pre clinical evaluation of amphiphilic silicone...
TRANSCRIPT
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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
3.2.1 Fractionation of GP226 40
3.2.3 Fibroblast Cell Culture 40
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3.2.4 Keratinocyte Cell Culture 40
3.2.5 WST‐1 Assay for Determination of Cell Viability 41
3.2.6 Cyquant Assay for Determination of Cell Proliferation 41
3.2.7 Analysis of Silicone and Protein Interaction 41
3.2.8 Analysis of Cell Morphology via Real‐Time Microscopy 42
3.2.9 Tunel Assay for the Detection of Apoptosis 42
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
4.2 Experimental Procedures 68
4.2.1 Synthesis of PDMS‐PEG oligomers 68
4.2.2 Fibroblast Cell Culture 68
4.2.3 Keratinocyte Cell Culture 68
4.2.4 Cyquant Assay for Determination of Cell Proliferation 68
4.2.5 Analysis of Cell Morphology via Real‐Time Microscopy 69
4.2.6 Tunel Assay for the Detection of Apoptosis 69
4.2.7 Statistical Analysis 69
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 Experimental Procedures 92
5.2.1 Fractionation of GP226 92
5.2.2 Synthesis of PDMS‐PEG oligomers 92
5.2.3 Fibroblast Cell Culture 92
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
5.2.14 Statistical Analysis 95
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
L i t e r a t u r e R e v i e w
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
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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
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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
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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
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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
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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
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(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
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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,
L i t e r a t u r e R e v i e w
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
L i t e r a t u r e R e v i e w
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 variable was tested in triplicate
Each assay was repeated with cells from three different patients.
Each variable was tested in triplicate
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
L i t e r a t u r e R e v i e w
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
M a t e r i a l s a n d M e t h o d s
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
M a t e r i a l s a n d M e t h o d 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.
M a t e r i a l s a n d M e t h o d s
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
M a t e r i a l s a n d M e t h o d s
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
M a t e r i a l s a n d M e t h o d s
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.
M a t e r i a l s a n d M e t h o d s
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
M a t e r i a l s a n d M e t h o d s
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.
3.2.1 Fractionation of GP226
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.
3.2.3 Fibroblast Cell Culture
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.
3.2.4 Keratinocyte Cell Culture
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.
3.2.6 Cyquant Assay for Determination of Cell Proliferation
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
untreated control, containing cells treated with medium but no silicone; and negative
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
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
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.
3.2.8 Analysis of Cell Morphology via Real‐Time Microscopy
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.
3.2.9 Tunel Assay for the Detection of Apoptosis
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.
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
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.
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
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.
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
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.
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
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.
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
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.
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
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.
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
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.
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
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.
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
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).
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
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.
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
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.
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
4.2 EXPERIMENTAL PROCEDURES
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.
4.2.1 Synthesis of PDMS‐PEG oligomers
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.
4.2.2 Fibroblast Cell Culture
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.
4.2.3 Keratinocyte Cell Culture
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.
4.2.4 Cyquant Assay for Determination of Cell Proliferation
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
untreated control, containing cells treated with medium but no silicone; and negative
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.
4.2.5 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, 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.
4.2.6 Tunel Assay for the Detection of Apoptosis
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.
4.2.7 Statistical Analysis
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
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
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.
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
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.
Untreated 3687 ± 195.8
Treatment GP226 PDMS7‐g‐PEG7 PDMS7
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
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
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
Treatment GP226 PDMS7‐g‐PEG7 PDMS7
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
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
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.
Untreated 2165 ± 124.2
Treatment GP226 PDMS7‐g‐PEG7 PDMS7
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
C h a p t e r 4 . 0 | 77
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.
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
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.
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
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.
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
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
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
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%).
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
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;
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
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)
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
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.
5.2 EXPERIMENTAL PROCEDURES
5.2.1 Fractionation of GP226
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.
5.2.2 Synthesis of PDMS‐PEG oligomers
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.
5.2.3 Fibroblast Cell Culture
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.
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
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.
5.2.14 Statistical Analysis
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.
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
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).
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
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).
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
Comparisons Increased Decreased
Superarray
(84 genes)
HSF vs nHSF 3 3
HSF Un vs 0.01% PDMS7‐g‐PEG7 (48 hours) 9 7
HSF Un vs 0.03% PDMS7‐g‐PEG7 (6 hours) 4 27
nHSF Un vs 0.03% PDMS7‐g‐PEG7 (48 hours) 16 12
HSF Un vs 0.03% PDMS7‐g‐PEG7 (6 hours) 9 13
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.
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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
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
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.
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
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.
C h a p t e r 5 . 0 | 105
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.
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
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.
C h a p t e r 5 . 0 | 107
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.
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
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.
C h a p t e r 5 . 0 | 109
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.
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
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.
C h a p t e r 5 . 0 | 111
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.
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
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
C h a p t e r 5 . 0 | 113
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
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
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
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
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
C h a p t e r 5 . 0 | 117
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
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
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
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
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
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
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
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
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
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
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
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
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
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
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
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 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
A p p e n d i c i e s
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.
A p p e n d i c i e s
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.
A p p e n d i c i e s
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.
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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.
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CHAPTER 8.0
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