPLC. Please refer to section 2.2.1 for further details relating to the fractionation process.

In all experiments performed, an equivalent amount of Fraction IV to what was originally

present in the unfractionated GP226 mixture, was assayed. Hence, a proportionate dilution

of Fraction IV (0.1%), compared to the unpurified GP226 (0.1%), was prepared in cell

culture medium and applied to the cultured cells. For further details relating to the

treatment dilutions, please see Table 2.1.


For full specifications of the synthesis of PDMS‐PEG oligomers used in this chapter, please

see section 2.2.2 as well as Appendix 1. PDMS7‐g‐PEG7 was prepared at a concentration of

0.01% and 0.03% in cell culture medium and applied to the cultured cells.


HSF and nHSF were independently and routinely cultured for use in the experiments

performed in this chapter. Please refer to section 2.3.1 for more specific cell culture details.

5.2.4 RNA Extraction

Total RNA was extracted from treated cell cultures using Qiagen RNeasy Mini kit. Briefly,

cultures of HSF and nHSF were prepared in T25cm2 cell culture flasks and cell culture

medium containing GP226 (0.1%), Fraction IV (0.1%) and PDMS7‐g‐PEG7 (0.01 and 0.03%)

were added 24 hours later. Cell cultures treated with GP226 and Fraction IV were

incubated for 48 hours while cell cultures with PDMS7‐g‐PEG7 were incubated for either 6

hours or 48 hours. Untreated controls, in which the cell cultures were treated with cell

C h a p t e r 5 . 0 | 93

culture medium containing no silicone for 6 or 48 hours, were also prepared. Following

treatment, RNA extractions were performed using the Qiagen RNeasy Mini kit. For more

specific details relating to this methodology, please see section 2.5.1.

5.2.5 Microarray

The HumanHT‐12 v3 Expression BeadChip microarray was used to determine differentially

expressed genes in cultures of HSF and nHSF following treatment with and without GP226

(0.1%) and Fraction IV (0.1%). The HumanHT‐12 v3 Expression BeadChip microarray was

performed with total RNA extracted from treated and untreated cell cultures by Katie

Nones at the Special Research Facility Microarray Service within the Institute of Molecular

Biosciences, University of Queensland. Following microarray, all data was kindly analysed

by Daniel Haustead, University of Western Australia, and Jacqui McGovern, Queensland

University of Technology. Please refer to sections 2.5.2 and 2.5.3 for more details relating

to the microarray process and the data analysis.

5.2.6 Gene Ontology, Canonical Pathway and Functional Network Analysis

Gene ontology, canonical pathway and functional network analyses were undertaken using

IPA tools (Ingenuity Systems, www.ingenuity.com). Please refer to section 2.5.4 for further

details relating to this analysis.

5.2.7 Super Arrays

In addition to the microarrays performed above, quantitative reverse transcription‐

polymerase chain reaction (qRT‐PCR) superarrays were also performed on HSF and nHSF

cultures exposed to PDMS7‐g‐PEG7 (0.01% and 0.03%). More specifically, arrays analysing

the human apoptosis pathway, were employed. For more specific details describing the

methodology required for cDNA synthesis and the subsequent superarray process, please

see sections 2.5.6 and 2.5.5, respectively.

5.2.8 Confirmation of Differential Gene Expression using Quantitative RT‐PCR

For full details of these protocols, please refer to chapter 2, sections 2.6.1 to 2.6.5. qRT‐PCR

was used to validate the microarray and superarray expression data by measuring absolute

expression levels of selected genes of interest.


5.2.9 Standard PCR Conditions

All standard PCR reactions were performed using the Platinum Taq PCR kit. Please refer to

section 2.6.1 for details describing the reaction and temperature cycling conditions. PCR

amplicon size was then confirmed using agarose gel electrophoresis and ethidium

bromide/UV visualization.

5.2.10 Primer Design

Primers were designed using Primer‐BLAST software, a web‐based primer designing tool

available through the National Center for Biotechnology Information

(www.ncbi.nlm.nih.gov/tools.primer‐blast/). Please refer to section 2.6.2 for specific details

used in primer design.

5.2.11 Reverse Transcription (RT) for qRT‐PCR

First strand cDNA synthesis was performed using SuperscriptTM III Reverse Transcriptase

(Invitrogen) with total RNA extracted from HSF and nHSF, either untreated or treated with

GP226, Fraction IV and PDMS7‐g‐PEG7. Section 2.6.3 provides more specific details relating

to this method.

5.2.12 PCR and Amplicon purification

PCR was performed as described previously (Section 5.2.9) using Platinum Taq and

sequence specific primers for each transcript to be analysed by qRT‐PCR. Please refer to

table 2.2 for the sequence specific primers used in these experiments and to section 2.6.4

for more details relating to this methodology.

5.2.13 qRT‐PCR

Microarray and superarray data was validated by using qRT‐PCR to measure absolute

expression levels of the selected genes of interest identified through microarray and

superarray studies. The primers used in qRT‐PCR were designed using Primer‐BLAST

(www.ncbi.nlm.nih.gov/tools/primer‐blast/) and are outlined in Table 2.2 and section

5.2.10. Standard curves, generated by 10‐fold serial dilutions of purified PCR target

amplicons and covering at least 7 logs of amplicon copy number, were used to achieve

absolute quantitation. For further details relating to the qRT‐PCR methodology and

standard conditions used, please refer to section 2.6.5. Expression of target genes for each

C h a p t e r 5 . 0 | 95

treated sample was normalized to 18S rRNA and compared to normalized untreated

samples.


To obtain replicate biological samples, qRT‐PCR experiments were repeated three times

using HSF and nHSF cells from three different patients. Within each experiment, each

variable was tested in triplicate. Data was expressed as a mean ± standard error of the

mean (SEM) and declared significant with analysis of variance followed by a pairwise t‐test

post hoc. Probability (P) values < 0.05 were considered significant.

5.3 RESULTS

5.3.1 Identification of Differential Gene Expression

The HumanHT‐12 v3 Expression BeadChip microarray was used to determine differentially

expressed genes in cultures of HSF and nHSF following treatment with and without GP226

and Fraction IV. Replicate experiments for the untreated control and each treatment type

were unable to be performed, meaning that statistical analysis could not be undertaken.

Instead the array was used as a screening technique to select genes of interest relating to

apoptosis. In order to identify silicone‐induced changes in gene expression, results were

first compared between each treatment and the untreated control. Table 5.1 depicts the

numbers of genes that were differentially expressed between the treated and untreated

controls in both the microarray and superarray studies. Comparisons were also made

between the different cell types, HSF and nHSF (Table 5.1). Only differentially expressed

genes with a fold change of ± 2.0 or greater within the microarray analysis results were

included to keep the data sets at a manageable size and to also ensure critical genes were

included. As is illustrated in Table 5.1, over 500 genes were found to be increased and over

600 genes were found to be decreased in HSF and nHSF following treatment with GP226

and Fraction IV. Less than 100 genes were found to be differentially expressed between

untreated HSF and nHSF in the microarray analyses.


Comparisons Increased Decreased Microarray (25000 genes)

HSF vs nHSF 91 8 HSF Un vs 0.1% GP226 577 687

HSF Un vs 0.1% Fraction IV 556 676

nHSF Un vs 0.1% Fraction IV 522 607

Table 5.1 – Summary of differential gene expression obtained through microarray analyses. Number of probe sets differentially regulated in the microarray analyses of HSF and nHSF following exposure to GP226 and Fraction IV, as well as between the untreated controls.

5.3.2 Gene Ontology and Functional Analysis of Microarray

The microarray investigations, examining the effect of GP226 and Fraction IV on HSF and

nHSF, produced large data sets of differentially regulated genes within fibroblasts in

response to the application of silicone. The Probe identification (ID), gene symbols,

GenBank accession numbers and relative fold changes for the differentially expressed

genes identified between HSF and nHSF, as well as in HSF following treatment with GP226

and Fraction IV, and nHSF in response to GP226, are depicted in A4.1, A4.2, A4.3 and A4.4,

respectively, within Appendix 4. It was evident that the differentially regulated genes,

especially in HSF and nHSF exposed to silicone, were from a diverse range of gene families,

with each having a variety of functions. In order to more fully appreciate the biological

pathways affected in fibroblasts in response to silicone treatment, the IPA knowledge base

was utilised. Data sets of the genes identified as being at least ± 2‐fold differentially

expressed between untreated fibroblast samples and in response to GP226 and Fraction IV

were uploaded separately into the application.

Functional analysis identified a number of biological functions that were the most closely

related to the differentially regulated genes. Molecular and cellular functions that showed

a P‐value below 0.05 and encompassed at least 5 genes of interest were considered

significant. Table 5.2 depicts the most significant molecular and cellular functions in the

data set of differentially expressed genes in HSF treated with GP226, and includes

processes involved in cell cycle, cell death, cellular growth and proliferation, cellular

movement and cellular development. Table A4.5, in Appendix 4, depicts the most

significant molecular and cellular functions in HSF and nHSF treated with Fraction IV. These

IPA analyses revealed that processes involved in the cell cycle and cell death functions were

the most significantly affected functional category, containing 205 and 302 focus genes

respectively.

C h a p t e r 5 . 0 | 97

Molecular and Cellular Functions Significance P‐value Focus Genes Cell Cycle 5.10E‐36 ‐ 1.94E‐04 205 Cell Death 4.49E‐20 ‐ 1.91E‐04 302 Cellular Growth and Proliferation 2.73E‐19 ‐ 1.92E‐04 339 Cellular Movement 6.03E‐18 ‐ 1.69E‐04 198 Cellular Development 2.99E‐15 ‐ 1.94E‐04 255

Table 5.2 Ontology of differentially expressed genes identified through microarray analysis in HSF exposed to GP226. Molecular and cellular functions identified to be most significantly related to the dataset of differentially regulated genes identified through microarray analysis in HSF treated with HSF for 48 hours.

The cellular and molecular functions that are described in Table 5.2 are clearly quite broad

classifications. Therefore IPA was used to further categorise the genes to more specific

functional roles relevant to each classification. As our studies were focused on apoptosis,

genes with more specific functions within the cell cycle, cell death and cellular growth and

proliferation categories were investigated. Table 5.3 demonstrated that a number of genes

in HSF treated with GP226 were significantly associated with these biological process

categories, including arrest in cell division process, arrest in cell cycle progression,

apoptosis and proliferation of fibroblasts. These genes were named focus genes. Section

A4.7, in Appendix 4, depicts the focus genes significantly associated with these biological

process categories in HSF and nHSF exposed to Fraction IV. Through these analyses, a

number of genes, including Collagen type I, alpha I (COL1A1), IL8, Apoptosis‐inducing

factor, mitochondrion‐associated, 2 (AIFM2), Neutral sphingomyelinase activation

associated factor (NSMAF), Insulin‐like growth factor 2 receptor (IGF2R) and SMAD family

member 7 (SMAD7), were associated with biological processes relevant to Cell Cycle, Cell

Death and Cellular Growth and Proliferation.

The disregulation of COL1A1 identified in the ‘arrest in cell division process’ was particularly

interesting, since collagen is one of the main forms of protein synthesised by dermal

fibroblasts. This result caused us to manually examine the initial microarray data obtained

to investigate the differential expression of other collagen types within each HSF and nHSF

sample treated with GP226 and Fraction IV. Upon manual inspection, many types of

collagen were found to be down‐regulated in HSF and nHSF exposed to GP226 and Fraction

IV (Table 5.4). Significantly, COL1A1 as well as Collagen type III, alpha I (COL3A1), the main

types of collagen within the skin, were both found to be down‐regulated following silicone

treatment when compared to the untreated controls (Uitto et al., 1981).


Sub‐Category Focus Genes

CELL CYCLE

Arrest in cell

division process

P‐value: 9.91E‐

19

ATF3, AURKA, AURKB, BARD1, BHLHE40, BIRC5, BMP2, BTG2, BUB1, BUB3, BUB1B, C11ORF82,

CAV1, CCNA2, CCNB1, CDC20, CDC25C, CDH13, CDK1, CDK2, CDKN3, CDKN1A, CDKN2C, CEBPB,

CEBPD, CENPE, CENPI, CHKA, CKAP2, CKS2, CKS1B, CLIP1, COL1A1, CYP1B1, CYP26B1, CYR61,

DCN, DDIT3, DLGAP5, EGR1, FANCD2, FBXO5, FOXM1, GADD45A, GAL, GAS1, GAS7, GDF15,

HIPK2, HOXA10, ID2, ID3, IFNB1, IGFBP5, IL6, IL8, ILK, IRF7, ITGA2, KIFC1, KRT19, MAD2L1,

MAGED1, MCM7, MET, NEK7, NFKBIA, NUF2, PITX2, PKD2 (includes EG:5311), PLAU, PLAUR,

PLK1, PPAP2C, PPP1R15A, PTTG1, RASSF1, RRM2B, SGOL1, SKP2, SMAD3, SMAD7, SPARC,

THBS1, TMPO, TP53INP1, TXNIP, TYMS, WEE1, ZWINT

Arrest in cell

cycle

progression

P‐value: 1.00 E‐

07

ATF3, AURKA, BIRC5, CAV1, CCNA2, CCNB1, CDK2, CDKN3, CDKN1A, CDKN2C, CKAP2, CKS2,

COL1A1, FANCD2, GADD45A, GAL, GAS1, GAS7, HIPK2, HOXA10, ID2, IFNB1, IL6, IL8, ILK, IRF7,

ITGA2, KRT19, MAD2L1, MAGED1, NFKBIA, PKD2, PLK1, PPP1R15A, RASSF1, SKP2, SMAD3,

SPARC, THBS1, TP53INP1, TYMS

CELL DEATH

apoptosis

P‐value: 4.98E‐

16

ABCC1, ADI1, ADM, AGT, AIFM2, ANGPT1, ARNT2, ASNS, ATF3, AURKA, B4GALT5, BARD1, BAX,

BCL2L12, BDKRB1, BEX2, BHLHE40, BIRC3, BIRC5, BMP2, BTG2, C11ORF82, CAST, CAV1, CBX5,

CCL5, CCNA2, CCNB1, CD14, CD55, CDC20, CDC45, CDC25C, CDC42EP3, CDCA2, CDK1, CDK2,

CDKN1A, CDKN2C, CEBPB, CEBPD, CENPF, CHKA, CKAP2, CLN3, CLN8, CNN2, CRABP2, CTGF,

CTH, CTSB, CTSD, CXCL2, CXCL12, CYBA, CYLD, CYP1B1, CYP26B1, CYR61, DAP, DCN, DDIT3,

DDIT4, DDX58, DNMT1, DUT, EEF1E1, EGR1, EPAS1, ETS2, F3, F2R, FAIM, FBXO32, FEN1, FOSL1,

FTH1, G6PD, GADD45A, GADD45G, GAL, GALNT5, GAS1, GCLC, GDF15, GLIPR1, GNA13,

GREM1, GRN, GSR, HIF1A, HIPK2, HIST1H1C, HK1, HK2, HMGA1, HMGB1L1, HMMR, HMOX1,

HOXA13, HSF2, HSPA2, HSPB8, ID1, ID2, ID3, IFI6, IFIH1, IFNB1, IGF2R, IGFBP3, IGFBP5, IKBIP,

IL6, IL8, ILK, IRAK1, IRS2, ITGA2, ITGB3, ITPR3, JUP, KIF14, KIT, LAMA4, LIG1, LMNB1, MAD2L1,

MAFB, MAGED1, MCM2, MCM10, MET, MME, MMP2, MMP3, MMP1 (includes EG:4312), MX1,

NAMPT, NCAM1, NCAPG2, NEDD9, NEK7, NFKB2, NFKBIA, NFKBIZ, NPC1, NPTX1, NRGN,

NSMAF, OAS1, OSGIN1, PDGFRB, PGF, PHLDA1, PHLDA2, PKMYT1, PLAT, PLAU, PLAUR, PLK1,

PMAIP1, PPP1R15A, PTGIS, PTGS2, PTPRE, PTTG1, RAD21, RASSF1, RCAN2, RHOB, RND3,

RRM2B, RTN4, S1PR3, SAT1, SDC4, SEMA3A, SFRP1, SGCG, SGK1, SKP2, SMAD3, SMAD7,

SOAT1, SOCS1, SOD2, SPARC, SPP1, SPRY2, SRPK2, SRXN1, STIL, STMN1, STX1A, TACC3, TCF4,

TGM2, THBS1, THRA, TK1, TNFAIP3, TNFRSF14, TNFRSF19, TNFRSF21, TNFRSF10A, TNFRSF10B,

TNFRSF11B, TNFSF9, TOP2A, TP53INP1, TPD52L1, TRIB1, TRIB3, TTK, TXNIP, TYMP, TYMS,

UACA, UBR4, UNC5B, VCL, VEGFC, WEE1, WISP1, WNT2, WNT5A, XAF1, ZFYVE16

CELLULAR

GROWTH AND

PROLIFERATION

Proliferation of

fibroblasts

P‐value:

2.28E‐07

AGT, BAX, CAV1, CCNA2, CCNF, CDK2, CDKN1A, CEBPB, DCN, GRN, HMMR, IL6, ILK, PMAIP1,

SKP2, SMAD3, SMAD7, SOD2, SPARC, SPRY2, TGIF1, TNFRSF10B, TNFRSF11B, TXNIP, WISP1,

ZMIZ1

Table 5.3 – Focus genes identified through microarray analysis to be differentially expressed in HSF exposed to GP226. Focus genes are those genes found to be differentially regulated in HSF exposed to GP226 for 48 hours and relevant to the biological processes of Cell Cycle, Cell death and Cellular Growth and Proliferation.

C h a p t e r 5 . 0 | 99

Gene HSF Un vs HSF

GP226 HSF Un vs HSF Fraction IV

nHSF Un vs nHSF Fraction IV

COL1A1 ‐7.00 ‐6.10 ‐8.12

COL1A2 ‐3.45 ‐3.37 ‐4.02

COL3A1 ‐22.29 ‐21.79 ‐20.82

COL5A1 ‐3.67 ‐2.80 ‐4.26

COL5A2 ‐4.72 ‐5.76 ‐4.94

COL5A3 ‐2.11 ‐2.21 ‐2.36

COL8A1 ‐4.03 ‐2.80 ‐4.27

COL11A1 ‐6.22 ‐2.04 ‐11.63

COL12A1 ‐2.09 ‐2.22 ‐2.71

COL13A1 2.31 2.43 2.31

COL15A1 2.55 3.68 2.83

Table 5.4 – Expression of collagen genes identified through microarray analysis to be differentially regulated in HSF exposed to GP226. Results are expressed as gene expression fold‐change in HSF and nHSF treated with GP226 and Fraction IV for 48 hours, compared to the untreated control.

5.3.3 Differentially Expressed Genes as depicted by Superarray

In addition to the microarray studies, qRT‐PCR superarrays were also performed with HSF

and nHSF cultures exposed to PDMS7‐g‐PEG7. While the microarray analyses investigated

the expression of genes over the whole genome and produced large data sets, the

superarray analyses only examined genes relating to the apoptosis pathway. Replicate

experiments for the untreated control and each treatment type in the superarray

experiments were unable to be performed hence statistical analysis could not be

undertaken. Nevertheless, meaningful data was obtained, with Table 5.5 depicting the

number of genes that were differentially expressed in the Superarray analyses of HSF and

nHSF following treatment with PDMS7‐g‐PEG7. These investigations were initially

undertaken on HSF exposed to 0.01% and 0.03% to observe which concentration induced

the most changes in gene expression. The concentration of 0.03% PDMS7‐g‐PEG7 was found

to stimulate the greatest change in gene expression (0.01% PDMS7‐g‐PEG7, 9 genes

increased, 7 genes increased; 0.03% PDMS7‐g‐PEG7, 16 genes increased, 12 genes

decreased). In view of this, further studies using 0.03% PDMS7‐g‐PEG7 were undertaken to

evaluate the effect of PDMS7‐g‐PEG7 on both HSF and nHSF for 6 and 48 hours. The two

timepoints of 6 and 48 hours were examined as differential changes in gene expression are

known to occur at varying times during chemical insult (Bobashev et al., 2002).


Comparisons Increased Decreased

Superarray

(84 genes)

HSF vs nHSF 3 3

HSF Un vs 0.01% PDMS7‐g‐PEG7 (48 hours) 9 7


nHSF Un vs 0.03% PDMS7‐g‐PEG7 (48 hours) 16 12


nHSF Un vs 0.03% PDMS7‐g‐PEG7 (48 hours) 28 4

Table 5.5 – Summary of differential gene expression obtained through superarray analyses. Number of probe sets differentially regulated in the superarray analyses of HSF and nHSF following exposure to PDMS7‐g‐PEG7 and between the untreated controls.

The resulting lists of genes and the fold‐change between treated and untreated samples, as

depicted in Table A4.7 in Appendix 4, were manually sorted and genes that were

differentially expressed between three or more samples selected for validation. While

genes with a fold change of ± 2.0 were included in microarray analyses, differentially

expressed genes with a fold change of ± 1.8 or greater within the superarray results were

included as it is a smaller array and to ensure all critical genes were included. Table 5.6 lists

the genes that were significantly expressed above the ± 1.8‐fold change threshold in three

or more samples. Interestingly, in most cases, genes, including Cluster of Differentiation 70

(CD70), Caspase 8‐like apoptosis regulator (CFLAR), Death‐associated protein kinase 1

(DAPK1), TNF receptor superfamily, member 6 (FAS) Lymphotoxin alpha (LTA), Myeloid cell

leukaemia sequence 1 (MCL1), Tumour necrosis factor alpha (TNF), Tumour necrosis factor

receptor superfamily member 10B (TNFRSF10B), TNF receptor‐associated factor 2 (TRAF2)

and TNF receptor‐associated factor 3 (TRAF3), were not ubiquitously up‐ or down‐regulated

in response to PDMS7‐g‐PEG7 treatment. Furthermore, the changes in gene expression

were not always similar between the samples from the 6 and 48 hour timepoints. Taken

together, along with the results of the microarray analyses, it is evident that many cellular

processes, including cell death and collagen synthesis pathways, are affected in HSF and

nHSF following exposure to the silicone treatments.

C h a p t e r 5 . 0 | 101

Gene

HSF vs

nHSF

HSF

Untreated

vs 0.01%

PDMS7‐g‐

PEG7

HSF Untreated vs 0.03%

PDMS7‐g‐PEG7

nHSF Untreated vs

0.03% PDMS7‐g‐PEG7

48 hours 48 hours 6 hours 48 hours 6 hours 48 hours

APAF1 ‐1.24 ‐1.67 ‐2.09 ‐2.06 ‐1.28 ‐2.35

BIK ‐1.08 1.03 ‐2.28 ‐2.70 ‐3.60 1.20

CD70 2.86 ‐3.59 ‐8.36 ‐2.76 2.72 3.67

CFLAR 1.01 11.02 ‐2.45 22.53 ‐1.16 1.16

CIDEA ‐2.14 1.20 ‐4.28 ‐1.88 ‐3.80 ‐1.77

DAPK1 ‐1.08 1.06 5.38 28.71 2.93 ‐1.28

FAS ‐1.07 ‐1.49 5.29 1.98 1.18 2.53

HRK 1.15 1.60 ‐1.63 8.62 2.43 134.80

LTA 1.98 1.07 ‐1.32 ‐4.84 2.00 20.79

MCL1 1.00 ‐42.39 ‐9.46 1.41 1.52 2.23

TNF ‐1.07 ‐1.06 ‐1.42 ‐18.13 3.64 ‐12.20

TNFRSF9 ‐1.06 1.13 ‐1.37 ‐98.38 ‐18.18 ‐139.88

TNFRSF10B ‐1.30 1.19 ‐2.01 2.48 2.61 7.83

TRAF2 ‐1.60 ‐2.63 ‐23.44 2.40 1.19 4.82

TRAF3 ‐1.38 ‐2.86 ‐10.49 1.53 1.69 2.88

Table 5.6 – Focus genes identified through superarray analysis to be differentially expressed in HSF and nHSF treated with PDMS7‐g‐PEG7. Focus genes are those genes found to be associated with apoptosis following superarray analysis of HSF and nHSF treated with PDMS7‐g‐PEG7 for 48 hours. Results are expressed as gene expression fold‐change in HSF and nHSF treated with PDMS7‐g‐PEG7 compared to the untreated control or in nHSF when compared to HSF. Genes highlighted in red represent those that were up‐regulated above the +1.8 fold change threshold while genes highlighted in blue represent those that were down‐regulated and below the ‐1.8 fold change threshold.

5.3.4 qRT‐PCR Validation of Differentially Expressed Genes

Several genes deemed biologically interesting because of their differential expression

following exposure to different silicone treatments and/or their relevance to apoptosis and

the specific cell and molecular functions of cell cycle, cell death and cellular growth and

proliferation, and collagen synthesis, were chosen for validation by qRT‐PCR. Total RNA was

isolated from cells, treated with and without silicone, from three different patients in order

to gain biologically relevant data. The differential expression of 16 transcripts identified by

microarray and superarray analysis were validated by qRT‐PCR, using 18S as an

normalisation control. The transcripts of 9 up‐regulated genes to be validated by qRT‐PCR

are illustrated in Table 5.7. Additionally, the transcripts of 5 down‐regulated genes to be

validated by qRT‐PCR are described in Table 5.8. Furthermore, the genes for TGFβ1, an


important cytokine in wound healing and the collagen synthesis pathway, as well as αSMA,

an important molecule in differentiating myofibroblasts, were also included in qRT‐PCR

experiments (Table 5.9) (Hinz et al., 2001; Varga et al., 1987).

Gene Symbol Gene Name Accession NumberMicroarray AIFM2

Homo sapiens apoptosis‐inducing factor, mitochondrion‐associated, 2, nuclear gene encoding mitochondrial protein, mRNA.

NM_032797.4

IGF2R

Homo sapiens insulin‐like growth factor 2 receptor, mRNA.

NM_000876.2

IL8 Homo sapiens interleukin 8, mRNA.

NM_000584.2

NSMAF

Homo sapiens neutral sphingomyelinase (N‐SMase) activation associated factor, mRNA.

NM_003580.2

SMAD7

Homo sapiens SMAD family member 7, mRNA. NM_005904.2

Superarray DAPK1

Death‐associated protein kinase 1

NM_004938

FAS

Fas (TNF receptor superfamily, member 6) NM_000043

TNF

Tumour necrosis factor alpha NM_000594

TNFRSF10B

Tumour necrosis factor receptor superfamily, member 10b

NM_003842

Table 5.7 – Up‐regulated Focus Genes selected for validation by qRT‐PCR.

Gene Symbol Gene Name Accession NumberMicroarray COL1A1 Homo sapiens collagen, type I, alpha 1 NM_000088.3

COL3A1 Homo sapiens collagen, type III, alpha 1 NM_000090.3

Superarray CFLAR CASP8 and FADD‐like apoptosis regulator NM_003879

TRAF2

TNF receptor‐associated factor 2 NM_021138

TRAF3

TNF receptor‐associated factor 3 NM_003300

Table 5.8 – Down‐regulated Focus Genes selected for validation by qRT‐PCR.

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Gene Symbol Gene Name Accession NumberMiscellaneous

aSMA ACTA2 actin, alpha 2, smooth muscle, aorta NM_001613.2

TGFβ1 Homo sapiens transforming growth factor, beta 1

NM_000660.4

Housekeeping gene

18S

18S ribosomal RNA NR_003286.2

Table 5.9 – Miscellaneous Focus Genes selected for validation by qRT‐PCR.

To fully validate the microarray and superarray data, relative mRNA levels were assessed

for the selected gene transcripts in samples from HSF and nHSF treated with and without

GP226 and Fraction IV at 48 hours, as well as PDMS7‐g‐PEG7 at 6 and 48 hours. Due to the

limited quantities of GP226 and Fraction IV available, the responses of differentially

regulated genes in HSF and nHSF could only be assessed at 48 hours. Target gene

expression for each sample of HSF and nHSF treated with GP226, Fraction IV or PDMS7‐g‐

PEG7 was normalised to 18S rRNA before being expressed as fold change relative to the

untreated samples. Figures 5.1 and 5.2 depict the expression of respective up‐regulated

and down‐regulated focus genes in HSF and nHSF following treatment with GP226 and

Fraction IV for 48 hours. Figures 5.3 and 5.4 depict the expression of the up‐regulated and

down‐regulated focus genes, respectively, in HSF and nHSF following treatment with

PDMS7‐g‐PEG7 for 6 and 48 hours. The differential expression of the miscellaneous focus

genes in HSF and nHSF cultures treated with GP226 and Fraction IV for 48 hours, as well as

PDMS7‐g‐PEG7 for 6 and 48 hours, are illustrated in Figure 5.5. Furthermore, comparisons

of fold expressions between HSF and nHSF were also made and are depicted in Figure 5.6.


Figure 5.1 – Validation of up‐regulated target genes in HSF and nHSF following treatment with GP226 and Fraction IV. Graphs of up‐regulated target gene validation, as determined by qRT‐PCR, in (A) HSF and (B) nHSF following treatment with GP226 and Fraction IV for 48 hours. RNA expression for each target gene was normalised to the 18S ribosomal gene and data from each treatment were expressed as the fold difference relative to the untreated control. Data are represented as mean ± SEM and were pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where expression is P<0.05 compared to the untreated control. The dashed line indicates the ± 1.8 fold change level in gene expression relative to the untreated control.

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Figure 5.2 – Validation of down‐regulated target genes in HSF and nHSF following treatment with GP226 and Fraction IV. Graphs of down‐regulated target gene validation, as determined by qRT‐PCR, in (A) HSF and (B) nHSF following treatment with GP226 and Fraction IV for 48 hours. RNA expression for each target gene was normalised to the 18S ribosomal gene and data from each treatment were expressed as the fold difference relative to the untreated control. Data are represented as mean ± SEM and were pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where expression is P<0.05 compared to the untreated control. The dashed line indicates the ± 1.8 fold change level in gene expression relative to the untreated control.


Figure 5.3 – Validation of up‐regulated target genes in HSF and nHSF following treatment with PDMS7‐g‐PEG7. Graphs of up‐regulated target gene validation, as determined by qRT‐PCR, in (A) HSF and (B) nHSF following treatment with PDMS7‐g‐PEG7 for 6 and 48 hours. RNA expression for each target gene was normalised to the 18S ribosomal gene and data from each treatment were expressed as the fold difference relative to the untreated control. Data are represented as mean ± SEM and were pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where expression is P<0.05 compared to the untreated control. The dashed line indicates the ± 1.8 fold change level in gene expression relative to the untreated control.

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Figure 5.4 – Validation of down‐regulated target genes in HSF and nHSF following treatment with PDMS7‐g‐PEG7. Graphs of down‐regulated target gene validation, as determined by qRT‐PCR, in (A) HSF and (B) nHSF following treatment with PDMS7‐g‐PEG7 for 6 and 48 hours. RNA expression for each target gene was normalised to the 18S ribosomal gene and data from each treatment were expressed as the fold difference relative to the untreated control. Data are represented as mean ± SEM and were pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where expression is P<0.05 compared to the untreated control. The dashed line indicates the ± 1.8 fold change level in gene expression relative to the untreated control.


Figure 5.5 – Validation of miscellaneous target genes in HSF and nHSF following treatment with GP226, Fraction IV and PDMS7‐g‐PEG7. Graphs of miscellaneous target gene validation, as determined by qRT‐PCR, in (A) HSF and (C) nHSF following treatment with GP226 and Fraction IV for 48 hours as well as in (B) HSF and (D) nHSF following treatment with PDMS7‐g‐PEG7 for 6 and 48 hours. RNA expression for each target gene was normalised to the 18S ribosomal gene and data from each treatment were expressed as the fold difference relative to the untreated control. Data are represented as mean ± SEM and were pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where expression is P<0.05 compared to the untreated control. The dashed line indicates the ± 1.8 fold change level in gene expression relative to the untreated control.

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Figure 5.6 – Validation of up‐regulated, down‐regulated and miscellaneous target genes in nHSF compared to HSF. Graphs of (A) up‐regulated, (B) down‐regulated and (C) miscellaneous target genes for nHSF compared to HSF at 6 and 48 hours. RNA expression for each target gene was normalised to the 18S ribosomal gene and data for nHSF were expressed as the fold difference relative to HSF. Data are represented as mean ± SEM and were pooled from three separate experiments, in which each treatment was performed in triplicate (n = 9). Significant differences (*) are illustrated where expression is P<0.05 compared to the untreated control. The dashed line indicates the ± 1.8 fold change level in gene expression relative to HSF.


Following treatment of HSF and nHSF with GP226 (Figure 5.1), expression of DAPK1, IGF2R,

IL8, NSMAF, SMAD7 and TNFRSF10B genes were up‐regulated. Interestingly, treating HSF

with Fraction IV (Figure 5.1) did not significantly affect many of the putative up‐regulated

focus genes. Thus, only the genes for SMAD7 and TNFRSF10B were significantly increased

when nHSF were treated with Fraction IV. While the IGF2R, IL8, NSMAF, SMAD7 and

TNFRSF10B genes were up‐regulated, they were not increased to the same extent as when

these same cells were treated with GP226. In terms of the down‐regulated focus genes

(Figure 5.2), both COL1A1 and COL3A1 were the only genes found to be down‐regulated

following treatment of HSF and nHSF with GP226 and Fraction IV. The genes for CFLAR and

TRAF3 were found to be significantly up‐regulated in nHSF treated with GP226, while only

TRAF3 was up‐regulated in HSF samples treated with GP226. The exposure of HSF and nHSF

to GP226 and Fraction IV did not significantly affect the expression of TRAF2, a ‘putative’

up‐regulated focus gene, nor αSMA or TGFβ1.

The effect of PDMS7‐g‐PEG7 on the expression of focus genes in HSF and nHSF were also

examined (Figures 5.3 and Figure 5.4). Furthermore, the effect of PDMS7‐g‐PEG7 on focus

genes was investigated at 6 and 48 hours. The 6 hours timepoint was included as many

genes changes occur quickly in vitro. Indeed, changes in gene expression were found to be

different at the two different time points. In most cases, gene expression of the focus

genes seemed to result in a greater fold change, compared to the untreated control, at 48

hours compared to the 6 hour treatment time. In HSF, the expression of AIFM2, FAS, IGF2R,

NSMAF and SMAD 7 genes were not different to the untreated control at 6 hours, but were

significantly increased at 48 hours. Expression of the genes for IL8 and TNFRSF10B was

increased in HSF treated with PDMS7‐g‐PEG7 at both 6 and 48 hours. In nHSF samples, the

FAS, NSMAF, SMAD7 and TNFRSF10B genes were significantly up‐regulated at 48 hours but

not at 6 hours treatment with PDMS7‐g‐PEG7. The expression of AIFM2, IGF2R and IL8

genes in nHSF treated with PDMS7‐g‐PEG7 was significantly increased at both 6 hours and

48 hours. Interesting, while most of the ‘putative’ up‐regulated focus genes were also

found to be up‐regulated in these qRT‐PCR validation studies, the DAPK1 gene was actually

found to be down‐regulated in HSF and nHSF following 48 hours treatment with PDMS7‐g‐

PEG7. The expression of TNF was not found to be differentially regulated in either HSF or

nHSF following treatment with PDMS7‐g‐PEG7 for either 6 or 48 hours.

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In terms of the down‐regulated focus genes, CLFAR was not found to be differentially

expressed in HSF treated with PDMS7‐g‐PEG7, but was significantly decreased at 6 but not

48 hours treatment with PDMS7‐g‐PEG7. Both the COL1A1 and COL3A1 genes were

significantly decreased in HSF and nHSF treated with PDMS7‐g‐PEG7 for 48 hours, with the

COL1A1 gene also being significantly decreased in nHSF treated with PDMS7‐g‐PEG7 for 6

hours. The TRAF2 gene was significantly decreased in HSF and nHSF treated with PDMS7‐g‐

PEG7 for 6 hours, but no differences were observed at 48 hours. The expression of the

TRAF3 gene was significantly up‐regulated following 48 hours exposure of PDMS7‐g‐PEG7 to

HSF and nHSF while expression was significantly decreased for nHSF only following 6 hours

treatment. The examination of the miscellaneous genes (Figure 5.5) demonstrated that

αSMA gene expression was not affected at 6 hours but was significantly decreased in HSF

and nHSF following 48 hours treatment with PDMS7‐g‐PEG7. It was also found that the

TGFβ1 gene was not significantly affected in either HSF or nHSF following treatment with

PDMS7‐g‐PEG7.

Comparisons of gene expression were also made between HSF and nHSF (Figure 5.6) it was

observed that many of the focus genes analysed, including AIFM2, FAS, IGF2R, NSMAF,

SMAD7, TNFRSF10B, COL3A1, TRAF3 and TGFβ1, were not differentially expressed between

the two cell types. It was observed, however, that the TNF gene was significantly decreased

in nHSF cultures, compared to HSF, at 6 hours. The IL8 gene was also found to be

significantly decreased in nHSF compared to nHSF at both 6 and 48 hours culture.

Furthermore, expression of the CFLAR, COL1A1, TRAF2 and αSMA genes in nHSF was

significantly decreased in cells exposed to the treatments for 48 hours, compared to HSF.

However, these differences were not observed at 6 hours.

The qRT‐PCR validation studies demonstrate that the expression of many of the target

genes were significantly affected in HSF and nHSF treated with the silicones when

compared to the untreated control. In most cases, the results of qRT‐PCR for all target

genes were consistent with those obtained from the microarray and superarray analyses.

For example, the IL8 gene was originally found to be up‐regulated in the microarray studies

and through validation via qRT‐PCR, the IL8 gene was also illustrated to be up‐regulated.


5.4 DISCUSSION

The focus of this chapter was on elucidating some of the mechanisms underpinning the

induction of apoptosis in fibroblasts following exposure to GP226, Fraction IV and PDMS7‐g‐

PEG7 in vitro. It is well established that changes in gene expression play major roles in

cellular biology (Risch and Merikangas, 1996). Therefore, gene microarray and superarray

experiments were used as screening techniques to observe the changes in fibroblast gene

expression following treatment with the silicones. While microarrays assess gene

expression over the whole genome, superarrays are a smaller scale array that analyse the

key genes involved in a specific biological function. In our studies, apoptosis superarrays

were used. It was anticipated that the use of these screening techniques would indicate

putative genes involved in the mechanistic pathways by which the silicone treatments were

exerting their effects. The gene microarray studies revealed over 1000 genes were found to

be differentially regulated in HSF and nHSF treated with the silicones, but only

approximately 20 genes were found to be differentially regulated in HSF and nHSF treated

with the silicones in the superarray studies. This can be explained by the fact that the

microarray assessed the whole genome (25000 genes) while the superarray investigated

only genes known to be involved in apoptosis (84 genes). The validation experiments,

conducted in cells from three different patients, further confirmed that many of the target

genes were indeed differentially regulated in fibroblasts following exposure to our silicone

treatments.

It was evident through the microarray and superarray screening process that genes related

to the TNF and TNF receptor (TNFR) superfamilies, including the genes for TNF FAS,

TNFRSF10B, TRAF2, TRAF3, NSMAF, CFLAR and DAPK1, were differentially regulated

following treatment with silicone. The TNF superfamily is a group of cytokines with

important functions in immunity, inflammation, differentiation, control of cell proliferation

and apoptosis (Shen and Pervaiz, 2006). TNF itself is the founding member of the

superfamily and is a pro‐inflammatory cytokine with a wide variety of functions in many

different cell types (Gupta, 2002). The TNF gene was demonstrated via the superarray

results to be up‐regulated following treatment with the silicone species and significant

differences were found between the untreated HSF and nHSF controls in our validation

experiments via qRT‐PCR. However, no significant differences were found through

validation between the untreated and silicone treated samples. This suggests that the

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silicone treatments were either affecting different members of the TNF superfamily or

other molecules involved in the induction of TNF‐induced apoptosis.

Members of the TNF family exert their biological effects through the TNFR superfamily. The

TNFR superfamily is compromised of cell surface receptors that share a stretch of

approximately 80 amino acids within their cytoplasmic region, named the death domain,

which is critical for recruiting other proteins required for the induction of apoptosis (Shen

and Pervaiz, 2006). Death receptors, including TNF receptor 1 (TNFR1), FAS, TNF receptor

superfamily member 10A (TNFRSF10A) and TNFRSF10B, are best characterized for their

role in the induction of apoptosis (Jin and El‐Deiry, 2005). Of note, the expression of the

genes for FAS and TNFRSF10B, were both identified through our superarray studies to be

up‐regulated in cell samples treated with the silicones. While the TNFRSF10B gene was

confirmed though qRT‐PCR to be significantly up‐regulated in HSF and nHSF treated with

GP226, Fraction IV and PDMS7‐g‐PEG7, the differentiated expression of the FAS gene was

not confirmed in the validation studies.

TNFRSF10B contains 2 extracellular cysteine‐rich repeats and the cytoplasmic death

domain (Walczak et al., 1997). TNFRSF10B engages a caspase‐dependant apoptotic

pathway and mediates apoptosis via the intracellular adaptor molecule FAS‐associating

protein with death domain (FADD) (Walczak et al., 1997). TNFRSF10B is a receptor for

TNFRSF10, which triggers apoptosis via interaction with its death receptors. Interestingly,

the gene for TNFRSF10B but not the gene for TNFRSF10’s other receptor, TNFRSF10A, was

found to be differentially regulated in our screening experiments. Furthermore, TNFRSF10

is characterised as a powerful activator of programmed cell death in tumour cells with

minimal toxicity against normal tissues (Almasan and Ashkenazi, 2003). While many human

tumour cell lines are sensitive to apoptosis induction by TNFRSF10, it has been reported

that most normal cells are not (Almasan and Ashkenazi, 2003). Despite this report, the

silicone treatments affected the expression of the TNFRSF10B gene in both HSF and nHSF,

however no significant differences in gene expression were evident between the HSF and

nHSF untreated controls.

Another gene family of interest, the TRAF gene family, are a family of six RING finger

containing proteins with a homologous TRAF domain at their C terminus and are widely

used as adaptor molecules by the TNFR superfamily (Bouwmeester et al., 2004; Rothe et


al., 1994). The TRAF2 and TRAF3 genes were found through the superarray results to be

down‐regulated in HSF and nHSF following treatment with silicone. Previous reports have

demonstrated that both TRAF2 and TRAF3 play an important role in cellular activation and

differentiation following engagement of a variety of TNFR superfamily members (Arch and

Thompson, 1998; Chang et al., 2002; Grammer and Lipsky, 2000; Hatzoglou et al., 2000;

Saitoh et al., 2003; Sinha et al., 2002). Members of the TRAF family can directly interact

with receptors containing no death domains, and can also interact with death domain

containing receptors through other death domain containing proteins (Hsu et al., 1996;

Rothe et al., 1994). For example, TRAF2 interacts with TNF receptor‐associated apoptotic

signal transducer (TRADD) to ensure the recruitment of inhibitor of apoptosis proteins (IAP)

for the direct inhibition of caspase activation (Zhang et al., 2010). Furthermore, TRAF2 has

also been reported to form a complex with TRAF1 to function as a mediator of anti‐

apoptotic signals from other TNF receptors (Zhang et al., 2010). The current qRT‐PCR

validation experiments demonstrated that the gene for TRAF2 is significantly decreased in

HSF and nHSF treated with PDMS7‐g‐PEG7 at 6 hours, but not following 48 hours treatment

with GP226, Fraction IV or PDMS7‐g‐PEG7. It is unknown whether this decrease in

expression is evident at 6 hours treatment with GP226 or Fraction IV as these were not

examined. However, taken together, it is plausible that the TRAF2 gene is down‐regulated

in the silicone treated samples early on; this may prevent anti‐apoptotic signals from

having effect. Decreased expression of the TRAF2 gene was also observed in untreated

nHSF when compared to untreated HSF, indicating that increased inhibition of apoptosis

was occurring in HSF and supporting the concept that the two cell types were different.

In addition to TRAF2, the initial gene superarray analyses revealed the TRAF3 gene to be

under expressed in HSF and nHSF following treatment with silicone. However, TRAF3 gene

expression was found though our validation experiments to be significantly increased

following HSF and nHSF being treated for 48 with GP226 and PDMS7‐g‐PEG7, but not

Fraction IV. This is most likely due to experimental errors occurring in the superarray

analyses. TRAF3 was first described as a molecule that participates in the signal

transduction of CD40, whereby it acts to mediate CD40 antibody secretion (Bradley and

Pober, 2001; Kuhne et al., 1997). TRAF3 is also recruited and used by the lymphotoxin‐β

receptor (LTβR) to induce apoptosis and can have an inhibitory effect on nuclear factor

kappa‐light‐chain‐enhancer of activated B cells (NF‐κB) activation through LTβR

(VanArsdale et al., 1997). Taken together, TRAF3 is one of the only members of the TRAF

C h a p t e r 5 . 0 | 115

family that has been reported to be involved in transducing, rather than inhibiting,

apoptotic effects, especially in LTβR‐induced apoptosis (VanArsdale et al., 1997).

The LTβR receptor, a member of the TNFR superfamily, is expressed on the surface of most

cell types (Chang et al., 2002). It has been reported that LTβR can induce IL8 release in

A375 human malignant melanoma cells (Degli‐Esposti et al., 1997; Hehlgans and Mannel,

2001). Furthermore, it was reported by Chang et al. (2002) that dominant negative mutants

of TRAF2, 3 and 5, could attenuate IL8 production induced by LTβR. Taken together, it

appears that the over expression of TRAFs is implicated in regulating the expression of IL8

through the LTβR. While our data demonstrated that treating fibroblasts with the silicones

led to decreased expression of the TRAF2 gene, increased expression of the TRAF3 gene

was observed. Interestingly, the IL8 gene was identified in the gene microarray studies, and

further validated through qRT‐PCR, to be highly up‐regulated in HSF and nHSF samples

exposed to our silicone treatments.

IL8 is often associated with inflammation, is secreted in a stimulus‐specific manner by a

variety of cell types and is regulated primarily at the level of gene transcription (Chang et

al., 2002; Gillitzer and Goebeler, 2001; Schauer et al., 2009). It has also been reported that

up‐regulation in IL8 protein occurs in fibrotic and malignant diseases, is increased by

oxidant stress and is a key mediator of proliferative responses (Schauer et al., 2009).

Interestingly, when comparing the untreated controls in our qRT‐PCR experiments, the

expression of the IL8 gene was found to be significantly decreased in nHSF compared to

HSF, which corresponds with the findings reported by Schauer et al. (2009) and indicates

that the two cell populations tested are in fact different. As mentioned above, up

regulation of IL8 gene expression was confirmed through the qRT‐PCR validation

experiments in HSF and nHSF treated with silicone. The upregulation of the IL8 gene by the

silicones could be considered to be less than ideal as increased inflammation in a

hypertrophic scar would not be desirable. However, when expression of the IL8 gene was

examined in HSF and nHSF cultures treated with PDMS7‐g‐PEG7 for both 6 and 48 hours, it

was observed that IL8 gene expression decreased between the 6 hour and 48 hour

timepoint. It has been reported that levels of IL8 peak during day 1 of wound healing and

decrease thereafter (Engelhardt et al., 1998). It is therefore possible that the initial peak of

IL8 gene expression is initially induced due to chemical insult and expression decreases

following this. It is unknown if IL8 gene expression levels continue to decrease following 48


hours. Further studies investigating this phenomenon, together with the effect of silicone

treatment on TRAF and IL8 expression in vivo, are clearly warranted.

Ceramide is well established as a bioactive lipid involved in the cellular responses to stress

and has been associated with the TNF superfamily (Clarke et al., 2008; Wu et al., 2005). TNF

has been reported to activate ceramide through proteins that bind to neutral

sphingomyelinase (NSMase) and its adapter protein, NSMAF, which specifically binds to a

distinct cytoplasmic domain of the TNFR1 called the neutral sphingomyelinase activation

domain (Adam‐Klages et al., 1996; Adam et al., 1996; Kolesnick and Kronke, 1998).

Importantly, this activation domain has the amino acid sequence, QKWEDSAHK (Adam‐

Klages et al., 1996; Adam et al., 1996). It is interesting to note that the cytoplasmic domain

of TNF receptor superfamily member 5 (TNFRSF5) contains a short sequence, QETLH, which

shares some homology to the neutral sphingomyelinase activation domain. It has been

hypothesised that NSMAF can also bind to this short sequence (Segui et al., 1999). In

addition, the threonine reside present in the middle of this sequence (Thr‐234) has been

described to be essential in various biological responses to TNF receptor superfamily

member 5 (TNFRSF5) ligation, such as: homotypic adhesion (Hostager et al., 1996);

interaction with TRAF2, TRAF3 and TRAF5 proteins (Hanissian and Geha, 1997; Hu et al.,

1994; Ishida et al., 1996; Lee et al., 1999; Pullen et al., 1998; Tsukamoto et al., 1999); c‐Jun

N‐terminal kinase activation (Lee et al., 1999); NF‐κB activation (Lee et al., 1999;

Tsukamoto et al., 1999); and importantly, apoptotic/growth inhibitory effects (Inui et al.,

1990; Segui et al., 1999). Moreover, overexpression of the full‐length NSMAF gene, as was

found in our studies, enhances NSMase activity in TNF‐treated cells, increasing the

occurrence of apoptosis (Adam‐Klages et al., 1996).

CFLAR, also known as a caspase 8 homologue, was first identified as a molecule that

interacts with FAS (Budd et al., 2006). The superarray analyses demonstrated that CFLAR

gene expression was decreased in HSF and nHSF following silicone treatment. As a

characteristic feature, CFLAR contains a tandem death effector domain, which is used to

inhibit death receptor‐induced apoptosis by binding to and preventing caspase 8 activation

(Budd et al., 2006; Krueger et al., 2001; Scaffidi et al., 1999). Furthermore, it has been

reported that TNF induces caspase‐dependent and independent Jun N‐terminal kinase

(JNK) activation and reactive oxygen species (ROS) accumulation in CFLAR knockout murine

embryonic fibroblasts (Nakajima et al., 2006). Despite these reports linking CFLAR to

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apoptosis, CFLAR gene expression was found to be differentially regulated in silicone

treated fibroblasts in some of our validation experiments. This includes decreased

expression in nHSF following treatment with PDMS7‐g‐PEG7 for 6 hours and increased

expression in nHSF following treatment with GP226 for 48 hours. In addition, a significant

difference in CFLAR gene expression was also observed in the untreated nHSF control when

compared to the untreated HSF control. Why these differences were not consistent

throughout all the validation studies, or why differential expression following silicone

treatment were not observed in any HSF samples is not clear.

The DAPK1 gene was another protein found to be over expressed in our superarray

analyses. The DAPK1 protein is the founding member of a newly classified family of Ser/The

kinases, whose members not only possess significant homology in their catalytic domains,

but also share cell death‐associated functions (Bialik and Kimchi, 2006). The C terminus of

DAPK1 contains a death effector domain, which when over expressed, acts as a dominant

negative regulator of apoptosis and protects cells from death induced by TNF or FAS (Bialik

and Kimchi, 2006; Cohen et al., 1999). Interestingly, the validation experiments revealed

that the DAPK1 gene was increased following treatment with GP226, yet was not

significantly affected following treatment with Fraction IV to HSF and nHSF. Furthermore,

decreased DAPK1 gene expression was observed in HSF and nHSF following treatment with

PDMS7‐g‐PEG7. These results are clearly inconsistent, making it difficult to understand the

role of DAPK1 in dermal fibroblasts treated with the silicones.

Through the evaluation of the results reported in this chapter, it has become evident that

apoptosis occurring through the participation of TNF and TNFR superfamily members can

be via many different pathways. In addition to what has already been discussed, TNF‐

induced apoptosis has also been reported to be modulated by mannose‐6‐phosphorylated

glycoproteins (Tardy et al., 2004). Furthermore, Chen et al. (2004) demonstrated that

decreased expression of mRNA for IGF2R, a receptor for mannose‐6‐phosphate (M6P), led

to increased cell proliferation and decreased cell susceptibility to TNF‐induced apoptosis.

The experiments reported in this chapter reveal that the expression of the IGF2R gene was

up‐regulated in HSF and nHSF following treatment with silicone, especially at the 48 hour

timepoint; this coincides with our previous results showing decreased proliferation and

increased apoptosis. IGF2R has been reported to be a growth inhibitor/tumour suppressor

(Probst et al., 2009) involved in the activation of TGFβ and also in cellular functions related


to apoptosis and tumorigenesis (Motyka et al., 2000). In addition, the IGF2R controls

concentrations of insulin‐like growth factor‐II (IGFII), a growth factor, through

internalization and lysosomal degradation. This has been reported to suppress mitogenesis

by reducing the availability of IGFII to mediate actions via the IGFI receptor (Jones and

Clemmons, 1995).

Reports that the IGF2R is involved in the activation of TGFβ are interesting in that TGFβ1

stimulates collagen production in normal human dermal fibroblasts (Varga et al., 1987).

The gene microarray studies demonstrate that many of the genes for different collagen

types, especially COL1A1 and COL3A1, the main types of collagen found in skin, were

decreased in HSF and nHSF samples exposed to GP226 and Fraction IV (Uitto et al., 1981).

This decrease in collagen production was also validated in the qRT‐PCR experiments,

whereby the COL1A1 and COL3A1 genes were found to be significantly decreased in HSF

and nHSF treated with GP226, Fraction IV and PDMS7‐g‐PEG7 for 48 hours. Furthermore,

the expression of the COL1A1 gene was also found to be significantly decreased in

untreated nHSF compared to the untreated HSF control. It has been reported that TGFβ1

overproduction at both mRNA and protein levels is associated with several

fibroproliferative disorders, such as pulmonary fibrosis, liver cirrhosis, glomerulonephritis

and scleroderma (Border and Noble, 1994; Broekelmann et al., 1991; Castilla et al., 1991;

Gruschwitz et al., 1990). However, despite our findings demonstrating differential COL1A1

gene expression between cell types and following silicone treatment, no significant

differences in TGFβ1 gene expression were found in any of our qRT‐PCR experiments. It is

possible that differences in the expression of genes for the other forms of TGFβ are present

in HSF and nHSF. For example, TGFβ1 and TGFβ2 are known to modulate collagen

synthesis, while TGFβ3 has been demonstrated to reduce scarring and is involved in scar

prevention (Bock et al., 2005; Shah et al., 1992).

The gene for αSMA was included in the qRT‐PCR validation studies as it is regulated by the

action of growth factors, like TGFβ1, and is the most widely employed marker of

myofibroblasts (Hinz et al., 2007). Following tissue injury, fibroblasts incorporate αSMA into

their stress fibres facilitating differentiation into contractile and secretory myofibroblasts

which contribute to tissue repair during wound healing (Hinz et al., 2001; Hinz et al., 2007).

However, the presence of myofibroblasts can severely impair organ function when

contraction and ECM protein secretion become excessive, such as in hypertrophic scars

C h a p t e r 5 . 0 | 119

(Hinz, 2007; Hinz et al., 2007). The qRT‐PCR studies showed that αSMA gene expression

was decreased in HSF and nHSF following 48 hours treatment with PDMS7‐g‐PEG7,

potentially meaning that PDMS7‐g‐PEG7 was decreasing the contractile and secretory action

of the fibroblasts in vitro. Additionally, the expression of the αSMA gene was significantly

decreased in untreated nHSF compared to untreated HSF after 48 hours culture. This

provides further support that the two cell types are actually distinct cell populations and

that the HSF have not reverted to normal fibroblast phenotype during in vitro culture.

The SMAD7 gene was also identified as a focus gene in the microarray studies and was

hence further validated via qRT‐PCR. These studies confirmed that the SMAD7 gene is up‐

regulated following silicone treatment. SMAD7 is another protein that interacts with TGFβ1

and regulates TGFβ1 signal via a negative feedback loop (Yan et al., 2009). In fact, the

TGFβ/SMAD signalling system is well known for an autoinhibitory loop that involves SMAD7

forming a complex with R‐SMADs, preventing their movement from the cell nucleus. The

transient induction of SMAD7 in response to TGFβ1 acts as an important negative feedback

inhibitor of TGFβ1 signalling transduction by blocking the activation of SMAD2 and SMAD3

proteins and preventing their interaction with activated TGFβ1 receptors and subsequent

phosphorylation (Xie et al., 2008). SMAD7 has well described anti‐fibrotic and anti‐

inflammatory activities and it has been reported that fibroblasts derived from keloid scars

have a decreased expression of SMAD7 (Tang et al., 2009). However, little is known about

the relationship between TGFβ1 and SMAD signalling in the hypertrophic scar (Xie et al.,

2008). Additionally, no significant differences in SMAD7 gene expression were found

between HSF and nHSF.

Finally, AIFM2 was identified as a target gene though our microarray studies, and was

confirmed via qRT‐PCR to be over‐expressed in HSF and nHSF following treatment with

silicone. There is only a small amount of information in the literature concerning AIFM2,

but it appears that apoptosis induced by AIFM2 is not p53‐dependant nor caspase‐

dependant (Marshall et al., 2005; Wu et al., 2002). Despite this, there is still debate as to

whether AIFM2 actually associates with the mitochondria, as it name suggests, or with the

plasma membrane (Bilyy et al., 2008).

Taken together, it is evident that many events are occurring in dermal fibroblasts when

they are treated with the silicone treatments. Significant differences in the expression of


many of the focus genes were also found between the two untreated controls, HSF and

nHSF. These findings were significant as the initial microarray analyses, which were

performed with fibroblasts derived from one patient only, identified less than 100 genes to

be differentially expressed between untreated HSF and nHSF. Furthermore, the validation

experiments indicated that the two cell populations were in fact distinct and that the HSF

had not differentiated back to normal fibroblasts during in vitro culture. Through analysis of

the literature, it has also become clear that many of the genes identified through the

microarray and superarray screening techniques are related to the TNF and TNFR

superfamilies. Indeed, while the gene expression data assembled is important, validation at

the protein level is now required. Proteins of interest to investigate further would include

αSMA, AIFM2, COL1A1, COL3A1, IGF2R, NSMAF, SMAD7, TRAF2, TRAF3 and TNFRSF10B.

The studies outlined in this chapter have, however, increased our knowledge base

regarding which genes and biological pathways may be affected through the application of

products containing silicone, particularly silicone‐PEG copolymers, to hypertrophic scars.

C h a p t e r 6 . 0 | 121

CHAPTER 6.0

GENERAL DISCUSSION AND CONCLUSIONS

The formation of hypertrophic scars is a frequent outcome of wound repair and often

results in disfigurement, pain, itching, reduced mobility and psychological trauma to those

suffering from them (Ferguson and O'Kane, 2004; Mutsaers et al., 1997). Nationally,

170,000 burn and scald injuries, all with the potential for adverse scarring, are reported

each year in Australia (Australian Bureau of Statistics, 2001). On a wider scale,

approximately 100 million people develop scars each year in the developed world. It is

estimated that 11 million of these scars develop abnormally and 4 million result from burn

injuries, of which 70% occur in children (Bayat et al., 2003). Of note, the survival rates of

burns patients have increased significantly in the last couple of decades, leading to a

pressing need for improved and optimised wound healing methods (Bloemen et al., 2009).

Moreover, a study performed by Li‐Tsang et al. (2005) demonstrated that the prevalence

rate of post‐surgical hypertrophic scarring was 74.7%. Clearly, scarring is a major medical

side effect and any improvement in scar management will impact on many lives, not only

increasing aesthetics, but also enhancing quality of lives by restoring physical movement to

affected areas and relieving emotional trauma that may accompany abnormal recovery

processes.

SGS are currently used as a treatment for hypertrophic scars, with many beneficial effects

being reported following use (Ahn et al., 1989; Berman and Flores, 1999; Eishi et al., 2003;

Gold et al., 2001; Quinn et al., 1985). Although widely used, no specific factor has been

proven to be the major contributor or inducer of the scar remediation properties attributed

to SGS. Research undertaken by our laboratory re‐investigated the chemical role of SGS on

scarring. It was demonstrated that small‐amounts of amphiphilic, linear, oligomeric

silicones with low molecular weight have the ability to migrate from commercially available

SGS into the human stratum corneum (Sanchez et al., 2005). In view of this, the

investigations described within this doctoral thesis were based around assessing the

potential scar remediating ability of low molecular weight amphiphilic silicone species. The

underlying hypothesis of this doctoral thesis was that low molecular weight and

amphiphilic silicones decrease cell viability by regulating apoptosis in dermal fibroblasts

and keratinocytes.

G e n e r a l D i s c u s s i o n a n d C o n c l u s i o n s

The research began by investigating the commercially available oligomeric silicone, GP226,

which is structurally similar to those found previously to traverse the stratum corneum

(Sanchez et al., 2005). These experiments were motivated by a need to validate the earlier

findings from our laboratory indicating that exposure of dermal fibroblasts to GP226 and its

five fractions decreased cell viability (Charters, 2007). Furthermore, these earlier studies

needed to be extended as the effect of the silicones were only assessed on fibroblasts and

did not include keratinocytes. In addition, cell viability, but not cell proliferation, was

investigated and the effects were only assessed at 48 hours. However, a report by

Hanasono et al., (2004) had shown that differences in cell growth curves occurred between

days 2 and 5 following application of silicone to dermal fibroblasts, prompting us to pursue

studies examining more time points.

The first studies reported in this thesis (Chapter 3.0) confirmed the previously reported

results (Charters, 2007) and illustrated that GP226, fractionated into five species of

differing molecular weights, reduces cell viability and proliferation of HSF, nHSF, KF, nKF

and HK at 24, 48, 72 and 168 hours (7 days). Interestingly, differing effects on cell viability

and proliferation were observed when the cells were treated with the five fractions of

GP226. It was observed that fractions III, IV and V were the most active components of

GP226, dose‐dependently decreasing viability and proliferation of all cell types. Of note,

application of Fraction IV to dermal fibroblasts and keratinocytes in vitro was found to

induce effects that were most similar to GP226. Unlike Hanasono et al., (2004), differences

in the trends observed for viability or proliferation of HSF, nHSF, KF and nKF were not

apparent following treatment with silicone at the four time points assessed. Therefore, 48

hours was selected as the best timepoint for subsequent experiments as it was the

timepoint that led to the greatest proliferation of cells in the majority of the untreated

control cultures.

Another key aspect of the analyses reported in Chapter 3.0 is that the effects of both HSF

and KF were examined. This was especially important as limited studies on the effect of

GP226 on keloid‐derived tissues had been conducted. Keloid and hypertrophic scars are

histologically and morphologically different (Bayat et al., 2003) and the fibroblasts derived

from keloid scars appear and behave differently to those derived from hypertrophic scars

(Hanasono et al., 2004). Our analyses of cell viability and proliferation, however,

C h a p t e r 6 . 0 | 123

demonstrated no major differences between HSF and KF or nHSF and nKF when compared

to their respective controls. Furthermore, differences in KF behaviour were not observed,

but this could have also been due to the fact that cells of passage 4‐7, rather than cells

derived directly from tissues, were utilised. As no pronounced differences between HSF

and KF in the studies reported in Chapter 3.0 were observed, the future analyses described

within this thesis investigated only HSF and nHSF.

In the experiments that demonstrated decreased cell viability and proliferation following

treatment with GP226 and its fractions, a concern became apparent that perhaps the

silicones were inducing aggregation of FCS proteins within the cell culture medium. This

hypothesis originated from the report that silicone oil has previously been implicated in the

induction of protein aggregation (Bernstein, 1987). Furthermore, Jones et al. (2005)

reported that silicone oil‐induced aggregation of proteins occurred dramatically with BSA,

possibly causing it to undergo gross conformational changes and rendering it non‐

functional. In our experimental situation, it was thought that the aggregation of FCS

proteins may possibly ‘starve’ the cells of their required nutrients and therefore be the

underlying cause for decreased cell viability and proliferation. To investigate this, HSF were

cultured with medium containing FCS of varying concentrations up to 20% and the cells

simultaneously treated with silicone. Of note, 20% FCS in cell culture medium is considered

a significant over supply of nutrients to the cells. However, following treatment with the

silicones in cell culture medium containing 20% FCS, cell density was still dramatically

decreased. Moreover, similar responses were obtained regardless of what concentrations

of FCS were present.

Following on from this, the cell morphology of HSF exposed to GP226, Fraction IV and a

commonly used surfactant, Tween 20, was examined in order to observe what was

happening morphologically to the cells during treatment. Real‐time microscopy was utilised

to observe cell morphology as it enables the cells to be imaged in high definition over an

extended period of time (Wilson et al., 1998). Through these experiments it became

evident that HSF exhibited morphology indicative of apoptosis, including cell shrinkage,

membrane blebbing and nuclear fragmentation following treatment with silicone. This

observation of apoptosis in HSF following treatment with GP226 and Fraction IV was then

confirmed using the Tunel assay. Interestingly, when HSF were exposed to Tween 20, the

cells appeared to be undergoing a form of apoptosis called anoikis; this is induced when


anchorage‐dependent cells detach from the surrounding ECM (Frisch and Screaton, 2001).

Tween 20 was included in these investigations to determine whether the action of GP226

and Fraction IV arose from their properties as surfactants. Given that the phenomenon of

anoikis was not observed in the HSF cultures exposed to GP226 or Fraction IV, it suggests

that their effect is independent of their surfactant properties.

Taken together, the results detailed in Chapter 3.0 demonstrated that one fraction, IV, was

largely responsible for the effects elicited by GP226. The methodology required for

obtaining pure samples of Fraction IV from GP226 involved separation via PSEC and HPLC.

Fraction IV represents only a small proportion (8%) of the complete GP226 mixture,

meaning that large amounts of GP226 starting material needed to be fractionated in order

to obtain small amounts of Fraction IV. In addition, the PSEC and HPLC methodology

required for the fractionation is labour‐intensive and time inefficient. In view of this, it

became apparent that the silicone treatments would need to be synthesised within our

laboratory in order to achieve the level of purity and quantity of silicones required for

experimentation. As Fraction IV had demonstrated the most consistent and dose

dependant results within Chapter 3.0, a series of amphiphilic silicones were synthesised

based on its structure. A small library of amphiphilic silicone oligomers, including PDMS15.2‐

PEG8, PDMS10.5‐PEG8, PDMS15.2‐PEG4 and PDMS7‐g‐PEG7, with low molecular weight and a

broad range of properties were synthesised by chemists within the laboratory. Importantly,

the amphiphilicity of these synthesised silicones, as indicated by their HLB factor in table

4.1, varied greatly. Of note, HLB indicates the balance of the degree of hydrophobicity and

hydrophilicty within a chemical structure, whereby lower HLB values indicate more

hydrophobic compounds, such as PDMS15.2‐PEG8 and PDMS15.2‐PEG4, while higher values

indicate more hydrophilic compounds, including PDMS10.5‐PEG8 and PDMS7‐PEG7.

The goal of the studies in Chapter 4.0 was to find a species of silicone that had either equal

or greater effects on HSF, nHSF and HK than GP226 and Fraction IV, in terms of cell

proliferation and apoptosis. As no major differences were observed in Chapter 3.0 between

our proliferation and viability data, only cell proliferation was investigated following

exposure to the synthetic silicones. These experiments indicated that each of the

synthesised silicones had differing effects on HSF and nHSF. In fact, PDMS7‐g‐PEG7 was the

only silicone to decrease both HSF and nHSF proliferation to levels below those obtained

for GP226 and was therefore chosen for further investigation. Interestingly, comparing all

C h a p t e r 6 . 0 | 125

the synthetic silicones together, PDMS7‐g‐PEG7 had the lowest molecular weight, was the

most hydrophilic and was the only synthesised silicone oligomer that consisted of a rake

structure. Further experimentation with PDMS7‐g‐PEG7 demonstrated that it could also

elicit dose dependant decreases in HSF, nHSF and HK proliferation when applied at a wider

range of concentrations. Not surprisingly, the results obtained with PDMS7‐g‐PEG7 were far

more consistant than those obtained with GP226 and its fractions. As PDMS7‐g‐PEG7 was

manufactured in house using high quality materials, its purity was able to be maintained at

a level suitable for application to cells cultured in vitro.

Given the earlier experiments that demonstrated GP226 and Fraction IV induced apoptosis

when applied to dermal fibroblasts, it seemed likely that application of PDMS7‐g‐PEG7 to

HSF, nHSF and HK also induced apoptosis. Indeed, induction of apoptosis by PDMS7‐g‐PEG7

was demonstrated through analyses of cell morphology and the Tunel assay in cultures of

HSF, nHSF and HK. Interestingly, decreased apoptosis of fibroblasts has been reported as a

major factor in the etiopathogenesis of hypertrophic scars (Appleton et al., 1996; Saray and

Gulec, 2005; Sayah et al., 1999). While the regulation of ECM deposition and remodelling

are key elements in hypertrophic scar formation, so too is tissue homeostasis, which is

maintained through a balance between cell proliferation and death (Bellemare et al., 2005;

Eckes et al., 2000; Moulin et al., 2004). Apoptosis is a method of controlled cell death that

has been widely investigated. The process is marked morphologically by cellular shrinking,

condensation and margination of the chromatin, as well as ruffling of the plasma

membrane, with eventual break‐up of the cell in apoptotic bodies (Taylor et al., 2008).

Investigations using real‐time and immunofluorescent microscopy revealed that GP226,

Fraction IV and PDMS7‐g‐PEG7 induce apoptosis in cultures of HSF, nHSF and HK. In

addition, in some of the cell morphology experiments it was observed that cell contents

were ruptured following the induction of apoptosis with the silicones, hence indicating

necrosis. However, it has been reported that the distinction between apoptosis and

necrosis in cells cultured in vitro can be confused because there is a lack of scavenging cells

present, and thus the phagocytic step after apoptosis may not occur. Instead, as was

observed, the apoptotic cell can eventually undergo secondary necrosis and release its

contents into the surrounding medium (Duke and Cohen, 1986).

These results concerning the induction of apoptosis following treatment with silicone

demonstrated to us that amphiphilic silicone oligomers with low molecular weight are


potential novel scar treatments that can remove the tissue bulk and prevent the excess

ECM deposition that is associated with abnormal scars. In fact, cytotoxic agents, such as

bleomycin, are currently used to reduce the extraneous tissue associated with hypertrophic

scarring (Espana et al., 2001; Meier and Nanney, 2006). Bleomycin is an apoptotic agent

that is also used in chemotherapy and has the potential to cause major side‐effects, such as

systemic toxicity, as well as pulmonary, renal and cutaneous fibrosis in the longer term

(Crooke and Bradner, 1976; Shastri et al., 1971). Furthermore, its use requires application

by injection, an invasive procedure with increased associated complications (Crooke and

Bradner, 1976; Shastri et al., 1971). Clearly, a non‐invasive topical silicone treatment that

stimulates an equivalent apoptotic effect with minimal side‐effects would be

advantageous.

The results obtained through the studies reported in Chapters 3.0 and 4.0 led us to propose

that the silicone species are able to interact with dermal fibroblast cells and affect key

cellular mechanisms pertinent to cell proliferation and apoptosis. However, the

mechanisms occurring in fibroblasts undergoing apoptosis during treatment were still

unknown. The focus of Chapter 5.0 therefore was to elucidate some of the mechanisms

underpinning the induction of apoptosis in fibroblasts following exposure to GP226,

Fraction IV and PDMS7‐g‐PEG7 in vitro. Although the earlier investigations reported in this

thesis involved studies with both fibroblasts and keratinocytes, the mechanistic

experiments focussed only on fibroblasts. As changes in gene expression play major roles in

cellular biology, the initial experiments focussed on investigating differential gene

expression in dermal fibroblasts following exposure to the silicone treatments (Risch and

Merikangas, 1996). To explore this, gene microarray and superarray experiments were

used as screening techniques to identify possible ‘target’ genes and mechanisms by which

the silicone treatments were mediating their effects. The HumanHT‐12 v3 Expression

BeadChip microarray was used to determine differentially expressed genes in cultures of

HSF and nHSF following treatment with and without GP226 and Fraction IV, while qRT‐PCR

apoptosis superarrays were performed on HSF and nHSF cultures exposed to PDMS7‐g‐

PEG7. Following analysis of the microarray and superarray results, the differential

expression of 16 ‘target’ transcripts were validated by qRT‐PCR, using 18S as an internal

control for normalisation. A summary of these results is illustrated in Table 6.1.

C h a p t e r 6 . 0 | 127

Focus Genes Silicone treated samples HSF vs nHSF

Microarray Superarray qRT‐PCR qRT‐PCR

αSMA ‐ ‐ Down ‐

AIFM2 Up ‐ Up ‐

CFLAR ‐ Up Inc Down

COL1A1 Down ‐ Down Down

COL3A1 Down ‐ Down ‐

DAPK1 ‐ Down Inc ‐

FAS ‐ Down Inc ‐

IGF2R Up ‐ Up ‐

IL8 Up ‐ Up Down

NSMAF Up ‐ Up ‐

SMAD7 Up ‐ Up ‐

TGFβ1 ‐ ‐ ‐ ‐

TNF ‐ Down ‐ Down

TNFRSF10B ‐ Up Up ‐

TRAF2 ‐ Down Down ‐

TRAF3 ‐ Down Up ‐

Table 6.1 – Summary of Results obtained for microarray, superarray and qRT‐PCR analyses. Genes depicted as being differentially regulated are indicative of changes in silicone‐treated samples compared to the untreated control or in untreated nHSF when compared to untreated HSF. Genes marked with – indicate those with no significant changes in gene expression or in terms of the superarray experiments, were not included in the array performed. Genes marked with Inc indicate results that were inconclusive or not significant.

It was evident through the microarray and superarray screening process that genes related

to the TNF and TNFR superfamilies, including the TNF, FAS, TRAF2, TRAF3, NSMAF,

TNFRSF10B and CFLAR genes, were differentially regulated following treatment with

silicone. Furthermore, many genes involved in the collagen production pathway, including

the genes for COL1A1, COL3A1, IGF2R and SMAD7, were also found to be differentially

regulated through the microarray analyses performed. Table 6.1 summarises that that

some of the investigated focus genes, including CFLAR, DAPK1 or FAS, were not significantly

differentially regulated or confirmed by the qRT‐PCR validation experiments. Similarly,

TRAF3 gene expression was found to be down‐regulated in our gene superarray analyses,

yet was significantly increased following treatment of GP226 and PDMS7‐g‐PEG7, but not

Fraction IV to HSF and nHSF in the qRT‐PCR experiments. These conflicting results may arise

from the fact that the microarray and superarray analyses were performed on HSF and


nHSF derived from one patient only, while the validation experiments were performed on

cells derived from three different patients. Despite this, the validation experiments

confirmed that many of the target genes were differentially regulated in fibroblasts

following exposure to the silicone treatments. Indeed, through the analysis of the peer‐

reviewed literature in Section 5.4, many links between the results obtained and cellular

processes important in apoptosis, proliferation and collagen production were found. Figure

6.1 depicts a summary of the pathways involved and predicted relationships made

between the protein products of the genes identified as being differentially regulated

following treatment of the silicone treatments to HSF and nHSF.

Figure 6.1 – Predicted relationships between differentially regulated genes identified in HSF and nHSF following treatment with the silicones. Genes for molecules in red indicate those found to be up‐regulated, blue indicates those found to be down‐regulated and green indicates those focus genes not found to be differentially regulated. Molecules in yellow indicate other important proteins that were not identified through microarray or superarray analyses to be differentially regulated at gene level in HSF and nHSF following treatment with silicone. Pathways marked with indicate an increased effect while pathways marked with indicate an inhibitory effect.

As is illustrated in Figure 6.1, although the gene for TNF was not differentially expressed in

fibroblasts following treatment with the silicones, apoptosis induced by members of the

TNF superfamily requires the interaction with many other molecules and can do this via

many different pathways. For example, many TNF family members, including TNF,

C h a p t e r 6 . 0 | 129

TNFRSF10 and LTβR, have been reported to exert their biological effects through TNFR

superfamily members, such as TNFRSF10B, and other adaptor molecules, NSMAF, TRAF2

and TRAF3 (Adam‐Klages et al., 1996; Adam et al., 1996; Bouwmeester et al., 2004; Gupta,

2002; Jin and El‐Deiry, 2005; Rothe et al., 1994; VanArsdale et al., 1997). As summarised in

Table 6.1 and Figure 6.1, the genes for all of these proteins were found to be differentially

expressed at the genomic level in dermal fibroblasts following treatment with the silicones.

Another gene identified to be up‐regulated following exposure to the silicones was AIFM2,

which has been reported to induce apoptosis that is neither p53‐dependant nor caspase‐

dependant (Marshall et al., 2005; Wu et al., 2002). Associations between AIFM2‐ and TNF‐

induced apoptosis or the TNFR superfamily were not apparent in the literature. It has also

been demonstrated that many of the protein products of the investigated genes belonging

to or associating with the TNF and TNFR superfamilies are involved in other pathways that

are not related to the induction of apoptosis. For example, IL8 gene expression, found to

be up‐regulated in the qRT‐PCR experiments, has been reported to be regulated through

the LTβR (Chang et al., 2002). Furthermore, Chen et al., (2004) also reported that

decreased expression of IGF2R mRNA can lead to increased cell proliferation and decreased

cell susceptibility to TNF‐induced apoptosis, which corresponds with our results of

increased IGF2R gene expression and increased apoptosis following application of the

silicones to HSF and nHSF.

In addition to the role of IGF2R in apoptosis, IGF2R is also involved in the activation of

TGFβ, a protein that has been widely reported to stimulate collagen production in normal

human dermal fibroblasts (Varga et al., 1987). Indeed, our investigations of differential

gene expression illustrated that the genes for both COL1A1 and COL3A1, the two main

types of collagen within skin, were decreased in silicone‐treated fibroblasts (Uitto et al.,

1981). Furthermore, links between TGFβ and αSMA and SMAD7, two other genes

investigated in our expression studies, have also been reported. αSMA, identified as being

decreased at the genomic level in HSF and nHSF following 48 hours treatment with PDMS7‐

g‐PEG7, is the most widely employed marker of myofibroblasts, the proliferative and

secretory fibroblasts responsible for collagen production during wound healing (Hinz et al.,

2001; Hinz et al., 2007). Moreover, It has been extensively reported that the TGFβ/SMAD

signalling system, involved in collagen production, includes an autoinhibitory loop that is

largely mediated by SMAD7 (Xie et al., 2008). Although little is known regarding the


relationship between SMAD7 and hypertrophic scars, it is known that SMAD7 has anti‐

fibrotic and anti‐inflammatory effects (Tang et al., 2009; Xie et al., 2008).

It is evident that the effect of exposure of dermal fibroblasts to the silicones is significant,

with many genes being differentially regulated following treatment. One observation that

became apparent through these studies was that many of the genes for the cytokines

investigated, including TNF, Fas and TGFβ1, were not found to be differentially regulated in

the qRT‐PCR validation experiments. Interestingly, many of the receptors reported to be

involved with these cytokines were found to be differentially regulated. The most likely

reason underlying this phenomenon is that the fibroblasts are regulating receptor

expression as a mechanism to induce apoptosis quickly following exposure to the silicone

treatments. It is also possible that the silicone treatments are directly interacting with the

cell membrane during treatment, inducing a mechanistic effect and regulating receptor

activity as a result. Indeed, other studies performed within our laboratory have attempted

to investigate the interaction and location of silicone within fibroblasts but the methods

adopted have proved troublesome thus far and the results are still inconclusive (Keddie, D.

and Farrugia, B. pers. comm.).

The gene expression studies performed in Chapter 5.0 also demonstrated that many

differences in gene expression were present between HSF and nHSF, the two main types of

fibroblasts investigated within this doctoral thesis. For example, the expression of the IL8

gene was found to be significantly decreased in nHSF compared to HSF. These results

corroborate those of Schauer et al. (2009), who reported that IL8 protein expression is up‐

regulated in fibrotic and malignant diseases, increased by oxidant stress and is a key

mediator of proliferative responses. In addition, expression of the TNF gene, a pro‐

inflammatory cytokine with many functions, as well as the TRAF2 gene, involved in

inhibiting apoptosis, were also found to be decreased in nHSF compared to HSF (Gupta,

2002). Finally, expression of the gene for αSMA was significantly decreased in untreated

nHSF compared to untreated HSF at 48 hours culture. As mentioned previously, αSMA is

the most widely employed marker of myofibroblasts (Hinz, 2007). The finding of decreased

expression of αSMA in nHSF compared to HSF indicates that HSF are indeed more

proliferative and secretory myofibroblastic in nature. Furthermore, the decreased

expression of the αSMA gene, along with IL8, TNFα and TRAF2, in nHSF demonstrates that

HSF and nHSF are actually distinct cell populations. Finally, the expression of the CFLAR

C h a p t e r 6 . 0 | 131

gene, a caspase homologue known to induce apoptosis, was also found to be decreased in

nHSF compared to HSF. However, it was anticipated that nHSF would undergo greater

apoptosis compared to HSF, as myofibroblasts are a more proliferative cell type. Why this

difference in CFLAR gene expression is not consistent with our other findings indicating that

HSF are more like proliferative myofibroblasts, is not clear. Importantly, however, these

results, which are summarised in Figure 6.2, suggest that HSF have not reverted back to a

‘normal’ fibroblast phenotype during their in vitro culture.

Figure 6.2 – Differential gene expression between HSF and nHSF and the effects on cellular processes involved in myofibroblast differentiation. Genes in blue indicate those found to be down‐regulated in nHSF compared to HSF while cellular processes in green indicate those important in myofibroblast differentiation. Pathways marked with indicate an increased effect while pathways marked with indicate an inhibitory effect.

The gene expression data reported in Chapter 5.0 is significant as they demonstrate that

the silicone treatments affect gene regulation. However, the studies were only performed

on dermal fibroblasts and the effect of these silicone treatments on gene expression in

keratinocytes, the other main cell type within skin, also needs to be assessed (Werner et

al., 2007). Furthermore, although the results presented are of great interest, gene

expression studies only examine the transcription of genes into RNA and the effects of


transcriptional changes with the cell. Validation at the protein level is therefore also

required since many changes occur during translation, following transcription, and these

influence protein production, conformation and their effects within cells (Chuaqui et al.,

2002). Nevertheless, the studies have provided significant clues regarding which genes and

biological pathways are affected through the application of silicone‐PEG copolymers to

cells, and indeed have implications for the application of silicone‐containing products to

hypertrophic scars. Furthermore, this is the first study to demonstrate a link between the

genes involved in TNF‐induced apoptosis, the genes for members of the TNFR superfamily

and silicone‐PEG copolymers for the treatment of hypertrophic scars.

6.1 LIMITATIONS AND FUTURE DIRECTIONS

Although the experiments within this thesis studied the effects of the silicone treatments

on fibroblasts in detail, it is evident that further experimentation is required to fully

characterise the biological effect of the silicones in vitro, as well as the fibroblasts used in

culture. For example, differential α‐smooth muscle actin expression of normal and scar‐

derived fibroblasts requires assessment at the protein level to investigate the effects of the

silicones and to quantify changes in cell phenotype due to the in vitro culture.

Methodological approaches such as immunocytochemistry or flowcytometry could be

employed to examine this. Expression of α‐smooth muscle actin is characteristic of

myofibroblast phenotype and these investigations are important as the percentage of

myofibroblast cells within the cultures and following treatment with silicone was not

examined within this thesis (Desmouliere et al., 2005). Furthermore, probing the cells for

both α‐smooth muscle actin and annexin V in flow cytometry experiments may be

particularly useful as this will enable the detection of apoptosis in a specific population of

cells. These experiments, among others, are important as it is not yet established how the

silicones affect protein expression and the regulation of biological pathways. In addition,

knowledge regarding keratinocytes and their responses to the treatments also warrants

further research. Given there is a close paracrine relationship between the dermis and

epidermis within skin, further experiments assessing the effects of the silicone treatments

on keratinocytes and dermal fibroblasts co‐cultured together are essential (Martin, 1997).

One of the other limitations of the research reported within this doctoral thesis relates to

the fact that the data were generated in vitro using 2D cell culture techniques. Firstly, cell

culture was performed on fibroblasts isolated and available for purchase from Cell

C h a p t e r 6 . 0 | 133

Research Corpration. Although 2D culture techniques using cells isolated from humans

produces more physiologically relevant results than when immortalised cell lines are used,

they can be hard to perform because the cells are more ‘fragile’ and susceptible to

infection (Freshney, 2010; Pan et al., 2009). Additionally, the fact that the cells used were

purchased from an external supplier means that we had no control over the experimental

techniques used to isolate and initially culture the cells. Additionally, the fibroblasts were

also only available for purchase from passage 4 onwards. While all cell culture experiments

were performed when cells were under passage 10, it is likely that changes in phenotype

occurred during culture (Freshney, 2010).

Another limitation of the studies performed herein is that the optimal concentrations of

the silicone treatments described have been investigated for in vitro use only; the optimal

concentration of each of these silicone treatments for in vivo use, as well as the resulting

effects, may well be quite different. Clearly, future experiments will require the use of 3D

cull culture models, such as the ex vivo 3D human skin equivalent (Regnier et al., 1990; Xie

et al., 2010) and in vivo porcine scar models (Cuttle et al., 2006) reported by others. The 3D

human skin equivalent model is one that utilises de‐epidermised dermis to create a model

of human skin that can be used in fundamental studies of wound repair and in preclinical

investigations of novel wound therapies (Xie et al., 2010). The porcine scar model is an

animal model, in which reproducible hypertrophic scars are created in pigs using deep

dermal partial thickness burns (Cuttle et al., 2006). Importantly, the skin of pigs is known to

be anatomically and physiologically similar to human skin (Meyer et al., 1978; Montagna

and Yun, 1964).

It is expected that further investigation of the silicone treatments using 3D human skin

equivalent models will illustrate decreased cellularity and an induction in apoptosis in the

dermal layer, as well as give additional insight into the effect of the silicone treatments on

keratinocytes within the epidermal layer. The use of the porcine scar models will also allow

the in vivo investigation of the effect of the silicones on both scarred and normal skin. This

is essential as the results obtained using 2D cell culture techniques demonstrated that the

silicone treatments elicited similar effects on both HSF and nHSF. Further advantages of

studies in in vivo models relate to their utility in examining the effect of the silicones on

inflammation; this is important as increased IL8 gene expression was observed in both HSF

and nHSF following treatment. Histological investigations could be used in conjunction with


the porcine scar model, such that inflammatory cells, including neutrophils, monocytes and

mast cells, could be identified by using traditional haematoxylin and eosin staining as well

as the leder stain (Leder, 1979). Finally, qRT‐PCR and immunohistochemistry experiments,

using both 3D human skin equivalent and porcine scar models, would also be employed to

investigate the expression of the genes of interest identified in this thesis at both the gene

and protein level.

Further investigations of the silicone treatments using 3D models are also required to

investigate their mode of action as it has been reported that the in vitro effects of silicone

are not likely to be observed in vivo (1995). This is because silicone oils, which are usually

hydrophobic, have difficulty penetrating the stratum corneum. In fact, it is generally

accepted that SGS are inert and do not instigate biological effects following treatment to

hypertrophic scars (Chang et al., 1995). However, biological effects were clearly evident

when GP226, Fraction IV and PDMS7‐g‐PEG7 were exposed to dermal fibroblasts. More

importantly, it has been demonstrated that these silicones can actually traverse the

stratum corneum (Dickfos, 2008; Gardoni, 2007). Despite these results, it is known that

GP226, Fraction IV and PDMS7‐g‐PEG7, are different to those present in SGS in that they are

amphiphilic silicone‐PEG copolymers with a low molecular weight. Although the

investigated silicones are of different structure to those present in SGS, the results

presented herein raise issues regarding the mechanism by which SGS work. The mechanism

of SGS action is a controversial area that is yet to be completely understood. Clearly, the

hypothesis that SGS can cause biological effects is not able to be dismissed at this time.

6.2 CONCLUSION

In summary, the results reported in this doctoral thesis indicate that low molecular weight

and amphiphilic silicone‐PEG copolymers, such as GP226, Fraction IV and PDMS7‐g‐PEG7,

have potential for use as a scar remediation therapy. It was demonstrated that the

mechanism of action lies in inducing apoptosis in dermal fibroblasts and keratinocytes. This

was further substantiated by gene expression studies in fibroblasts following treatment

with the silicones; these demonstrated that many genes pertinent to apoptosis and

scarring were implicated. More specifically, many genes relating to the TNF‐induced

apoptosis and collagen production pathways were affected. Future studies will therefore

focus on further examining these mechanistic pathways at the protein level in both

fibroblasts and keratinocytes. This will not only clarify the mechanism of silicone action but

C h a p t e r 6 . 0 | 135

will facilitate the optimisation of these treatments prior to clinical trials evaluating these

novel silicones for scar remediation. It is anticipated that this will ultimately assist in the

design of a novel scar therapy with faster and improved outcomes for patients suffering

from hypertrophic scars.

A p p e n d i c i e s

C h a p t e r 7 . 0 | 137

CHAPTER 7.0

APPENDICIES

APPENDIX 1

PDMS‐PEG oligomers were synthesized by Daniel Keddie, Marilla Dickfos, Brooke Farrugia

and Tim Dargaville using methodology described previously (Keddie et al., 2010; submitted;

Dickfos, 2008). Brief descriptions of the methods used are outlined below. All solvents

used were of AR grade and purchased from Sigma‐Aldrich unless otherwise stated.

Synthesis of PEG sidechains

Discrete allyl‐PEG sidechains to be used for block (ABA) PDMS‐PEG oligomers were

synthesized step‐wise starting from diethylene glycol (99%) and diethylene glycol methyl

ether (99%) and coupling these with p‐toluenesulfonyl (98%) activation. When the desired

number of ethylene glycol repeat units had been achieved, allyl groups were added to the

hydroxyl termini. This was achieved by stirring the discrete PEG sidechains with 0.25 mol

equivalent Na+ followed by dropwise addition via cannula to an ice cooled flask under

argon containing 0.25 mol equivalent allyl bromide. Non‐discrete commercially sourced

PEGs (Sigma‐Aldrich) were used to prepare rake PDMS‐PEG oligomers, whereby allyl groups

were added using the same procedure.

Synthesis of silicone (PDMS) backbone

α,ω‐Dihydro(polydimethylsiloxane)s were synthesized via acid catalysed ring‐opening

polymerization of octamethylcyclotetrasiloxane (D4, 99%) and capped with

tetramethyldisiloxane (TMDS, 97%). α,ω‐dihydro(polydimethylsiloxane)s of differing

molecular weight were synthesized by altering the ratio of D4 to TMDS used. For example,

the PDMS10.5 polymer was synthesized using the following process; to a mixture of D4 (30 g,

31.41 mL, 101 mmol), TDMS (13.8 g, 18.16 mL, 102 mmol) and trifluoromethanesulfonic

acid (300 μL, 509 mg, 3.4 mmol, 3.4 mol %) was combined and the reaction mixture stirred

at 100 °C for 5 hours. Following this, the reaction was quenched on NaHCO3 (2 g) with

stirring. The reaction mixture was diluted with diethyl ether in order for filtering to be

carried out, and the ether was subsequently removed under reduced pressure. Kugelrohr


distillation was used to remove the resulting unwanted cyclic and to give the linear PDMS

as a colourless, viscous liquid (25.4 g, Mn = 850).

Hydrosilation of PEG to backbone – block PEG‐PDMS oligomers

To synthesis the block PDMS‐PEG oligomers, discrete allyl‐PEG (3.35 g, 7.9 mmol, ~2.2

equiv.) and α,ω–dihydro(polydimethylsiloxane) (3.05 g, Mn = 850, ~3.58 mmol) were

dissolved in toluene and heated to 90°C under argon in a Schlenk vessel (Sigma‐Aldrich).

Speier’s catalyst (70 µL, 2 % wt/v H2[PtCl6].6H2O in i–PrOH; Sigma‐Aldrich) was then added

and the resulting solution heated at 130 °C in a sealed Schlenk tube for 16 hours. Following

this, activated charcoal was added to the solution and stirred for 16 hours. The product was

filtered and the solvent was removed. The reaction product was purified by an n–pentane

and methanol wash to ensure the block copolymer was a colourless, viscous liquid (4.83 g).

Different block PEG‐PDMS copolymers were obtained by changing the PEG or PDMS

reagents to generate the desired products.

Hydrosilation of PEG to backbone – rake PEG – PDMS oligomers

To synthesise the rake PDMS‐PEG oligomers, Karstedt’s catalyst (Sigma‐Aldrich) was

combined with a solution of non discrete allyl‐PEG (0.08 g, 2.2x10‐4 mol, ~1 equiv) and α,ω–

dihydro(polydimethylsiloxane) (0.055 g, 7x10‐5 mol, ~0.3 equiv) in tetrahydrofuran (THF).

The resulting reaction mixture was stirred for 3 hours at 80 ˚C. Following cooling, activated

charcoal (0.1 g) was added and the mixture stirred at room temperature overnight. The

solution was then filtered and the (THF) evaporated.

C h a p t e r 7 . 0 | 139

APPENDIX 2

Treatment

time Untreated 0.01% GP226

0.03%

GP226 0.1% GP226 0.3% GP226 1% GP226

Viability

24 hours 1.27 ± 0.08 1.25 ± 0.17 1.19 ± 0.16 1.19 ± 0.17 1.31 ± 0.22 1.27 ± 0.35

48 hours 1.70 ± 0.16 1.51 ± 0.10 1.27 ± 0.08 1.10 ± 0.11 0.83 ± 0.19 0.23 ± 0.05

72 hours 1.95 ± 0.17 1.42 ± 0.03 1.37 ± 0.09 1.23 ± 0.06 0.84 ± 0.19 0.11 ± 0.03

168 hours 2.03 ± 0.05 1.22 ± 0.03 1.09 ± 0.05 0.99 ± 0.06 0.03 ± 0.02 0.05 ± 0.01

Proliferation 24 hours 1779 ± 46.1 1538 ± 99.9 1371 ± 79.7 1304 ± 100 1192 ± 73.7 1073 ± 90.2

48 hours 2641 ± 87.0 1855 ± 167 1572 ± 115 1450 ± 116 1292 ± 133 967.7 ± 121

72 hours 2939 ± 120 1911 ± 172 1549 ± 167 1420 ± 113 1096 ± 123 801.8 ± 123

168 hours 3380 ± 102 1889 ± 88.1 1519 ± 103 1383 ± 106 620.3 ± 109 110.0 ± 30.1

Table A2.1 – Numerical Values of HSF Viability and Proliferation following treatment with GP226 for 24, 48, 72 and 168 hours. Data are expressed as an average of the true values (Viability, absorbance at 620 nm, ref at 620 nm; Proliferation, Fluorescence, ex at 480 nm, em at 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate.

Treatment

time Untreated

0.01%

Fraction I

0.03%

Fraction I

0.1%

Fraction I

0.3%

Fraction I

1%

Fraction I

Viability

24 hours 1.27 ± 0.08 1.15 ± 0.11 1.19 ± 0.10 1.16 ± 0.09 1.11 ± 0.13 0.55 ± 0.27

48 hours 1.70 ± 0.16 1.84 ± 0.19 1.87 ± 0.22 1.91 ± 0.22 1.73 ± 0.30 1.12 ± 0.56

72 hours 1.95 ± 0.17 2.25 ± 0.19 2.03 ± 0.27 1.83 ± 0.14 2.00 ± 0.21 1.00 ± 0.48

168 hours 2.03 ± 0.05 1.89 ± 0.07 1.80 ± 0 08 1.50 ± 0.10 1.37 ± 0.14 0.62 ± 0.33

Proliferation 24 hours 1779 ± 46.1 1700 ± 91.5 1724 ± 109 1607 ± 135 1452 ± 164 1125 ± 200

48 hours 2641 ± 87.0 2737 ± 165 2766 ± 156 2326 ± 208 1928 ± 298 1579 ± 246

72 hours 2939 ± 120 3000 ± 168 2897 ± 207 2466 ± 270 1800 ± 333 1337 ± 278

168 hours 3380 ± 102 3252 ± 141 3323 ± 110 2662 ± 194 1701 ± 358 1367 ± 237

Table A2.2 – Numerical Values of HSF Viability and Proliferation following treatment with Fraction I for 24, 48, 72 and 168 hours. Data are expressed as an average of the true values (Viability, absorbance at 620 nm, ref at 620 nm; Proliferation, Fluorescence, ex at 480 nm, em at 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate.


Treatment

time Untreated

0.01%

Fraction II

0.03%

Fraction II

0.1%

Fraction II

0.3%

Fraction II

1%

Fraction II

Viability

24 hours 1.27 ± 0.08 1.16 ± 0.11 1.18 ± 0.12 1.21 ± 0.11 1.37 ± 0.15 0.84 ±0.39

48 hours 1.70 ± 0.16 1.87 ± 0.22 1.82 ± 0.20 1.91 ± 0.17 2.17 ± 0.17 1.22 ± 0.59

72 hours 1.95 ± 0.17 1.99 ± 0.23 1.77 ± 0.16 2.04 ± 0.15 2.38 ± 0.14 1.22 ± 0.58

168 hours 2.03 ± 0.05 1.89 ± 0.07 1.73 ± 0.06 1.74 ± 0.07 1.84 ± 0.11 0.76 ± 0.36

Proliferation 24 hours 1779 ± 46.1 1853 ± 83.1 1818 ± 98.7 1724 ± 108 1506 ± 144 1244 ± 217

48 hours 2641 ± 87.0 2830 ± 134 2582 ± 167 2336 ± 147 2081 ± 208 1598 ± 335

72 hours 2939 ± 120 2910 ± 141 2805 ± 146 2347 ± 147 1930 ± 258 1416 ± 357

168 hours 3380 ± 102 3420 ± 178 3080 ± 155 2540 ± 95.5 2166 ± 153 1392 ± 361

Table A2.3 – Numerical Values of HSF Viability and Proliferation following treatment with Fraction II for 24, 48, 72 and 168 hours. Data are expressed as an average of the true values (Viability, absorbance at 620 nm, ref at 620 nm; Proliferation, Fluorescence, ex at 480 nm, em at 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate.

Treatment

time Untreated

0.01%

Fraction III

0.03%

Fraction III

0.1%

Fraction III

0.3%

Fraction III

1%

Fraction III

Viability

24 hours 1.27 ± 0.08 1.17 ± 0.12 1.20 ± 0.13 1.30 ± 0.09 1.25 ± 0.09 1.09 ± 0.15

48 hours 1.70 ± 0.16 1.75 ± 0.22 1.75 ± 0.20 1.81 ± 0.19 1.32 ± 0.08 1.01 ± 0.22

72 hours 1.95 ± 0.17 2.02 ± 0.25 1.92 ± 0.16 1.53 ± 0.22 1.45 ± 0.03 1.11 ± 0.22

168 hours 2.03 ± 0.05 2.11 ± 0.04 1.85 ± 0.07 1.60 ± 0.15 1.23 ± 0.09 0.94 ± 0.20

Proliferation 24 hours 1779 ± 46.1 1853 ± 78.7 1832 ± 81.5 1697 ± 87.1 1454 ± 108 1063 ± 184

48 hours 2641 ± 87.0 2821 ± 158 2696 ± 128 2275 ± 153 1955 ± 252 1383 ± 265

72 hours 2939 ± 120 2952 ± 111 2650 ± 194 2093 ± 158 1707 ± 150 1112 ± 196

168 hours 3380 ± 102 3425 ± 166 3227 ± 168 2187 ± 139 1827 ± 34.7 1137 ± 230

Table A2.4 – Numerical Values of HSF Viability and Proliferation following treatment with Fraction III for 24, 48, 72 and 168 hours. Data are expressed as an average of the true values (Viability, absorbance at 620 nm, ref at 620 nm; Proliferation, Fluorescence, ex at 480 nm, em at 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate.

C h a p t e r 7 . 0 | 141

Treatment

time Untreated

0.01%

Fraction IV

0.03%

Fraction IV

0.1%

Fraction IV

0.3%

Fraction IV

1%

Fraction IV

Viability

24 hours 1.27 ± 0.08 1.11 ± 0.10 1.13 ± 0.11 1.19 ± 0.09 1.09 ± 0.08 0.016 ± 0.01

48 hours 1.70 ± 0.16 1.48 ± 0.18 1.50 ± 0.18 1.43 ± 0.12 1.02 ± 0.03 0.003 ± 0.01

72 hours 1.95 ± 0.17 1.55 ± 0.07 1.63 ± 0.09 1.47 ± 0.02 1.14 ± 0.01 0.011 ± 0.01

168 hours 2.03 ± 0.05 1.21 ± 0.09 1.25 ± 0.14 1.30 ± 0.10 0.91 ± 0.05 ‐0.01 ± 0.01

Proliferation 24 hours 1779 ± 46.1 1569 ± 105 1537 ± 105 1440 ± 95.4 1132 ± 62.5 406.3 ± 39.4

48 hours 2641 ± 87.0 2077 ± 184 2094 ± 219 1884 ± 148 1399 ± 104 578.7 ± 102

72 hours 2939 ± 120 2034 ± 127 1771 ± 140 1942 ± 143 1406 ± 96.3 324.9 ± 45.1

168 hours 3380 ± 102 2022 ± 71.5 1918 ± 121 2052 ± 68.9 1501 ± 25.5 137.4 ± 19.3

Table A2.5 – Numerical Values of HSF Viability and Proliferation following treatment with Fraction IV for 24, 48, 72 and 168 hours. Data are expressed as an average of the true values (Viability, absorbance at 620 nm, ref at 620 nm; Proliferation, Fluorescence, ex at 480 nm, em at 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate.

Treatment

time Untreated

0.01%

Fraction V

0.03%

Fraction V

0.1%

Fraction V

0.3%

Fraction V

1%

Fraction V

Viability

24 hours 1.27 ± 0.08 1.14 ± 0.14 1.01 ± 0.05 0.01 ± 0.01 0.02 ± 0.01 0.03 ± 0.01

48 hours 1.70 ± 0.16 1.22 ± 0.06 1.11 ± 0.03 0.06 ± 0.05 0.02 ± 0.01 0.02 ± 0.01

72 hours 1.95 ± 0.17 1.54 ± 0.12 1.37 ± 0.04 0.01 ± 0.01 0.02 ± 0.02 0.03 ± 0.02

168 hours 2.03 ± 0.05 1.11 ± 0.10 1.17 ± 0.11 ‐0.03 ± 0.01 0.00 ± 0.01 ‐0.02 ± 0.01

Proliferation 24 hours 1779 ± 46.1 1203 ± 72.2 1298 ± 67.3 590.2 ± 90.6 363.0 ± 94.0 1061 ± 32.6

48 hours 2641 ± 87.0 1529 ± 110 1809 ± 77.0 738.8 ± 82.0 506.2 ± 106 708.0 ± 127

72 hours 2939 ± 120 1453 ± 98.0 1803 ± 64.2 677.8 ± 87.7 278.9 ± 38.9 349.0 ± 101

168 hours 3380 ± 102 1427 ± 59.2 1918 ± 99.2 766.0 ± 181 158.2 ± 17.4 127.0 ± 53.9

Table A2.6 – Numerical Values of HSF Viability and Proliferation following treatment with Fraction V for 24, 48, 72 and 168 hours. Data are expressed as an average of the true values (Viability, absorbance at 620 nm, ref at 620 nm; Proliferation, Fluorescence, ex at 480 nm, em at 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate.


Treatment

time Untreated 0.01% PEG 0.03% PEG 0.1% PEG 0.3% PEG 1% PEG

Viability

24 hours 1.27 ± 0.08 1.20 ± 0.11 1.24 ± 0.12 1.22 ± 0.12 1.278 ± 0.14 1.29 ± 0.11

48 hours 1.70 ± 0.16 1.80 ± 0.22 1.82 ± 0.20 1.83 ± 0.22 1.86 ± 0.22 2.02 ± 0.19

72 hours 1.95 ± 0.17 1.99 ± 0.22 2.00 ± 0.26 1.88 ± 0.24 2.03 ± 0.21 2.02 ± 0.17

168 hours 2.03 ± 0.05 1.97 ± 0.05 1.82 ± 0.07 1.78 ± 0.07 2.00 ± 0.09 2.11 ± 0.07

Proliferation 24 hours 1779 ± 46.1 1772 ± 96.0 1671 ± 76.6 1617 ± 104 1739 ± 67.3 1722 ± 93.3

48 hours 2641 ± 87.0 2719 ± 180 2527 ± 174 2634 ± 139 2641 ± 112 2724 ± 169

72 hours 2939 ± 120 2828 ± 137 2766 ± 153 2748 ± 157 2884 ± 145 2979 ± 177

168 hours 3380 ± 102 3260 ± 135 3354 ± 257 3026 ± 282 3483 ± 155 3871 ± 119

Table A2.7 – Numerical Values of HSF Viability and Proliferation following treatment with PEG for 24, 48, 72 and 168 hours. Data are expressed as an average of the true values (Viability, absorbance at 620 nm, ref at 620 nm; Proliferation, Fluorescence, ex at 480 nm, em at 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate.

Treatment

time Untreated

0.01%

PDMS 0.03% PDMS

0.1%

PDMS

0.3%

PDMS

1%

PDMS

Viability

24 hours 1.27 ± 0.08 1.12 ± 0.09 1.15 ± 0.11 1.19 ± 0.12 1.17 ± 0.12 1.21 ± 0.11

48 hours 1.70 ± 0.16 1.80 ± 0.23 1.75 ± 0.24 1.76 ± 0.23 1.79 ± 0.21 1.76 ± 0.22

72 hours 1.95 ± 0.17 1.95 ± 0.21 1.87 ± 0.18 1.91 ± 0.18 1.93 ± 0.22 1.96 ± 0.19

168 hours 2.03 ± 0.05 1.66 ± 0.06 1.74 ± 0.04 1.82 ± 0.04 1.62 ± 0.11 1.64 ± 0.05

Proliferation 24 hours 1779 ± 46.1 1521 ± 146 1548 ± 119 1550 ± 128 1592 ± 152 1568 ± 124

48 hours 2641 ± 87.0 2077 ± 250 2120 ± 256 2228 ± 279 2323 ± 303 2151 ± 282

72 hours 2939 ± 120 2861 ± 282 2840 ± 276 2800 ± 270 2919 ± 239 2923 ± 265

168 hours 3380 ± 102 2960 ± 147 2943 ± 115 3022 ± 181 3176 ± 148 3065 ± 101

Table A2.8 – Numerical Values of HSF Viability and Proliferation following treatment with PDMS for 24, 48, 72 and 168 hours. Data are expressed as an average of the true values (Viability, absorbance at 620 nm, ref at 620 nm; Proliferation, Fluorescence, ex at 480 nm, em at 520 nm) pooled from three separate experiments, in which each treatment was performed in triplicate.

C h a p t e r 7 . 0 | 143

APPENDIX 3

For Appendix 3, please refer to the folder named Appendix 3 on the attached DVD.

APPENDIX 4

For Appendix 4, please refer to the file named Appendix 4 on the attached DVD.

R e f e r e n c e s

C h a p t e r 8 . 0 | 145

CHAPTER 8.0

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