investigations on keloid pathogenesis and therapy · interactions in keloid scar pathogenesis and...
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i
INVESTIGATIONS ON KELOID PATHOGENESIS AND THERAPY
Anandaroop Mukhopadhyay (B.Pharm), The Tamilnadu Dr M.G.R Medical University, INDIA)
THESIS SUBMITTED FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
DEPARTMENT OF PHARMACY
NATIONAL UNIVERSITY OF SINGAPORE
2007
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ACKNOWLEDGEMENT
It is my pleasure to thank the many people who made this thesis possible.
To start with, I would like to thank my Ph.D. supervisor Associate Professor Chan Sui
Yung for her excellent guidance and support. Had it not been for her, I wouldn’t have had
the courage to explore new horizons. Her critical evaluation of my research and constant
encouragement invigorated me to push the limits.
It is difficult to overstate my gratitude to my co-supervisor Assistant Professor Phan Toan
Thang. He took me in his research team knowing fully well that I had no experience in
skin biology. I thank him for having confidence in me and providing me the opportunity
to be a part of the cutting edge research in skin pathogenesis in his laboratory. With his
enthusiasm, his inspiration, and his great efforts to explain things clearly and simply, he
helped to make research fun for me. Throughout the period of writing my thesis, he
provided encouragement, sound advice, good teaching, good company, and lots of good
ideas. I would have been lost without him.
I am deeply indebted and thankful to my parents Subhash Chandra Mukhopadhyay and
Gopa Mukhopadhyay for instilling in me, values, that helped me comprehend the
importance of knowledge and pursue it. My brothers Abhiroop Mukhopadhyay and
Ritobrata Banerjee have always been my mentors and I am thankful to them for their
invaluable advice. I would also like to thank my uncle Dr M.G Banerjee for always
believing in me, and the rest of my family members for their support. Finally I am
grateful to all my friends and lab mates for making my graduate life enjoyable.
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TABLE OF CONTENTS
Acknowledgement
List of Publications
List of Figures
Abbreviations
Summary
1 BACKGROUND AND INTRODUCTION........................................................................................4 1.1 OVERVIEW OF THE WOUND HEALING PROCESS..............................................................5 1.2 KELOID SCAR............................................................................................................................7
1.2.1 Clinical Characteristics of Keloid ...........................................................................................7 1.2.2 Epidemiology...........................................................................................................................9 1.2.3 Histopathology ........................................................................................................................9 1.2.4 Molecular Pathogenesis ........................................................................................................11 1.2.5 Current Treatment .................................................................................................................16
1.3 OBJECTIVES OF THE PRESENT STUDY..............................................................................19
2 MATERIALS AND METHODS ......................................................................................................21 2.1 MEDIA AND CHEMICALS .....................................................................................................22 2.2 RECOMBINANT GROWTH FACTORS AND ANTIBODIES................................................22 2.3 PREPARATION OF NORMAL AND KELOID TISSUE EXTRACTS..............23 2.4 IMMUNOHISTOCHEMISTRY ................................................................................................24
2.4.1 Preparation of Paraffin Sections of Normal and Keloid Tissues...........................................24 2.4.2 Pretreatment of Paraffin Sections for Immunohistochemistry...............................................24 2.4.3 Probing with Antibody and Developing ................................................................................25
2.5 CELL CULTURE.......................................................................................................................25 2.5.1 Keloid Keratinocyte and Fibroblast Database......................................................................25 2.5.2 Keloid and Normal Keratinocytes from Keloid scar and Normal Skin .................................26 2.5.3 Keloid and Normal Fibroblast from Keloid Scar and Normal Skin ......................................26 2.5.4 Keratinocyte-fibroblast Coculture.........................................................................................27
2.6 CELL COUNTING ....................................................................................................................29 2.7 TREATMENT OF FIBROBLASTS WITH GROWTH FACTOR.............................................29 2.8 TREATMENT OF CELLS WITH GLEEVEC...........................................................................29 2.9 SERUM STIMULATION..........................................................................................................30 2.10 MTT ASSAY .............................................................................................................................30 2.11 FIBROBLAST-POPULATED COLLAGEN LATTICE (FPCL) ..............................................31
2.11.1 Preparation of Fibroblast-Populated Collagen Lattices ..................................................31 2.11.2 Macroscopic Evaluation of FPCL Contraction ................................................................32
2.12 WESTERN BLOTTING ............................................................................................................32 2.13 IMMUNOASSAY......................................................................................................................33
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2.14 FLOW CYTOMETRY...............................................................................................................33 2.15 RNASE PROTECTION ASSAY.................................................................................................34 2.16 RATE OF ATP SYNTHESIS.....................................................................................................34 2.17 MEASUREMENT OF INTRACELLULAR ATP......................................................................35 2.18 STATISTICAL ANALYSIS ......................................................................................................35
3 ROLE OF EPITHELIAL-MESENCHYMAL INTERACTIONS IN WOUND CONTRACTION AND SCAR CONTRACTURE ..................................................................................................................36
3.1 INTRODUCTION......................................................................................................................37 3.2 RESULTS ..................................................................................................................................39 3.3 DISCUSSION ............................................................................................................................46
4 ROLE OF ACTIVIN-FOLLISTATIN SYSTEM IN KELOID PATHOGENESIS.....................50 4.1 INTRODUCTION......................................................................................................................51 4.2 RESULTS ..................................................................................................................................53 4.3 DISCUSSION ............................................................................................................................67
5 ROLE OF PROTEOGLYCANS IN KELOID PATHOGENESIS................................................73 5.1 INTRODUCTION......................................................................................................................74 5.2 RESULTS ..................................................................................................................................77 5.3 DISCUSSION ............................................................................................................................94
6 ROLE OF STEM CELL FACTOR AND C-KIT SYSTEM IN KELOID PATHOGENESIS..101 6.1 INTRODUCTION....................................................................................................................102 6.2 RESULTS ................................................................................................................................104 6.3 DISCUSSION ..........................................................................................................................117
7 INVESTIGATION OF GLEEVEC AS A THERAPEUTIC AGENT FOR KELOID SCARS 124 7.1 INTRODUCTION....................................................................................................................125 7.2 RESULTS ................................................................................................................................127 7.3 DISCUSSION ..........................................................................................................................134
General Conclusion
Appendix 1
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LIST OF PUBLICATIONS
Publication Papers
1. Mukhopadhyay A, Tan EK, Khoo YT, Chan SY, Lim IJ, Phan TT. Conditioned medium
from keloid keratinocyte/keloid fibroblast coculture induces contraction of fibroblast-
populated collagen lattices. Br J Dermatol 2005 Apr; 152(4):639-45.
2. Phan TT, Lim IJ, Aalami O, Lorget F, Khoo A, Tan EK, Mukhopadhyay A, Longaker
MT Smad3 signaling plays an important role in keloid pathogenesis via epithelial-
mesenchymal interactions. J Pathol 2005 Oct; 207(2):232-42.
3. Khoo A, Ong CT, Tan EK, Mukhopadhyay A, IJ Lim, TT Phan. The role of Connective
Tissue Growth Factor (CTGF) in the biology of epithelial-mesenchymal interactions of
keloid pathogenesis. J Cell Physiol 2006 Aug; 208(2):336-43
4. Mukhopadhyay A, Chan SY, Lim IJ, Phillips DJ, Phan TT. The role of Activin System in
Keloid Pathogenesis. Am J Physiol-Cell Physiol 2006 Sep 13(in press)
5. Ong CT, Khoo A, Tan EK, Mukhopadhyay A, Do DV, Han CH, IJ Lim, TT Phan.
Epithelial-mesenchymal interactions in keloid pathogenesis modulate vascular
endothelial growth factor expression and secretion. J Pathol 2007; 211:95-108
6. Do DV, Mukhopadhyay A, Lim IJ, Phan TT. Roles of epithelial-mesenchymal
interactions in keloid scar pathogenesis and carcinogenesis. A review. Current Signal
Transduction Therapy (invited review paper, in press).
7. Mukhopadhyay A, Fan S, Do DV, Khoo A, Ong CT, Lim IJ, Phan TT. Role of HGF /c-
Met system in keloid pathogenesis. Br J Dermatol, (in press).
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Submitted manuscripts
1. Mukhopadhyay A, Khoo A, Chan SY, Aalami O, Lim IJ, Phan TT. Specific Target of
Sp1 Transcription Factor: A Novel Therapeutic Approach for Keloids.
2. Mukhopadhyay A, Wong MY, Chan SY , Do DV , Khoo A , Ong CT , Cheong HH , Lim
IJ , Phan TT. Syndecan-2 and Decorin - Proteoglycans with a difference: Implications in
keloid pathogenesis.
Presentations
1. NHG Annual Scientific Congress, Singapore, October 2004
2. AAPS (American Association of Pharmaceutical Scientists) Annual Meeting and
Exposition, Baltimore, USA, November 2004
3. 17th Pharmacy Congress and Intervarsity Symposium, Singapore, July 2005
4. 3rd Meeting of the WHSS (Wound Healing Society of Singapore), Singapore, August
2005
5. Inaugural AAPS-NUS Student Chapter Symposium, Singapore, September 2005
6. Controlled Release Society Conference, Vienna, Austria, July 2006
7. Biostar Congress, Stuttgart, Germany, October 2006
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LIST OF FIGURES
FIGURE 1: SCHEMATIC REPRESENTATION OF DIFFERENT STAGES OF WOUND REPAIR (ADAPTED FROM
WERNER ET AL., 2003). ...........................................................................................................................6
FIGURE 2: KELOID FORMATION IN DIFFERENT PATIENTS (ADAPTED FROM MARNEROS ET AL., 2004)...............8
FIGURE 3: H&E STAINING OF NORMAL SKIN AND KELOID SCAR TISSUE .........................................................10
FIGURE 4: SCHEMATIC REPRESENTATION OF PATHWAYS POTENTIALLY RESULTING IN ACCUMULATION OF
COLLAGEN IN FIBROTIC SKIN DISEASES (ADAPTED FROM UITTO ET AL., 2007)......................................16
FIGURE 5: COCULTURE OF EPIDERMAL KERATINOCYTES AND DERMAL FIBROBLASTS AS AN IN VITRO MODEL
TO STUDY EPITHELIAL-MESENCHYMAL INTERACTION.....................ERROR! BOOKMARK NOT DEFINED.
FIGURE 6: EFFECT OF CONDITIONED MEDIA COLLECTED FROM KELOID KERATINOCYTE (KK)/KELOID
FIBROBLAST (KF) CO CULTURES ON COLLAGEN CONTRACTION BY KFS ...............................................42
FIGURE 7: COMPARISON OF CONTRACTION OF COLLAGEN GEL LATTICE INCORPORATED WITH KELOID
FIBROBLASTS (KFS) AND NORMAL FIBROBLASTS (NFS). ......................................................................43
FIGURE 8: Α-SMOOTH MUSCLE ACTIN (Α-SMA) EXPRESSION BY KELOID FIBROBLASTS (KFS) (A) OR NORMAL
DERMAL FIBROBLASTS (NFS) (B). .........................................................................................................44
FIGURE 9: EFFECT OF ANTI TRANSFORMING GROWTH FACTOR (TGF)-Β1 NEUTRALIZING ANTIBODY ON
COLLAGEN CONTRACTION BY KELOID FIBROBLAST (KF) STRAIN 48 (KF48) INDUCED BY CONDITIONED
MEDIA COLLECTED FROM KELOID KERATINOCYTE (KK) STRAIN 48 (KK48)/KF48 COCULTURE. .........45
FIGURE 10: INCREASED LOCALIZATION OF ACTIVIN-A AND FOLLISTATIN IN THE BASAL LAYER OF EPIDERMIS
IN NORMAL AND KELOID TISSUES..........................................................................................................57
FIGURE 11: EXPRESSION OF ENDOGENOUS ACTIVIN-ΒA MRNA DERIVED FROM NORMAL AND KELOID TISSUES.
.............................................................................................................................................................58
FIGURE 12: ELEVATED LEVELS OF ACTIVIN-A OBTAINED FROM KELOID FIBROBLAST CULTURES...................59
FIGURE 13: INCREASED PROLIFERATION OF NORMAL FIBROBLASTS WHEN COCULTURED WITH ACTIVIN-A
OVEREXPRESSING HΒAHACAT CELLS. .................................................................................................60
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FIGURE 14: INCREASED PROLIFERATION OF NORMAL AND KELOID FIBROBLASTS TREATED WITH RHACTIVIN-
A. .........................................................................................................................................................61
FIGURE 15: EFFECT OF ACTIVIN-A, FOLLISTATIN OR TGF-Β1 ON COLLAGEN, FIBRONECTIN AND Α-SMA
EXPRESSION..........................................................................................................................................63
FIGURE 16: ACTIVIN-A EXPRESSION INCREASED BY EPITHELIAL-MESENCHYMAL INTERACTIONS RESULTING
IN A FIBROTIC PHENOTYPE. ...................................................................................................................72
FIGURE 17: ELEVATED LEVELS OF SYNDECAN-2 IN TISSUE EXTRACTS OBTAINED FROM KELOID TISSUE. .......81
FIGURE 18: SERUM GROWTH FACTORS UPREGULATED SYNDECAN-2 EXPRESSION IN NORMAL AND KELOID
FIBROBLASTS. .......................................................................................................................................82
FIGURE 19: INCREASED ECTODOMAIN SHEDDING OF SYNDECAN-2 IS OBSERVED IN KELOID COCULTURES AS
COMPARED TO MONOCULTURES............................................................................................................83
FIGURE 20: EXOGENOUS RHFGF-2 STIMULATES SHEDDING OF SYNDECAN-2 FROM FIBROBLAST CELL
SURFACE INTO THE CONDITIONED MEDIA..............................................................................................84
FIGURE 21: INCREASED LOCALIZATION OF FGF-2 IN THE BASAL LAYER OF EPIDERMIS AND DERMIS IN KELOID
TISSUE. .................................................................................................................................................85
FIGURE 22: ELEVATED LEVELS OF FGF-2 IN TISSUE EXTRACTS OBTAINED FROM KELOID TISSUE. .................86
FIGURE 23: COCULTURED NORMAL AND KELOID KERATINOCYTES EXPRESS INCREASED LEVELS OF FGF-2 AS
COMPARED TO MONOCULTURED KERATINOCYTES. ...............................................................................87
FIGURE 24: COCULTURED NORMAL AND KELOID FIBROBLASTS EXPRESS INCREASED LEVELS OF FGF-2 AS
COMPARED TO CELLS IN MONOCULTURE...............................................................................................88
FIGURE 25: DOWNREGULATION OF DECORIN IN TISSUE EXTRACTS OBTAINED FROM KELOID TISSUE..............89
FIGURE 26: SERUM GROWTH FACTORS DOWNREGULATE DECORIN EXPRESSION IN NORMAL AND KELOID
FIBROBLASTS. .......................................................................................................................................90
FIGURE 27: INCREASED SECRETORY DECORIN IN COCULTURED NORMAL FIBROBLASTS AND NOT KELOID
FIBROBLASTS. .......................................................................................................................................91
FIGURE 28: EFFECT OF DECORIN ON COLLAGEN, FIBRONECTIN AND Α-SMA EXPRESSION..............................92
FIGURE 29: ECTODOMAIN SHEDDING OF SYNDECAN-2 BY FGF-2 STIMULATED BY EPITHELIAL-MESENCHYMAL
INTERACTIONS RESULTING IN A FIBROTIC PHENOTYPE........................................................................100
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FIGURE 30: ELEVATED LEVELS OF SCF IN TISSUE EXTRACTS OBTAINED FROM KELOID TISSUE. ...................107
FIGURE 31: SERUM GROWTH FACTORS UPREGULATED SCF EXPRESSION IN NORMAL AND KELOID
FIBROBLASTS. .....................................................................................................................................108
FIGURE 32: INCREASED LEVELS OF SCF IN CONDITIONED MEDIA OBTAINED FROM KELOID COCULTURES AS
COMPARED TO MONOCULTURES..........................................................................................................109
FIGURE 33: ELEVATED LEVELS OF C-KIT AND PHOSPHO C-KIT (TYR 703, TYR 721) IN TISSUE EXTRACTS
OBTAINED FROM KELOID TISSUE. ........................................................................................................110
FIGURE 34: INCREASED LOCALIZATION OF C-KIT IN THE BASAL LAYER OF EPIDERMIS OF KELOID TISSUE. ..111
FIGURE 35: C-KIT UPREGULATED IN KELOID KERATINOCYTES AS COMPARED TO NORMAL KERATINOCYTES.
...........................................................................................................................................................112
FIGURE 36: INCREASED ECTODOMAIN SHEDDING OF C-KIT IN KELOID KERATINOCYTE/KELOID FIBROBLAST
COCULTURES. .....................................................................................................................................112
FIGURE 37: TACE OVEREXPRESSED IN KELOID SCAR AS COMPARED TO NORMAL SKIN................................115
FIGURE 38: UPREGULATION OF TACE IN KELOID FIBROBLAST COCULTURES AS COMPARED TO NORMAL
COCULTURES. .....................................................................................................................................116
FIGURE 39: C-KIT SYSTEM IN KELOID PATHOGENESIS..................................................................................122
FIGURE 40: NO SIGNIFICANT DIFFERENCE IN THE PROLIFERATIVE POTENTIAL OF KELOID KERATINOCYTES
AND KELOID FIBROBLASTS..................................................................................................................129
FIGURE 41: GLEEVEC DOWNREGULATES EXPRESSION OF PHOSPHO AKT, PHOSPHO MTOR, PHOSPHO C-KIT
(TYR 721). ..........................................................................................................................................130
FIGURE 42: GLEEVEC DOWNREGULATES EXPRESSION OF COLLAGEN, FIBRONECTIN, Α-SMA, VEGF, HDGF,
TGF-Β1, SCF, FGF-2. ........................................................................................................................131
FIGURE 43: GLEEVEC SIGNIFICANTLY REDUCES THE CONTRACTION OF COLLAGEN GEL LATTICE
INCORPORATED WITH KELOID FIBROBLASTS. ......................................................................................132
FIGURE 44: GLEEVEC SIGNIFICANTLY REDUCES THE INTRACELLULAR ATP AND THE RATE OF ATP SYNTHESIS
IN KELOID FIBROBLASTS. ....................................................................................................................133
FIGURE 45: SUMMARY OF VARIOUS GROWTH FACTORS INVOLVED IN KELOID PATHOGENESIS AND THE
POSSIBLE THERAPEUTIC TARGETS. ......................................................................................................139
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ABBREVIATIONS
ALK
Activin receptor – like kinase
BMP
Bone morphogenetic protein
CTGF Connective tissue growth factor
DMEM
Dulbecco’s modified Eagle medium
ECM
Extracellular matrix
EGF
Epidermal growth factor
ELISA
Enzyme-Linked Immunosorbent Assay
ERK
Extraregulated Kinase
FCS
Fetal calf serum
FGF-2
Basic fibroblast growth factor
FPCL
Fibroblast populated collagen lattice
FPD Fibroproliferative disorder
GADPH
glyceraldehyde-3-phosphate dehydrogenase
HBSS
Hank’s balanced salt solution
HDGF Hepatocyte derived growth factor
HSPG
Heparan Sulfate Proteoglycan
IgG
Immunoglobulin G
IgM
Immunoglobulin M
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JNK
Jun-Amino terminal-Kinase
KF
Keloid fibroblasts
KGM
Keratinocyte growth medium
KK
Keloid keratinocytes
MAPK
Mitogen-Activated-Protein-Kinase
mTOR Mammalian target of rapamycin
MTT
3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide
NF
Normal fibroblasts
NK
Normal keratinocytes
PBS
Phosphate buffered saline
SCF Stem cell factor
TGF-β1
Transforming growth factor-beta1
VEGF
Vascular endothelial growth factor
α-SMA
Alpha-Smooth Muscle Actin
1
SUMMARY
Keloid scars are a result of such an aberration in wound healing, and are characterized by
overabundant collagen/ECM deposition. Several treatment modalities have been explored
but none of the treatments are effective. In spite of the large number of growth factors,
cytokines and chemokines, that have been studied and are now known to play a role in its
pathogenesis, the precise pathobiology leading to this scar formation remains largely
unknown. Thus to identify new targets for therapeutic intervention, it is imperative to
identify key modulators which are aberrantly expressed in keloids and to understand and
assess the role of these modulators towards this fibrotic phenotype.
This study aims to identify key modulators of cellular dynamics in keloid scars and
investigate the effect of epithelial-mesenchymal interactions on the expression profile of
these modulators using various in vitro models. This study further explores different
growth factors or chemical drugs as potential therapeutic agents for keloid scars.
Bearing in mind the importance of wound contraction and scar contracture in the wound
healing process and scar formation respectively, an in vitro coculture model and a
fibroblast-populated collagen lattice was employed to study the role of epithelial-
mesenchymal interaction towards a contractile phenotype. It was observed that
epithelial-mesenchymal interactions increased the contractile response of both normal
and Keloid fibroblasts in vitro. In addition KF’s were shown to have an elevated
contracile response to signals from the extracellular envirionment than normal
fibroblasts, due to the increased expression of α-SMA, a contractile protein.
2
Further studies were performed to investigate the molecular effectors of keloid scars and
the effect of epithelial-mesenchymal interaction on the expression profile of these
effectors.
Activin-A, a dimeric protein and a member of the TGF-β superfamily, has been shown to
regulate various aspects of cell growth and differentiation in the repair of the skin
mesenchyme and the epidermis. Thus the study aimed at investigating the role of activin
and its antagonist, follistatin, in keloid pathogenesis. The findings strongly suggest that
activin-A is a potent inducer of fibroblast activation and involved in the pathogenesis of
keloids. They also emphasize the importance of follistatin in regulating activin-A
bioactivity and suggest a possible therapeutic potential of follistatin in the treatment and
prevention of keloids.
Most of the growth factors and cytokines involved in the wound healing process seem to
be immobilized at the cell surface and extracellular matrix via binding with
proteoglycans, making them important modulators of cell dynamics. Thus the expression
of two proteoglycans, namely syndecan-2 and decorin, were investigated in order to
elucidate their roles in keloid pathogenesis.
It was demonstrated that syndecan-2 and FGF-2 were not only overexpressed in keloid
tissues but could also interact with each other resulting in the shedding of syndecan-2
which could in turn activate a whole cascade of events resulting in keloidogenesis. In
addition, decorin appeared to be downregulated in keloid tissues, and this protein with
strong antifibrotic effects could have the potential to be used as a therapeutic agent for
keloids.
3
Finally the ubiquity of SCF and its tyrosine kinase receptor c-KIT in different kinds of
cells and tissues, and their importance in wound healing and cancer led to their
investigation as potential players in keloid pathogenesis. The study demonstrated that
both SCF and c-KIT were upregulated in keloid scar tissues in vivo. They were also
upregulated in cultured fibroblasts on stimulation with serum highlighting their possible
role in the initial phases of the healing process when the wound is flooded with serum. In
addition, we demonstrated that epithelial-mesenchymal interactions, mimicked by
coculture of keratinocytes and fibroblasts in vitro, not only stimulated secretion of
soluble form of SCF in keloid cocultures but also brought about shedding of the
extracellular domain of c-KIT perhaps by upregulation of TACE which was also shown
to be elevated in keloid tissues and keloid cocultures. Although the increased
phosphorylation of c-KIT suggests the activation of SCF/c-KIT pathway in keloid
cocultures, the exact role of the shed c-KIT is yet to be understood.
Having identified a tyrosine kinase receptor system as a major player in keloid
pathogenesis, we investigated the role of an established tyrosine kinase inhibitor, gleevec,
as a possible therapeutic agent for keloid scars. It was observed that although gleevec did
not have a significant effect on the proliferative potential of KF and KK, it was as able to
downregulate the expression of ECM components as well as profibrotic factors in KF via
inhibition of the c-KIT mediated PI3 kinase signaling pathway. It was also able to render
the KF less bioenergetic by reducing the intracellular ATP levels and their rate of ATP
synthesis. Thus given its effects, it has a potential of being a therapeutic agent for keloids.
4
1 BACKGROUND AND INTRODUCTION
5
1.1 OVERVIEW OF THE WOUND HEALING PROCESS The wound healing process is a highly orchestrated process comprising of overlapping
phases of inflammation, granulation tissue formation, angiogenesis and remodeling of the
wound matrix (Clark et al., 1996). After wounding, the healing cascade immediately
ensues with the release of various growth factors and cytokines from the serum of the
disrupted blood vessels and degranulating platelets. The disruption of the blood vessels
results in the formation of a fibrin clot which not only works as a barrier for invading
microorganisms but also as a mesh for the binding of inflammatory cells, fibroblasts and
growth factors (Tuan et al., 1998). Within a few hours of injury, inflammatory cells like
neutrophils followed by monocytes and lymphocytes infiltrate the tissue and phagocytose
the bacteria. In addition to their role in defence against microorganisms, inflammatory
cells also serve as a pool for several growth factors and other macromolecules which are
required to intiate the proliferative phase of the healing process. The proliferative phase
involves the migration and proliferation of the keratinocytes from the wound edge
followed by the proliferation of fibroblasts in the area neighbouring the wound. These
fibroblasts then migrate into the wound area over a provisional wound matrix, deposit
large amounts of ECM and subsequently transform into myofibroblasts with a contractile
phenotype (Werner et al., 2003). Myofibroblasts are a group of actin rich fibroblasts and
are known to play an important role in wound contraction. Proto-myofibroblasts
differentiate to myofibroblasts by the production of α-SMA in order to generate more
force for contracture (Tomasek et al., 2002; Roy et al., 2001). At the same time,
angiogenic factors are released which promote angiogenesis by stimulating endothelial
cell proliferation leading to capillary tube formation. The resulting wound connective
6
tissue is called granulation tissue with nerve sprouting occuring at the wound edge.
Finally, a transition from granulation tissue to mature scar occurs, characterized by
continued collagen synthesis and collagen catabolism. This process requires a balance
between matrix biosynthesis and matrix degradation. A disruption in this balance either
due to excessive matrix deposition or decreased matrix degradation leads to keloid and
hypertrophic scars (Nedelec et al., 1996; Raghow et al., 1994).
Figure 1: Schematic representation of different stages of wound repair (Adapted from Werner et al., 2003). A: 12–24 h after injury the wounded area is filled with a blood clot. Neutrophils invade into the clot. B: at days 3–7 after injury, neutrophils undergo apoptosis. Instead, macrophages are abundant in the wound tissue at this stage of repair. Endothelial cells migrate into the clot; they proliferate and form new blood vessels. Fibroblasts migrate into the wound tissue, where they proliferate and deposit extracellular matrix to form a granulation tissue. Keratinocytes proliferate at the wound edge and migrate down the injured dermis and above the provisional matrix. C: 1–2 wk after injury the wound is completely filled with granulation tissue. Fibroblasts have transformed into myofibroblasts, leading to wound contraction and collagen deposition. The wound is completely covered with a neoepidermis.
12–24 h
Day 3-7
1-2 Week
7
1.2 KELOID SCAR
1.2.1 Clinical Characteristics of Keloid
Keloid scars represent a pathological response to cutaneous injury and occur only in
humans. The term ‘keloid’ is derived from a Greek word “khele” meaning crab claw
(Ladin et al., 1995). Keloids usually appear as firm broad nodules, often erythematous
and with a shiny surface (Marneros et al., 2004). A keloid scar extends beyond the
confines of the original wound, does not regress spontaneously, grows in pseudotumor
fashion with distortion of the lesion and tends to recur after excision. They are
notoriously resistant to therapy. Numerous treatment modalities are available, none of
which are consistently effective (Poochareon et al., 2003).
Fibroproliferative disorders (FPD) involve various pathologic fibrotic conditions which
constitute a leading cause of mortality in the United States (from a 5 year running
average of the US bureau of vital statistics analysis of cause of death, 45% of deaths in
1988 included death due to any form of fibrosis of any organ or tissue) (Kozak et al.,
1988; Bitterman et al., 1991).
The keloid is a dermal form of FPD and although it has much less morbidity as compared
to FPD of other organs, it causes considerable functional and aesthetic problems
particularly after thermal injury (Tredget et al., 1990; Engrav et al., 1987). They have a
propensity to occur in melanocyte-rich regions like face, neck, deltoid, presternal area
and ear lobes (Kim et al., 2000; Crockett et al., 1964; Bayat et al., 2004). They have also
8
been reported to appear on the cornea (Shukla et al., 1975; Shoukrey et al., 1993; Lahav
et al., 1982).
Keloids are often compared with hypertrophic scars. Though their gross appearance is
similar, hypertrophic scars unlike keloid scars, remain confined to the borders of the
original wound and most of the times retain their shape (Mutalik et al., 2005) [21]. In
addition, though hypertrophic scars develop within a few weeks after skin injury, keloid
scars often show a delayed onset.
Figure 2: Keloid formation in different patients (Adapted from Marneros et al., 2004)
9
1.2.2 Epidemiology
There is believed to be a genetic and racial predisposition for the development of keloids
in darker-skinned races compared to lighter skinned people. Some 15-20% of blacks,
hispanics, and asians are afflicted with the disorder. Studies have reported incidence
ratios between the two groups to range from 2:1 to 19:1 (Oluwasnami et al., 1974; Omo-
Dare et al., 1975). Both autosomal dominant and autosomal recessive genetic inheritance
have been proposed but not confirmed (Omo-Dare et al., 1975). Although keloids are
known to occur sporadically, some data suggest familial occurrence (Bloom et al., 1956).
Differences in occurrence of keloids based on age and gender has also been reported. It
has been shown to be predominant in females and in populations between the ages of 10
and 30 (Cosman et al., 1961; Murray et al., 1981). Until now, no susceptibility genes for
keloid formation have been identified. TGF-β1, β2, β3, and TGF-β receptor
polymorphisms have bene studies but has not yielded any statistically significant
associations with keloids in case control studies (Bayat et al., 2002, 2003, 2004,2005 ).
The Difficulty in identifying genes specific to keloids among patients with keloids may
reflect genetic heterogeneity, whereby different genes contribute to keloid formation in
different families (Robies et al., 2007)
1.2.3 Histopathology
Contrary to the appearance of organized fibres in normal skin, keloids have stretched
collagen fibres aligned in the epidermal plane (Kelly et al., 1988). Excessive deposition
of collagen and other extracellular matrix components along with increased blood vessels
10
and cells are observed in the histological sections of keloids. However the microvessels
in keloids seem to be occluded perhaps to the increase proliferation of endothelial cells
(Kischer et al., 1992). Keloid scars are also characterized by presence of tongue-like
advancing edge underneath normal-appearing epidermis and papillary dermis (Lee et al.,
2004). Keloid sections are also characterized by a thickened epidermis and prominent
disarray of fibrous fascicles/nodules. Mast cells, macrophages, epidermal langerhan cells
have all been demonstrated to be elevated in keloids. However their role in keloid
pathogenesis is not well understood. At higher magnification using scanning electron
microscopy, the random nature of fiber orientation, varying fiber length and poor bundle
formation in keloids are observed.
Figure 3: H&E staining of normal skin and keloid scar tissues
Normal Skin Keloid Scar
11
1.2.4 Molecular Pathogenesis
The wound healing process involves a complex interplay of cells, mediators, growth
factors, and cytokines, leading to inflammation, cell proliferation, ECM deposition,
contraction and remodeling. Thus a disruption in the cellular harmony leads to either a
delayed healing response or execessive healing. The over healing response is marked by
an over exuberant deposition of collagen and other extracellular matrix components like
fibronectin by fibroblasts resulting in a scar which can either be keloidic or hypertophic.
Previous studies have demonstrated that keloid-derived fibroblasts produced 20 times
more type1 collagen than normal fibroblasts in vitro (Ladin et al., 1995; Ala-kokko et al.,
1987; Di cesare et al., 1990).
Keloids are characterized by an increased proliferation of fibroblasts in vitro (Calderon et
al., 1996). Recent studies have highlighted the importance of regulation of apoptosis, in
scar establishment and the development of a pathological scar. Apoptosis-related genes
and proteins have been shown to be down-regulated or mutated in keloid tissues and
fibroblasts (Sayah et al., 1999; Chodon et al., 2000).
The bioactivity of fibroblasts in fibrogenesis or excessive scar formation is regulated by a
number of growth factors, of which the TGF-β family is thought to play a central role by
its regulation of fibroblast proliferation, differentiation and matrix production. It is
believed to enhance the production of ECM elements, such as fibronectin and collagen,
and to upregulate the cellular expression of matrix receptor integrin (Tuan et al., 1998;
Niessen et al., 1999; Kim et al., 2000).
12
TGF-β1 in particular has been widely implicated as a key modulator of cellular dynamics
in fibrosis. Many groups have demonstrated increased levels of TGF–β in keloid tissues
and other fibrotic disorders (Lee et al., 1999; Younai et al., 1994; Polo et al., 1999).
Peltonen and co workers (1991) demonstrated that TGF-β1 mRNA and TGF-β1 protein
were associated with excessive collagen synthesis and ECM accumulation in keloids.
Shah’s group (1994) showed that blocking of TGF–β1 and TGF–β2 by the application of
antibodies to cutaneous wounds markedly reduced scarring. Liu and team (2002)
demonstrated, by an in vitro experiment, that the effects of TGF-β1 might be modulated
by an autocrine loop and that this autocrine regulation might play an essential role in
keloid formation and development. Further, Chodon’s group (2000) reported that
dysregulation in the Fas-mediated apoptosis, which occurred in normal wound healing,
may be a factor contributing to scar formation and suggested a role for TGF-β1 in this
resistance.
Various other molecular pathways have been invoked in keloid pathogenesis. Platelet-
derived growth factor (PDGF) α-receptor expression was demonstrated to be elevated in
KF and this corresponded to an increase in mitotic response of the fibroblasts on
exposure to the PDGF isoforms (Haisa et al., 1994). An over expression of insulin-like
growth factor-I (IGF-1) receptor in KF was also demonstrated to enhance their invasive
potential highlighting the role of IGF-1 in keloid pathogenesis (Yoshimoto et al., 1999).
An elevated expression of interleukin (IL)-6 (Xue et al., 2000) and VEGF (Wu et al.,
2004; Le et al., 2004) has also been reported in KF.
13
As the wound milieu consists of various cell types, it would be foolhardy to implicate
fibroblasts alone as main players in fibrosis. Other cell types have also been shown to
play a role in keloid pathogenesis directly or indirectly by influencing the fibroblasts. An
elevated presence of mast cells was observed in keloids suggesting their possible role in
keloid pathogenesis (Russel et al., 1977). Upon further investigation it was observed that
an elevation of mast cell histamine resulted in an up-regulation of procollagen type 1
production in KF (Kikuchi et al., 1995; Tredget et al., 1998). Another cell type which has
been shown to play an important role in pathogenesis of scars is endothelial cells. Along
with fibroblasts they are known to be important in production of collagen (Milsom et al.,
1973; Sollberg et al., 1991; Diegelmann et al., 1977). It has been observed that
overproliferation of endothelial cells leads to occlusion of microvessel lumens in keloids
resulting in a severe hypoxic condition (Kischer et al., 1982). The resulting hypoxia
further stimulates endothelial cell proliferation and collagen production by fibroblasts by
releasing growth factors like HIF-1α and VEGF (Knighton et al., 1983; Kischer et al.,
1992; Zhang et al., 2006).
It is now becoming clear, that epithelial-mesenchymal interactions, initially applied to
normal skin homeostasis (Maas-Szabowski et al., 2000; Mackenzie et al., 1994; Fusenig
et al., 1994), are also instrumental in modulating fibroblast behaviour in keloids by
paracrine influences. Previous laboratory-based findings showed that keloid-derived
keratinocytes enhanced both normal and keloid-derived fibroblast proliferation and
collagen synthesis in a co-culture system (Lim et al., 2001; Funayama et al., 2003; Xia et
al., 2004). NF and KF cocultured with KK in defined serum-free media showed a higher
rate of proliferation compared to that of coculture with NK (Lim et al., 2001; Funayama
14
et al., 2003; Xia et al., 2004). It was subsequently observed that KF co-cultured with KK
produced increased amounts of both collagens I and collagen III compared with coculture
with NK (Xia et al., 2004; Lim et al., 2002; Lim et al., 2003). Mitogen activation protein
kinase (MAPK) and phosphatidyl-inositol-3 kinase (PI-3K) pathway activation have been
observed in excessively proliferating KF cocultured with KK. When these fibroblasts
were treated with MAPK-p44/42 specific inhibitor and PI-3K specific inhibitor, it caused
complete nullification of collagen I and III production, and significantly decreased
fibronectin and laminin β2, all of which is produced in excess in keloidogenesis (Lim et
al., 2003). Electron microscopic analysis also demonstrated that the appearance of
collagen–extracellular matrix (ECM) produced by NF cocultured with KK in vitro,
approximated the morphological appearance of in vivo keloid scar tissue (Lim et al.,
2002). Epithelial-mesenchymal interactions have been demonstrated to influence the
expression of various transcription factors, growth factors and cytokines in vitro. Phan
and his group (2005) demonstrated that epidermal-dermal interactions in keloids resulted
in activation of the TGFβ-Smad axis which in turn played a crucial profibrotic role in
keloid pathogenesis. Lim and his co-workers (2006) further demonstrated the modulation
of yet another transcription factor STAT-3 by epithelial-mesenhcymal interactions in
keloidic cells. Ong’s team (2007) used a two chambered coculture system to mimic
epithelial-mesenchymal interaction and reported the expression and secretion of VEGF to
be upregulated in KF on being coculutred with KK. Biologically active CTGF was also
shown to be secreted by KF only in the presence of overlying keratinocytes (Khoo et al.,
2006).
15
In addition to the increase of profibrotic molecules, anti fibrotic growth factors and
enzymes have also been demonstrated to be downregulated in keloids. The balance
between matrix metalloproteinase (MMP) and their inhibitors TIMP is critical for over all
ECM turnover. Concentrations of collagenase inhibitors, α-globulins and plasminogen
activator inhibitor -1 have been shown to be elevated in both in vitro and in vivo keloid
samples, whereas the levels of degradative enzymes are frequently decreased
(Diegelmann et al., 1977; Tuan et al., 1996). It has also been observed that MMP activity
differs between KF and NF and these differences affect the phenotype (Uchida et al.,
2003).
Although attempts are being made in understanding the molecular basis of keloid
pathogenesis, the complexity of the repair process and the lack of proper in vitro and in
vivo animal models have hindered progress in revealing the mechanisms underlying
keloid scar formation. Thus it is imperative that new in vitro and in vivo methods are
developed to assists researchers to delineate the dynamics of growth factors, deemed
important in normal wound healing process, in pathological scars.
16
Figure 4: Schematic representation of pathways potentially resulting in accumulation of collagen in fibrotic skin diseases (Adapted from Uitto et al., 2007). The net deposition of collagen is a balance between the rate of synthesis and the rate of degradation, both of which can be modulated by a number of factors, such as cytokines. These factors can be evoked by stimuli such as trauma to the skin. When superimposed on the individual’s genetic background, an imbalance in the flux through these pathways can result in collagen accumulation manifesting as tissue fibrosis.
1.2.5 Current Treatment
Several modalities have been explored for either the treatment of keloid scars or
prevention of their recurrence after surgery. However the several treatment strategies
used, remain unsatisfactory. The current treatments available are as follows:
Surgery: Surgical excision of hypertrophic and keloid scars when combined with steroid
injection reduces the recurrence rate. However, excision alone in keloids is met with high
17
recurrence rate of 45 to 100% (Berman et al., 1995; 1996). Due to this, various adjunct
therapies have been explored to reduce the problem of recurrence.
Silicone Gel Sheeting: Silicone gel is a crosslinked polymer of dimethylsoloxane and is
not only an effective adjunct to excision of keloids but also a prophylaxis to abnormal
scarring in some incisions (Al-Attar et al., 2006). Despite the wide appreciation about the
effect of silicone gel sheeting among the physicians, the mode of action of silicone is still
unknown. Although silicone gel is comfortable, it requires active patient compliance and
long-term application which is challenging on mobile and angled anatomical sites (Sproat
et al., 1992).
Corticosteroid Injection: Injection of triamcinolone is efficacious for the treatment of
keloid scars in which it is used as a first-line therapy (Shons et al., 1983; Tang et al.,
1992). Corticosteroids are usually combined with surgical excision of the scar (Berman et
al., 1996). However intralesional injections of corticosteroids are highly painful. In
addition other side effects are skin atrophy, loss of pigmentation, and telangiectasia
(Sproat et al., 1992).
Pressure Therapy: Pressure therapy is an effective therapy with minimal side effects for
keloids but its practical use is limited to ear lobes. It is generally used as a post operative
adjunct for earlobe keloids (Brent et al., 1978; Russell et al., 2001).
Radiotherapy: Radiotherapy has been used as monotherapy, and in combination with
surgery, for hypertrophic scars and keloids. It is effective in reducing the recurrence rate
of scars. However, monotherapy remains controversial because of anecdotal reports of
carcinogenicity following the procedure (Thomas et al., 2002; Norris et al., 1995).
18
Cryotherapy: Cryotherapy is an option considered in treatment of very small scars like
those arising from very severe acne (Layton et al., 1994). It uses rapid, repeated cooling
and rewarming of tissue causing cell death and tissue sloughing (Al-Attar et al., 2006).
Side-effects produced are slow healing of the wound along with tissue necrosis, altered
pigmentation, moderate skin atrophy and pain. The response rate is around 51 to 74
percent and may require more than one application (Thomas et al., 2002).
New emerging therapies: Recently, a number of novel experimental therapeutic
approaches have been explored, based on results from in vitro experiments from cells
derived from keloid tissue [eg. Interferon γ (Granstein et al., 1990; Larrabee et al., 1990),
imiquimod (Berman et al., 2002), 5-fluorouracil (Manuskiatti et al., 2002 ; Fitzpatrick et
al., 1999), bleomycin (Yamamoto et al., 2006).
Although new treatment modalities are constantly been explored, most of them are
accepted on a broad consensus rather than backed by a good knowledge of the
pathogenesis of keloid and hypertrophic scars. Adding to the problem, there is an evident
shortage of randomized controlled trials to assess the efficacies of existing therapeutic
interventions. It is also not possible to incorporate variables which widely exist among
the patients in actual clinical setting, like differences in age, race, involved body part,
associated systemic diseases like diabetes, local infection, in a volunteer-based study
using artificial methods of induction of wound. Results based on animal/animal cell
studies may not be reliable because animal tissue/cell behaviors are much different from
that of humans and more over, diseases like keloid exclusively occur in humans. These
19
limitations make the assessment and comparison of different treatment methods very
difficult.
Thus, for more effective and specific treatments, it is imperative that molecular
mechanisms that cause keloid formation are elucidated and better understood. Increased
understanding at each level of pathogenesis may lead to development of new therapies, in
form of synthetic drugs or specific growth factors, to downregulate profibrotic molecules.
Monoclonal antibodies, growth factor receptor antagonists, antisense oligonucleotides are
some of the treatment modalities that hold promise.
1.3 OBJECTIVES OF THE PRESENT STUDY We hypothesise that epithelial–mesenchymal interactions have an important governing
role in keloid formation, and that keloid formation may be the consequence of abnormal
keratinocyte control over fibroblasts, mediated by the key factors like activin-A,
proteoglycans, Stem cell factor , rather than a defect of fibroblasts themselves.
The broad objective of this study is to identify key modulators of cellular dynamics in
keloid scars and investigate the effect of epithelial-mesenchymal interactions on the
expression profiles of these modulators using various in vitro models. Depending on the
profibrotic or antifibrotic effects of these modulators, they could either be targets for
therapeutic intervention using chemical agents or themselves be used as natural
therapeutic agents for keloid scars, respectively.
20
The specific objectives of this study are as follows:
1. To investigate the role of epithelial-mesenchymal interactions, especially
fibroblast response to keratinocyte paracrine stimulation in wound contracture.
2. To investigate the role of various growth factor systems in keloid pathogenesis
a) Activin-A/ follistatin system
b) Proteoglycans
c) SCF/c-Kit system
3. Finally to investigate both growth factors and synthetic agents which could be
used as therapeutic agents for keloids
a) Natural modulators: decorin, follistatin
b) Synthetic modulators: gleevec (Imatinib mesylate)
21
2 MATERIALS AND METHODS
22
2.1 MEDIA AND CHEMICALS
Dulbecco’s modified eagle medium (DMEM), Hank’s balanced salt solution (HBSS),
fetal calf serum (FCS), streptomycin, penicillin, gentamicin and fungizone were
purchased from Gibco. Keratinocyte growth medium (KGM) was purchased from
Clonetics (USA). Phosphate buffered saline without Ca2+ and Mg2+ (PBS), epidermal
growth factor (EGF), cholera toxin and hydrocortisone were purchased from Sigma
Chemical Co (USA). Dispase II was purchased from Boehringer Mannheim (USA).
Rhodamine counter stain was obtained from Difco (USA). Tris base was purchased from
J.T Baker. Triton X-100, ethylenediaminetetraacetic acid (EDTA), 30% acrylamide/bis
solution (37.5:1 2.6%C) and glycine were purchased from Biorad. Sodium Chloride
(NaCl), nonidet P-40 (NP-40), sodium dodecyl sulphate, hydrogen peroxide (H2O2),
bovine serum albumin (BSA), tween-20, potassium chloride (KCl), potassium phosphate
(K3PO4), magnesium chloride (MgCl2), MTT [3-(4,5-Dimethylthiazol-2-yl)-2,5-
diphenyltetrazolium bromide], N,N –dimethylformamide (DMF) and paraformaldehyde
were all purchased from Sigma Chemical Co (USA). Methanol and acetic acid were
purchased from Lab-Scan.
2.2 RECOMBINANT GROWTH FACTORS AND ANTIBODIES
Mouse anti-α- SMA monoclonal antibody (Sigma), mouse anti-TGF- β1 neutralizing
antibody (R&D systems), Mouse anti-Collagen I,III monoclonal antibody
(Monosan®antibodies-The Netherlands), rabbit anti-fibronectin monoclonal antibody
(BD Transduction Laboratories), mouse anti-activin-A monoclonal antibody (Oxford
23
Bio-Innovations, Oxford, UK) and mouse anti-follistatin (Gift from Dr David Phillips,
Monash University, Melbourne, Australia), rabbit-anti syndecan-2 polyclonal antibody
(Santa Cruz Biotechnology), rabbit-anti FGF-2 polyclonal antibody (Santa Cruz
Biotechnology), mouse anti-decorin monoclonal antibody (R&D systems), rabbit-anti
TACE polyclonal antibody (Santa Cruz Biotechnology), rabbit anti-c-Kit (tyr 703)
polyclonal antibody (Affinity BioReagents), rabbit-anti c-Kit (tyr 721) polyclonal
antibody (Affinity BioReagents), mouse-anti c-Kit monoclonal antibody (Neomarkers),
mouse-anti c-Kit monoclonal antibody (R&D systems), rabbit-anti VEGF polyclonal
antibody (Neomarkers), HDGF (Gift from M.D Anderson Cancer Center, U.S) , mouse-
anti Phospho Akt (Cell signaling), mouse-anti Phospho mTOR (Cell signaling), Labeled
chicken- anti mouse IgG antibody (Molecular probes), goat-anti rabbit IgG H&L (HRP)
polyclonal antibody (abcam), rabbit-anti mouse IgG H&L (HRP) polyclonal antibody
(abcam), rabbit-anti mouse IgM H&L (HRP) polyclonal antibody (abcam), recombinant
human activin-A (R&D systems), recombinant human follistatin (R&D systems),
recombinant human decorin (R&D systems), recombinant human TGF-β1 (R&D
systems), recombinant human FGF-2 (R&D systems).
2.3 PREPARATION OF NORMAL AND KELOID TISSUE EXTRACTS
Normal and keloid skin tissues were cut into small pieces using a scalpel to about 120
mg. 200 µl of lysis buffer containing 20 mM Tris-HCl (pH 7.5), 1% v/v Triton X-100,
100 mM NaCl, 0.5% w/v Nonidet P-40 and 1mg/ml protease inhibitor cocktail
(Boehringer Mannheim, Mannheim, Germany) were added to the tissue sections and
24
sonicated. This was followed by centrifugation at 13000 x g for 10 min. Supernatant was
collected while the pellet was discarded. Protein concentration of the tissue extracts were
determined by the Bradford method.
2.4 IMMUNOHISTOCHEMISTRY
2.4.1 Preparation of Paraffin Sections of Normal and Keloid Tissues
Normal and keloid tissue samples were fixed in 4% v/v formalin in PBS for 24 hr (longer
for larger samples but never more than a week) followed by immersion in 70% v/v
ethanol overnight, in histocassettes. This was followed by paraffin embedding as per
standard protocol (BD biosciences). Sections were cut at 5 microns onto slides.
Tissue sections were dried onto the slides at 37 oC overnight.
2.4.2 Pretreatment of Paraffin Sections for Immunohistochemistry
The paraffin sections were dewaxed or deparaffinized in two changes of xylene followed
by re-hydration at 100%, 95% and 70% v/v ethanol gradients. The antigens were then
retrieved by immersing slides in 0.01 M citrate buffer, pH 6.0, heating in a microwave
oven (high for 2.5 min, low for 5 min), cooling at 4°C for 20 min, and washing in water
for 5 min. Endogenous peroxidase was blocked in 3% v/v H2O2, and non-specific binding
blocked for 1 hr (CAS block, Zymed Laboratories, South San Francisco, CA, USA).
25
2.4.3 Probing with Antibody and Developing
Sections were then incubated in specific antibodies for 1-2 hrs at room temperature. After
washing, slides were incubated in universal secondary antibody provided in the vectastain
kit (Vector Labs). Slides were washed in Tris-buffered NaCl (TBS) 0.05% w/v Tween-20
pH 7.5, then MilliQ H2O. The sections were then treated with “vectastain elite RTU elite
reagent” for five minutes. The reaction product was developed with a 3, 3’-
diaminobenzidine tetrahydrochloride substrate kit (Dako), and sections counterstained
with hematoxylin. All wash steps were in TBS/0.05% w/v Tween-20. Antibodies were
diluted in 1% w/v BSA/TBS.
2.5 CELL CULTURE
2.5.1 Keloid Keratinocyte and Fibroblast Database
Keratinocytes and fibroblasts were randomly selected from a specimen bank of
keratinocyte/fibroblast strains derived from excised keloid specimens. All patients had
received no previous treatment for keloids before enrollment. Before informed consent
was obtained, a full history was taken and an examination performed, complete with
photographic documentation. The tisssues were separated into normal and keloids by an
approved surgeon and pathologist. Approval by the National University of Singapore-
Institutional Review Board (NUS-IRB) was obtained for this study.
26
2.5.2 Keloid and Normal Keratinocytes from Keloid scar and Normal Skin
Excised keloid scar and normal skin specimens were repetitively washed in PBS
containing 150 µg/ml gentamicin and 7.5 µg fungizone, until the washing solution
became clear. The tissue was then divided into pieces of approximately 5 mm × 10 mm
and the epidermis was scored. Dispase 5mg/ml in HBSS was added and skin was
incubated overnight at 4ºC. The epidermis was carefully scraped off with a scalpel the
next day and placed in trypsin 0.25% w/v /glucose 0.1% w/v /EDTA 0.02% for 10 min in
the incubator. Trypsin action was quenched by DMEM/10% FCS. The suspended cells
were transferred into tubes and centrifuged at 1000 rpm for 8 min. The cells were seeded
in Keratinocyte Culture Medium (80 ml DMEM supplemented with 20 ml FCS, EGF 10
ng/ml, cholera toxin 1 × 10-9 M and hydrocortisone 0.4 µg/ml) at 1 × 105 cells/cm2 for 24
hrs before changing to Keratinocyte Growth Medium (KGM). The cell strains were
maintained and stored at -150ºC. Only cells from second and third passages were used for
the experiments.
2.5.3 Keloid and Normal Fibroblast from Keloid Scar and Normal Skin
Remnant dermis from the keloid scar and normal skin were either minced or incubated in
a solution of collagenase type 1 (0.5 mg/ml) and trypsin (0.2 mg/ml) at 37ºC for 6 hrs.
Cells were pelleted and grown in tissue culture flasks. Alternatively the skin tissue
samples were chopped into pieces of 1-2 mm2. The pieces were then transferred to a 100
mm tissue culture dish previously coated with a thin layer of DMEM/10% v/v FCS.
Culture medium enough to cover the explants were then added and topped up after 2-3
27
days. After 4-7 days the fibroblasts outgrew from the tissue. Fibroblast cell strains were
maintained and stored at -150ºC until use. Only cells from the second and third passages
were used for the experiments.
2.5.4 Keratinocyte-fibroblast Coculture KK obtained from randomly selected keloid strains were seeded at density of 1 × 105
cells/cm2 on 6-well transwell clear polyester membrane inserts with 0.4 µm pore size and
area 0.3 cm2 area (Costar Corp, USA). Ten days before co-culture, cells were maintained
in serum-free Keratinocyte Growth Medium (KGM) until 100% confluent in monolayer.
The medium was then changed to Dulbecco’s modified Eagle’s medium (DMEM)
supplemented with 10% heat inactivated fetal bovine serum (FBS) and
penicillin/streptomycin. The cells were raised to air-liquid interface to allow
keratinocytes to stratify and reach terminal differentiation.
KF obtained from randomly selected keloid strains were seeded in 6-well plates at a
density of 1 × 104 cells/ml in DMEM/10% v/v FCS for 24 hrs and then in serum-free
medium for another 48 hrs. Cells on both the membrane inserts and the wells were
washed twice with phosphate buffer saline (PBS) to remove the old medium before
combination of the inserts and plates for co-culture in serum-free DMEM. Controls
comprised of one series of non co-cultured keloid fibroblasts and one series non co-
cultured keloid keratinocytes. At day 5, the inserts with the cultured keratinocytes were
removed, and the conditioned media were collected, pooled and stored at -80ºC for later
28
analysis. The fibroblasts and keratinocytes were also harvested for protein extraction for
western blot analysis. (Fig. 5)
Figure 5: Coculture of epidermal keratinocytes and dermal fibroblasts as an in vitro model to study epithelial-mesenchymal interaction
FFiibbrroobbllaasstt ssuubbccuullttuurree KKeerraattiinnooccyyttee ssuubbccuullttuurree
KKeerraattiinnooccyyttee-- FFiibbrroobbllaasstt CCooccuullttuurree
29
2.6 CELL COUNTING
Before the cells were seeded into culture flasks for experiments, aliquots of the cell
suspension were mixed with trypan blue in a ratio of (1:4) and counted in a Neubauer’s
haemocytometer. All the cells (non-viable cells stained blue, viable cells became opaque)
were counted in the four corner squares of the hemocytometer. Since the volume of each
square is 10-4 cm3 use the following formula was used to calculate the number of cells in
the cell suspension
Cells per ml = the average count per square x the dilution factor x 104
Total cell number = cells per ml x the original volume of fluid from which cell sample
was removed.
2.7 TREATMENT OF FIBROBLASTS WITH GROWTH FACTOR
NF and KF from different patients were seeded in 6-well plates at a density of 1 × 104
cells/ml in DMEM/10% v/v FCS for 24 hrs and then in serum-free medium for another
48 hrs. The cells were subsequently treated with varying concentrations of activin-A (100
ng/ml; (R&D Systems), follistatin (FS-288, 100 ng/ml; R&D Systems) or TGF-β1 (5, 10
ng/ml; R&D Systems), decorin (0, 250, 500, 1000, 2000 ng/ml; R&D Systems), FGF-2
(10 ng/ml; R&D Systems) respectively, to test the effect of these growth factors on the
proliferation of fibroblasts and expression of key ECM proteins.
2.8 TREATMENT OF CELLS WITH GLEEVEC
30
NF and KF from different patients were seeded in 6-well plates at a density of 1 × 104
cells/ml in DMEM/10% v/v FCS for 24 hrs and then in serum-free medium for another
48 hrs. The cells were subsequently treated with varying concentrations of gleevec (0.5
µg/ml, 1 µg/ml, 2.5 µg/ml, 5 µg/ml, 10 µg/ml, and 20 µg/ml), to test its effect on the
proliferation of fibroblasts and expression of key ECM proteins.
2.9 SERUM STIMULATION
When fibroblasts are exposed to serum, the signal is interpreted to be a physiological
wounding signal. The serum stimulation model is an in-vitro model used to mimic the
early phases of wounding. NF and KF were seeded in 6-well plates at a density of 1 × 104
cells/ml in 10% v/v FCS for 24 hrs and subsequently seeded in serum-free medium for
another 48 hrs. The fibroblasts were exposed to either DMEM supplemented with 10%
v/v FCS or DMEM alone for 24 hrs before being harvested for analysis.
2.10 MTT ASSAY
The MTT assay is a commonly used colorimetric assay to quantify cell numbers. It is
widely used in studies involving cell proliferation or cell toxicity. MTT [3-(4,5-
Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium] is cleaved by an active succinate-
tetrazolium reductase system present in the mitochondrial respiratory chain of a living
and metabolizing cell into blue formazon crystals which can be solubilized and
absorbance measure. The relationship between cell number and absorbance is linear.
31
In our experimental set up, the treated cells in 96-well plates were incubated with 10 µl of
MTT (5 mg/ml) in 100 µl of DMEM to give a final concentration of 0.5 mg/ml in each
well for about 2hrs. The medium was removed and the blue crystals were soubilized by
Hansen’s method (1989) using 20% w/v SDS in a solution of DMF: water (1:1 v/v) and
shaking in an orbital shaker. The absorbance of the solution was then measured directly
by using a plate reader at 570 nm.
2.11 FIBROBLAST-POPULATED COLLAGEN LATTICE (FPCL)
2.11.1 Preparation of Fibroblast-Populated Collagen Lattices
Collagen lattices were prepared from type 1 collagen extracted from rat tail tendons in
0.1% (v/v) acetic acid. A working solution was constituted by diluting the collagen to 1.6
mg/ml on the day of experiment. Both NF and KF cell strains were trypsinised,
resuspended in medium and counted using a hemocytometer. A collagen-fibroblast (for
both NF and KF) suspension was prepared in serum-free DMEM and 2 ml of the
suspension was dispensed into 32-mm petri dishes. The final concentration of collagen
was 0.8 mg/ml with a cell density of 6.7 × 104 cells/ml. The collagen lattices were
subjected to different treatments and the area of the lattice calculated.
32
2.11.2 Macroscopic Evaluation of FPCL Contraction
The lattice diameters were assessed at various time-points for several days by placing the
dish on transparent graph paper and measuring two diameters at right angles to one
another. The area of collagen gel populated with fibroblasts was photographed. The
average area was calculated from the mean of the diameters and expressed as a
percentage of initial lattice area.
Percentage of initial area = A2/A1 × 100,
where A1 = initial gel area and A2 = area at the observed interval
2.12 WESTERN BLOTTING
To study the changes in the expression of investigative proteins in keratinocytes and
fibroblasts under different culture conditions and under treatment with different drugs,
cells were lysed in lysis buffer containing 20 mM Tris-HCL (pH 7.5), 1% v/v Triton X-
100, 100 mM NaCl, 0.5% w/v Nonidet P-40 and 1 mg/ml protease inhibitor cocktail
(Antipaindihydrochloride, Aprotinin, Bestatin, Chymostatin, E-64, EDTA-Na, Leupeptin,
Pefabloc-SC, Pefstatin, Phosphormidon; Boehringer Mannheim, Mannheim, Germany).
100 µg of protein was extracted and electrophoresed on 8% w/v and 14% w/v sodium
dodecyl sulphate-polyacrylamide gels using the Protein II system (Bio-Rad). The proteins
were subsequently transferred to a nitrocellulose membrane and blotted for antibodies
against the proteins under investigation. This was followed by treatment with respective
33
secondary antibodies. The blots were then visualized with a chemiluminescence-based
photoblot system (Amersham Biosciences).
2.13 IMMUNOASSAY
An immunoassay is a biochemical test generally used to measure the level of antigens
present in a biological fluid like serum or urine. It has an advantage of being highly
specific as well as sensitive. Due to this reason, often very low volume of sample is
required to detect the antigen.
In our study, ELISA kits for detection of investigative proteins were purchased and
assays performed on conditioned media (100 µl) obtained from NF, NK, KF and KK.
2.14 FLOW CYTOMETRY
FACS analysis was performed to detect the investigative protein’s expression. Briefly,
about 5*105 to 1*106cells were washed twice with ice-cold PBS at 250 g for 5 min. The
cells were then incubated with antibodies against the proteins for 30 min at 4°C followed
by incubation with an FITC-conjugated secondary antibody in for 30 minutes at 4°C in
the dark. Cells treated with an isotype control or without primary antibody were used as
negative controls. All washing steps involved washing with flow cytometry buffer
containing PBS, 2% v/v FBS and 0.1% v/v sodium azide. Thereafter, the cells were fixed
in 2% w/v paraformaldehyde in PBS for 15 minutes at 4°C in the dark. A FACScan
(Becton Dickinson, Sunnyvale, CA) flow cytometer was used for the analysis.
34
2.15 RNase PROTECTION ASSAY
The RNase protection assay is a highly sensitive method used to detect and quantify
specific mRNA transcripts in a mixture of total RNA or mRNA. The assay involves
hybridising a radioactive probe complementary to the gene sequence of interest, and then
adding RNase to the mixture. The RNase digests the entire single stranded RNA leaving
behind the double stranded probe-target hybrid. This hybrid is then electrophoresed on a
denaturing TBE-urea polyacylamide gel and detected.
In this study, the total RNA was isolated from the tissue lysate. RNase protection assay
was performed as described by Werner’s group (1993) using a radiolabelled antisense
probe complementary to the activin-βA mRNA (Hübner et al., 1996). Hybridization of
the same RNA with a gapdh riboprobe served as a loading control.
2.16 RATE OF ATP SYNTHESIS
The measurement of ATP by the luciferin-luciferase reaction was done as previously
described. (Vincent et al., 2004; Zhang et al., 2004) and was extended in this study to
digitonin-permeabilized whole fibroblasts. KF were permeabilized with 65 µg
digitonin/106 cells in buffer (150 mM KCl, 25 mM Tris-HCl, 2 mM EDTA, 0.1% w/v
BSA, 10 mM potassium phosphate, 0.1 mM MgCl2, pH 7.4). The reaction mixture
contained 1 mM ADP in a total volume of 0.25 ml of the above buffer. The reaction was
started with 50,000 cells, treated with the drug or only treated with DMEM with 5% v/v
serum for 24 hrs, followed by incubation at 37oC for 5 min and stopped by boiling for 3
min. Following centrifugation, 25 µl supernatant was added to a well in a 96-well plate
35
together with 100 µl of 250x-diluted FL-AAM (Sigma). The chemiluminescence
generated was read in a luminometer (PerkinElmer Victor) as described previously by
Ng’s group (2006). The amount of ATP biosynthesized in 5 min was extrapolated from a
standard curve of ATP of 5 to 100 ρmol, a range which produced a linear response.
2.17 MEASUREMENT OF INTRACELLULAR ATP Intracellular ATP was determined from the whole cell lysate following the luciferin-
luciferase assay described above. Normalization was done with the protein content of the
lysate determined by the Bradford method.
2.18 STATISTICAL ANALYSIS Statistical significance between groups was assessed using either Student’s t-test or a
one-way analysis of variance with Tukey’s post-hoc test. All tests were performed using
MiniTab® software. Except for ELISA measurements where the mean value of data
from 5 different cell populations in each group was used, for all other results the cell
population representative of other populations with at least 5 different clinical samples
studied was conducted. Differences at the 95% level ( P value < 0.05) were considered to
be significant.
36
3 ROLE OF EPITHELIAL-MESENCHYMAL INTERACTIONS IN WOUND CONTRACTION AND
SCAR CONTRACTURE
37
3.1 INTRODUCTION The process of wound contraction, which involves the centripetal movement of the
wound edges by means of a retractile activity of the granulation tissue, is very critical for
proper healing of any cutaneous wound. However, unlike in other mammals, the
contraction of wound is often less beneficial in humans leading to minimal scarring in
some cases to major body deformations (Hunt et al., 1979; Rudolph et al., 1992).
Although it is widely accepted that the contractile force resides in the granulation tissue
which fills the wound (Carell et al., 1916), the exact mechanism is still not clearly
understood.
Epithelial-mesenchymal interactions have recently been explored as a possible factor
contributing to excessive scar formation. Given their emerging role in keloid
pathogenesis ((Lim et al., 2001; 2002; 2003; Funayama et al., 2003; Xia et al., 2004)
they might well be an important player in modulating wound contraction and scar
contracture. During wound contraction, fibroblasts differentiate into myofibroblasts
expressing α-smooth muscle actin. In normal wound repair, these myofibroblasts undergo
apoptosis and disappear but under pathological conditions these myofibroblasts become
apoptosis resistant, remain in the extracellular matrix and continue to remodel, leading to
sustained wound contracture (Tomasek et al., 2002). Previous studies reported the
modulation of underlying mesenchymal cell apoptosis by the epithelium (Hurle et al.,
1986). It has been further reported that epithelial–mesenchymal interaction upregulates
TGF-β1 (Funayama et al., 2003; Xia et al., 2004), a growth factor with important roles in
38
wound and scar contraction (Kamamoto et al., 2003; Spyrou et al., 2002). In addition,
TGF-β has also been found to inhibit apoptosis of different types of cells including
myofibroblasts (Smith et al., 1999; Kim et al., 1998; Genestier et al., 1998) by various
mechanisms including the downregulation of c-myc, preventing the decline of Bcl-2
(Zhang et al., 1999) and through activation of the protein kinase pathway (Chin et al.,
1999). Taken a step further, epithelial-mesenchymal interactions would similarly affect
wound contracture by modulating apoptosis of myofibroblasts. The exact mechanism for
this however remains unclear.
New investigative tools, arising from tissue bioengineering have stimulated new studies
related to scar formation. Bell’s group (1979) described an in vitro model for the study of
wound contraction using collagen gels populated by fibroblasts. The advantage of this
model is that it includes the two fundamental dermal participants in scar formation: the
ECM and the fibroblasts, unlike the traditional in vitro models where the cells are grown
as a monolayer. Harris and co-workers (1981) demonstrated that the contraction of the
collagen lattice occurred as a consequence of fibroblast migration through the matrix.
This process was termed ‘tractional remodeling’ (Grinell et al., 1994).
In this study, we hypothesized the role of epithelial-mesenchymal interactions, especially
fibroblast response to keratinocyte paracrine stimulation in wound contracture. To test
our hypothesis, we investigated the contractile response of collagen lattices populated by
KF and NF, respectively to the addition of conditioned media obtained from KK/KF
coculture. To complement this data, immunohistochemical analysis of the fibroblasts
exposed to KK/KF conditioned media was performed to assess the expression of alpha-
smooth muscle actin in order to test our second hypothesis that fibroblasts exposed to this
39
abnormal media might differentiate into the myofibroblasts in keloid scar tissue. The
effect of anti-TGF β1 neutralizing antibody on collagen lattice contraction induced by
KK/KF coculture conditioned media was also studied to investigate the role of this
fibroinductive cytokine.
3.2 RESULTS • Keloid or normal dermal fibroblast populated collagen lattice exhibited
significant contraction when treated with KK/KF coculture conditioned media
Study groups comprised collagen matrices populated with KF or NF exposed to the
conditioned media collected from KK/KF coculture or non-cocultured KKs (KK48,
KK43, KK30, KK25) or KFs (KF48, KF43, KF30, KF25). Lattice areas were calculated
at intervals of 1, 2, 3, 6, 24 and 48 hrs to determine the contraction percentage.
Conditioned media from the non-cocultured KK and non-cocultured KF controls induced
slight contraction of collagen lattices populated by KFs (Fig. 6a), but no contraction in
the group of collagen lattices populated with normal dermal fibroblasts (Fig. 6b).
Conditioned media obtained from KK/KF coculture, however, induced significantly
greater collagen lattice contraction, with nearly 90% or 60% contraction of the collagen
gel at 48 hrs compared with approximately 20% or 5% contraction for the controls, with
KF or NF, respectively (Fig. 6a,b).
40
• Contraction of collagen lattice populated by KF was more significant as
compared to that populated by NF
The FPCLs were photographed and the area calculated as previously mentioned, for side-
by-side analysis. It was observed that collagen lattices populated with KFs demonstrated
significantly greater contraction when treated with KK/KF conditioned media compared
with collagen lattices populated with NF (Figs 6 and 7)
• Expression of α-smooth muscle actin was increased in keloid-derived fibroblasts
treated with keloid keratinocyte/keloid fibroblast coculture conditioned media
KF and NF were seeded at a density of 1000 cells per well in eight-well Laboratory-tek
chamber slides in DMEM/10% FCS for 24 hrs. The medium was then replaced by serum-
free DMEM for 48 hrs. After 48 hrs, conditioned media pooled from KK/KF coculture
and non-coculture samples were added. This was followed by addition of anti-α-SMA
antibody (Sigma, A-2547). A rhodamine counterstain (Difco) was added to the
fluorescein isothiocyanate-labelled secondary antibodies to reduce the nonspecific
background staining. A bright green fluorescence marked the positive expression of α-
SMA, while cells demonstrating minimal or no expression were stained dull orange.
Compared with control groups treated with non-coculture conditioned media, increased
expression of α-SMA (marked by bright green fluorescence) was seen in both KF
(Fig. 8a) and NF (Fig. 8b) study groups treated with conditioned media obtained from
KK/KF coculture. Actin fibres were more elongated and stressed in the cells treated with
KK/KF coculture conditioned media compared with those treated with non-coculture
41
conditioned media. KFs demonstrated significantly increased expression of α-SMA
compared with similarly exposed NF (Fig. 8).
• Contraction of KF populated collagen lattice by KK/KF coculture conditioned
media is attenuated by anti-TGF-β1 neutralizing antibody.
Study groups comprised collagen matrices populated by KFs from sample KF48 with
conditioned media collected from KK48/KF48 coculture, non-cocultured KKs, non-
cocultured KFs and KK48/KF48 coculture with anti-TGF-β1 antibody. At each interval
(6, 24 and 48 hrs) the percentage lattice contraction was calculated. Significant
contraction was seen in collagen lattice exposed to conditioned medium obtained from
KK/KF coculture compared with non-coculture samples. The addition of anti-TGF-β1
antibody to the conditioned medium obtained from KK/KF coculture, however,
significantly decreased the rate and extent of contraction of the collagen lattice (Fig. 9).
42
Figure 6: Effect of conditioned media collected from (KK)/(KF) cocultures on collagen contraction by KFs Collagen matrices composed of KFs (a) or NFs (b) were treated with conditioned media collected from non-cocultured KKs ,non-cocultured KFs (□) and KK/KF coculture (■). At each interval of study (1, 2, 3, 6, 24 and 48 h) the area of the gel was calculated as a percentage of the initial area. Each bar shows mean ± SEM of eight lattices pooled from four independent experiments using four different KF (a) or NF (b) cell strains.
43
Figure 7: Comparison of contraction of collagen gel lattice incorporated with KFs and NFs. The fibroblast-populated collagen lattices were collected after 48 hrs, stained with rhodamine B and photographed. (a) NF1 and (b) KF48 were treated with KK48/KF48 coculture conditioned media. (c) NF2 and (d) KF30 were treated with KK30/KF30 coculture conditioned media. (e) NF3 and (f) KF25 were treated with KK25/KF25 coculture conditioned media. (g) NF4 and (h) KF43 were treated with KK43/KF43 coculture conditioned media.
44
Figure 8: α-Smooth muscle actin (α-SMA) expression by KFs (a) or NFs (b). KFs (a) and NFs (b) were seeded at a density of 1000 cells per well in eight-well Lab-tek chamber slides in Dulbecco's modified Eagle's medium (DMEM)/10% fetal calf serum for 24 hrs. Medium was then replaced by serum-free DMEM for 48 hrs. After 48 hrs conditioned media pooled from keloid keratinocyte (KK)/KF coculture and non-coculture samples were added. This was followed by addition of anti-α-SMA antibody. A rhodamine counterstain was added to the fluorescein isothiocyanate-labelled secondary antibodies at a 1: 40 dilution. A bright green fluorescence marked the positive expression of α-SMA, while cells demonstrating minimal or no expression were stained dull orange. Each set represents KFs (a) or NFs (b) treated with conditioned media collected from KK/KF coculture.
45
Figure 9: Effect of anti transforming growth factor (TGF)-β1 neutralizing antibody on collagen contraction by KF48 induced by conditioned media collected from KK48)/KF48 coculture. Collagen matrices composed of KF48 were treated with conditioned media collected from KK48/KF48 coculture (checked ■), KK non-coculture (■), KF non-coculture (□) or KK48/KF48 coculture with anti-TGF-β1 antibody ( ). At each interval of study (6, 24 and 48 hrs) the area of the gel was calculated as a percentage of the initial area. Results are shown as mean ± SEM.
46
3.3 DISCUSSION In recent years, significant work has been done to support the hypothesis that epithelial–
mesenchymal interactions have an important governing role in keloid formation, and that
keloid formation may be the consequence of abnormal keratinocyte control over
fibroblasts rather than a defect of fibroblasts themselves (Yang et al., 2003). In this study
an established two-chamber coculture model was used to obtain conditioned media and
the effects of the latter was tested on FPCLs, an established three-dimensional wound
contraction and scar contracture model, to investigate the effect of paracrine secretions
from epithelial–mesenchymal interactions on collagen lattice contraction. Conditioned
media obtained from KK/KF coculture induced greater and faster contraction of collagen
lattices populated with KFs compared with lattices populated with NFs. It was also
observed that conditioned media obtained from non-cocultured KFs and KKs produced
decreased contraction of the collagen lattice. This underscores the importance of
paracrine secretions resulting from epithelial–mesenchymal interactions in wound repair,
and supports the role of overlying epidermal keratinocytes in modulation of KF
behaviour.
Transforming growth factor-beta has often been considered to be a key player in the
pathogenesis of keloids (Kim et al., 2000). Increased mRNA expression of TGF-β1 and
collagen has been observed in keloids (Kim et al., 2000). Previous studies demonstrated
that KFs produced increased TGF-β (Fusenig et al., 1994; Maas-Szabowski et al., 1999;
Mackenzie et al., 1994). Tissue fibrosis is thought to be the result of excessive ECM
deposition induced by TGF-β which signals fibroblasts to increase the synthesis of matrix
47
proteins, decrease the production of matrix-degrading proteases and increase the
production of inhibitors of these proteases (Border et al., 1996). There is evidence
suggesting that TGF-β1 can inhibit apoptosis in various types of cells (Smith et al., 1999;
Kim et al., 1998; Genestier et al., 1998). Chodon’s group (2000) reported that
dysregulation in the Fas-mediated apoptosis which occurs in normal wound healing may
be a factor contributing to scar formation and suggested a role for TGF-β1 in this
resistance; specifically that of prolonged proliferative phase due to generation of an
apoptosis resistant phenotype. As the epithelium regulates underlying mesenchymal cell
apoptosis (Hurle et al., 1986), epithelial-mesenchymal interactions could be one factor
responsible for this decreased apoptosis. Funayama and his co-workers (2003)
demonstrated through a similar coculture model that the keloid derived fibroblasts
brought about a greater proliferation and diminished apoptosis when cocultured with NK
and KK. They also demonstrated an upregulation of TGF-β1 in the fibroblasts cocultured
with keloid derived keratinocytes. Xia and his team (2004) demonstrated that the
upregulation of TGF-β1 in a previous study and observed that this was the net effect of an
increased TGF-β1 mRNA transcription in response to paracrine signaling between KK
and KF which was amplified in keloid versus normal cells. Their group postulated that an
aberration in this paracrine regulation in Keloid-derived cells could have resulted in the
generation of a positive feedback loop between KKs and KFs even after reepithelisation,
increasing the expression of TGF-β1 and related factors such as CTGF, which potentiate
TGF-β signaling.
TGF-β1 has also been demonstrated to enhance the contraction of collagen gels in vitro
(Montesano et al., 1998; Finesmith et al., 1990) and wound contraction in vivo
48
supporting its role in wound contraction. The effect of TGF-β is mainly through the Smad
family of proteins, although a Smad independent pathway involving signaling through
MAPK, ERK, JNK pathways has also been proposed (Derynck et al., 2003). In the
present study, we observed attenuation of FPCL contraction with the addition of anti-
TGF-β1 neutralizing antibody into conditioned media from KK/KF co-culture, indicating
the importance of TGF-β1 in keloid scar contracture.
Myofibroblasts, a group of actin rich fibroblasts, are known to play an important role in
wound contraction. Wound contraction is a very important step in wound repair involving
the conversion of proto-myofibroblasts to differentiated myofibroblasts by the production
of α-SMA in order to generate more force for contracture (Tomasek et al., 2002; Roy et
al., 2001). In normal wound repair, these myofibroblasts disappear by apoptosis but in
pathological conditions the myofibroblasts persist and continue to remodel the ECM,
resulting in connective tissue contracture (Tomasek et al., 2002). Since increased
expression of TGF-β1 directly induces α-SMA expression (Desmouliere et al., 1993;
1995; Ronnov-Jessen et al., 1993) and epithelial–mesenchymal interactions modulate
TGF-β1 expression, we hypothesized that epithelial–mesenchymal interactions could also
indirectly modulate α-SMA expression in vitro. In the present study
immunohistochemical analysis of α-SMA expression by KFs and NFs exposed to
conditioned media from KK/KF coculture and non-coculture conditioned media controls,
demonstrated increased expression of α-SMA in both KF and NF groups, supporting the
hypothesis.
49
To summarize, it has been demonstrated that the paracrine output from epithelial-
mesenchymal interactions between KKs and KFs modulate contraction of FPCL, strongly
suggesting its role in scar and wound contraction. TGF-β1 is likely to play an important
role in contraction of FPCL induced by epithelial-mesenchymal interactions. These data
suggest a therapeutic approach involving targeting the epidermal layer of the keloid
rather than its deeper constituents for the treatment of this difficult condition.
50
4 ROLE OF ACTIVIN-FOLLISTATIN SYSTEM IN KELOID PATHOGENESIS
51
4.1 INTRODUCTION
The bioactivity of fibroblasts in fibrogenesis or excessive scar formation is regulated by a
number of growth factors. The transforming growth factor-β (TGF-β) family, comprising
the TGF-β’s, bone morphogenetic proteins (BMPs) and activins, is thought to play a
central role by regulating fibroblast proliferation, differentiation and matrix production
(Derynck et al., 2003; Kim et al., 2000; Niessen et al., 1999; Tuan et al., 2000). Although
a series of studies has demonstrated the importance of TGF-β as a key player in keloid
pathogenesis, the function and role of activin-A have yet to be established.
Activins, like other members of the TGF-β superfamily, are dimeric proteins consisting
of two βA subunits (activin-A), two βB subunits (activin-B) or a βA and a βB subunit
(activin-AB) (Massague et al., 1990; Vale et al., 1990). Activin-A signals through
heterodimeric complexes of two receptor types - type I receptors, also called activin
receptor-like kinases (ALKs), and type II receptors which are transmembrane serine–
threonine kinases (Mathews et al., 1993). The biological action of activin-A is also
regulated by follistatin, a soluble activin-binding glycoprotein, which inhibits activin’s in
vitro and in vivo functions (Massague et al., 1990; Phillips et al., 2003).
Activin-A was discovered initially as a regulator of pituitary function (Vale et al., 1986).
Subsequent research has shown it to affect proliferation and differentiation of various cell
types. For example, it has been shown to stimulate proliferation of lung fibroblasts and
their differentiation into myofibroblasts (Ohga et al., 1996). It has also been
demonstrated to play a role in the proliferation and differentiation of keratinocytes in
vitro (Seishima et al., 1999). Activin-A inhibits proliferation of vascular endothelial cells
52
(Mathews et al, 1994; McCarthy et al., 1993; Kozian et al, 1997) and has been shown to
cause cell death in primary hepatocyte cultures (Schwall et al., 1993), B cell hybridomas
and mouse and human myeloma cells (Nishihara et al., 1993, 1995).
The first evidence for a role of activin-A in wound healing came from the studies by
Hübner’s group (1996). They demonstrated an increased induction of activin-A
expression within 24 hr after injury, which remained high until the end of the repair
process. Transgenic mice, which overexpressed the activin-βA subunit specifically in the
epidermis, had epidermal hyperthickening and dermal fibrosis (Munz et al., 1999). After
skin injury, these mice showed enhanced granulation tissue formation with strong
deposition of ECM below the hyperproliferative keratinocytes. By contrast, reduced
granulation tissue formation and thus delayed healing was observed in mice
overexpressing the activin antagonist, follistatin, in the epidermis (Wankell et al., 2000).
Several studies have implicated activin-A as an important player in fibrotic disorders.
Sugiyama’s team (1998) demonstrated an upregulation of activin-A in cirrhotic and
fibrotic rat livers. Furthermore, there was an increased expression of activin-A in various
pulmonary conditions associated with interstitial pulmonary fibrosis (Matsuse et al.,
1996). Activin-A has also been reported to act as a profibrotic cytokine in renal disease
(Gaedeke et al., 2005). Thus, although in vitro studies suggest that activin-A may play a
role in the repair of the mesenchyme and epithelium, a disruption in the control of
activin-A expression could lead to the development of tissue fibrosis.
The present study characterized the role of activin-A synthesis and signaling in keloid
pathogenesis. It evaluated the activin-follistatin system in regulating cellular proliferation
53
and expression of ECM components in keloid scars, using an extensive array of
techniques such as RNAse protection assay, immunohistochemistry, and co-cultures.
Overall, the expression and production of activin-A and its biological regulator,
follistatin, were perturbed in the development of human keloids.
4.2 RESULTS • Increased activin-A and follistatin in the basal layer of epidermis in keloid
tissues
Immnohistochemical analysis on paraffin sections of normal skin tissues and keloid scar
tissues was performed as previously mentioned. The sections were probed with
antibodies against activin-A (Oxford Bio-Innovations, Oxford, UK) and follistatin (Gift
from Dr David Phillips, Monash University, Melbourne, Australia) followed by
secondary antibodies. Both activin-A and follistatin were found to be localized to the
basal layer of the epidermis in both normal and keloid skins. However, an increased
intensity of these proteins was observed in keloid tissues compared with normal skin
tissues (Fig. 10). This suggested that the activin-follistatin regulatory system might be
implicated in keloid pathogenesis.
• Keloid scars expressed higher levels of activin-A mRNA compared with normal
skin
The expression profile of activin-A was studied using a RNase protection assay in keloid
scars and normal skin tissues (Fig. 11). Normal skins and treated scars expressed very
54
low levels of activin-A mRNA. In contrast, keloid scar samples had abundant levels of
activin-A mRNA.
• Increased production of activin-A and follistatin by KF.
NF and KF were cultured and conditioned media from these cultures were collected.
Activin-A was measured in conditioned media samples using a specific enzyme linked
immunosorbent assay (ELISA) (Knight et al., 1996) according to the manufacturer’s
instructions (Oxford Bio-Innovations, Oxfordshire, UK) with some modifications for
media samples as described previously (Buzzard et al., 2003). The average intra-plate %
coefficient of variation (CV) was 8.1% (n = 4 plates) and the inter-plate %CV was 6.4%.
The lower limit of detection was 0.01 ng/ml. Follistatin concentrations were measured
using a discontinuous radioimmunoassay as described previously (O’Connor et al., 1999)
with modifications for media samples. Conditioned media showed a linear dose-response
curve which was parallel to the standard. The average intra-assay %CV was 7.7% (n = 3
assays), the inter-assay %CV was 6.4% and the limit of detection was less than 1.94
ng/ml. When cultured in isolation, KFs demonstrated a 29-fold higher secretion of
activin-A compared with NFs (P < 0.0001; Fig. 12a). In contrast, NK produced
significantly higher activin-A than KK (P < 0.0001; Fig. 12a). For follistatin, there were
significant differences in secreted levels between KK and NK (P=0.019; Fig. 12b); KFs
demonstrated a 5-fold increase in secreted levels of follistatin compared with NFs (P <
0.0001; Fig. 12b).
55
• Over-expression of activin-A in HaCaT keratinocytes increased the proliferation
of underlying fibroblasts during co-culture
As the activin-follistatin regulatory system was shown to be upregulated in keloid tissues,
we determined if HaCaT keratinocytes overexpressing activin-A (hβAHaCaT) modulated
underlying fibroblasts in a paracrine manner. Normal human dermal fibroblasts
cocultured with these overexpressing hβAHaCaT keratinocytes (Gift from Prof Sabine
Werner, ETH, Zurich, Switzerland) demonstrated a 66% increase in proliferation
compared with non-cocultured NFs (P < 0.0001; Fig. 13). In comparison NFs cocultured
with neoHaCaT cells (cells transfected with the empty plasmid) (Gift from Prof Sabine
Werner, ETH, Zurich, Switzerland) demonstrated only a 30% increase (P < 0.0001; Fig.
13).
• Activin-A increased proliferation in NFs and KFs and stimulated expression of
ECM components
Given that keratinocytes overexpressing activin-A stimulated the proliferation of
underlying fibroblasts, the direct effect of activin-A on proliferation and ECM production
were tested. Treatment of NFs and KFs with activin-A (100 ng/ml) resulted in a 23 % (p
< 0.001) and 46% (p < 0.0001) increase, respectively, in proliferation compared with
untreated controls (Fig. 14). This was comparable to the proliferation brought about by
treatment with TGF-β1, although at a lower dose (10 ng/ml).
It was then determined if activin-A and follistatin affected the production of ECM
components like collagen, fibronectin and α-SMA in NFs and KFs. Using Western
56
blotting, an increase in collagen and α-SMA was observed in both NF and KF treated
with activin-A compared with untreated cells (Fig. 15a, 15b, 15c, 15d, 15g, 15h).
Elevated collagen and fibronectin levels in the conditioned medium in response to
activin-A was confirmed in KFs (Fig. 15e, 15f). To assess the neutralizing ability of
follistatin in this system, exogenous follistatin was co-administered with activin-A which
resulted in a marked reduction in the expression levels of collagen and α-SMA.
Follistatin treatment alone, unlike activin-A, significantly reduced the secreted levels of
collagen, strongly suggesting that follistatin was blocking the stimulatory effects of
endogenously produced activin-A (Fig. 15e). Activin when combined with TGF-β did not
show significant difference in expression of both collagen and α-SMA when compared to
activin only treatment (Fig 15g, Fig 15h).
57
Figure 10: Increased localization of activin-A and follistatin in the basal layer of epidermis in normal and keloid tissues. Paraffin sections of normal (a, b) and keloid (c, d) skin tissues were prepared and labeled with antibodies specific for activin-A (a, c) and follistatin (b, d) and observed with microscopy (100x). In each panel, the insert shows the same tissue labeled with a non-immune mouse antibody of the appropriate immunoglobulin isotype instead of the specific activin or follistatin antibody. Keloid tissues had more intense labeling in the basal layer of the epithelium compared with normal tissues. The localizations of activin-A and follistatin (or its lack in the negative control inserts) is shown by arrows. The dermis and epidermis are represented by ‘D’ and ‘E’, respectively
(a
(c
(b
(d
E
E
E
E
D
D D
D
E
E
E
D
D D
E
D
20µm 20µm
20µm 20µm
58
Figure 11: Expression of endogenous activin-βA mRNA derived from normal and keloid tissues. RNA was isolated from normal and keloid human tissues and analyzed by RNase protection assay for the presence of activin-A mRNA. Lanes 1, 2, 12: NS1, NS2, NS8 (normal skin). Lanes 3, 4, 5, 6, 7: KS4, KS5, KS6, KS7, KS11 (keloid scars from burn patients). Lane 8, 9: Keloid treated-skin before surgical excision. Lane 10, 11: KS5, KS19 (keloid scars from earlobe).
59
Figure 12: Elevated levels of activin-A obtained from keloid fibroblast cultures Conditioned media obtained from NF (NF1,NF3,NF4,NF5,NF9), KF (KF16,KF31,KF32,KF43,KF48), NK (NK1,NK3,NK4,NK5,NK9) normal and KK (KK16,KK31,KK32,KK43,KK48) were assayed using immunoassays specific for activin-A (a) or follistatin (b). Data are presented as mean ± SD.
60
Figure 13: Increased proliferation of NF when cocultured with activin-A overexpressing hβAHaCaT cells Immortalized hβAHaCaT cells overexpressing the βA subunit of activin (striped bar) and neoHaCaT cells with only the resistance plasmid (solid bar) were co-cultured with normal NF. The control cultures (open bar) comprised of non-cocultured KF. Data are means ± SD.
61
Figure 14: Increased proliferation of NF and KF treated with rhActivin-A Cultures of NF (a, b) or KF (c, d) were grown until 50% confluence and then serum starved for 48 hrs. The fibroblasts were then treated with activin-A (100 ng/ml, solid bars) TGF-β1 (1 ng/ml dotted bars), TGF-β1 (10 ng/ml, grey bars) then subjected to MTT proliferation assay. Untreated samples were used as control (open bars). Values are mean ± SD absorbance at 570 nm as a percentage of control.
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63
Figure 15: Effect of activin-A, follistatin or TGF-β1 on collagen, fibronectin and α-SMA expression NF or KF were cultured in the absence or presence of different concentrations of activin-A and TGF-β1. Cells were harvested, lysed and both cell lysate and conditioned media collected for Western blot analysis for collagen, fibronectin or α-SMA. (a) and (b) Lane 1: Untreated NF; Lane 2: NF + activin-A; Lane 3: NF + TGF-β1; Lane 4: NF4 + follistatin. (c) and (d) Lane 1: Untreated KF; Lane 2: KF + activin-A; Lane 3, 4: KF + TGF-β1; Lane 5: KF + follistatin; Lane 6: KF + follistatin + activin-A. (e) and (f) Lane 1: Untreated KF; Lane 2: KF + activin-A; Lane 3: KF + TGF-β1; Lane 4: KF + follistatin. All of the blots were probed with a β-actin antibody to confirm equal loading. For conditioned media 4 ml of conditioned media was concentrated from each treatment group to 100 µl which was used for analysis. Histograms represent % OD volume of the bands in the corresponding blots. Solid bars represent collagen; striped bars represent α-SMA; Open bars represent fibronectin. (g) Lane 1: Untreated KF; Lane 2: KF + activin-A; Lane 3: KF + TGF-β1; Lane 4: KF + activin-A (100 ng/ml) + TGF-β1 5 ng/ml. (h) Lane 1: Untreated KF; Lane 2: KF + activin-A 100 ng/ml; Lane 3: KF + activin-A 50 ng/ml; Lane 4: KF + activin-A (100 ng/ml) + TGF-β1 5 ng/ml; Lane 5: KF + TGF-β1 10 ng/ml.
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65
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4.3 DISCUSSION Activin-A, a member of the TGF-β superfamily, has been detected in high levels in
various inflammatory (Jones et al., 2004) and fibrotic disorders (Hübner et al., 1996;
Sugiyama et al., 1998; Matsuse et al., 1996, Gaedeke et al., 2005). To confirm its role in
keloid formation, the present study showed that activin-A was indeed overexpressed in
keloid tissues. For example, immunolocalization of activin-A was found in the basal
layers of the epithelium in human skin tissue, but interestingly, keloid tissues showed
more intense immunolocalization compared with normal tissues. Since epithelial–
mesenchymal interactions have recently been explored as a possible important factor
contributing to excessive scar formation, in our experimental set-up although there is no
staining for the fibroblasts, one cannot rule out the paracrine effect of Activin- A
expressed by KK on KF.
The activin-follistatin system is an important regulatory system controlling cellular
proliferation and differentiation in many epithelio-mesenchymal organs, including the
kidney, prostate, mammary gland, lung, pancreas, and salivary gland (Ritvos et al.,1995;
Hildén et al., 1994; Roberts et al., 1994; Tuuri et al., 1994; Cancilla et al., 2001; Liu et
al., 1996; Zhao et al., 1996; Miralles et al., 1998; Furukawa et al., 1995; Yamaoka et al.,
1998; Maldonado et al., 2000; Bläuer et al., 1996). When the localization of follistatin in
keloid and normal tissues was investigated, it was found to have similar pattern to that of
activin. The increase in follistatin in the basal layer of keloid tissue is likely due to a
short-loop negative feedback mechanism operative between activin-A and follistatin
(Phillips et al., 1998). Taking into consideration the localization and expression of both
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activin-A and follistatin in the basal layer of the epidermis, where mitotically-active
keratinocytes are present, one could hypothesize that the activin-follistatin system
modulates keratinocyte dynamics in tissue repair and keloid pathogenesis (Werner et al.,
2001).
Activin-A, in addition to being a potent activator of fibroblasts, has also been shown to
have an antiproliferative effect on various types of cells, including epithelial cells,
lymphocytes, prostate cancer cells, and others (Chen et al., 2002). However, it has been
observed that many carcinoma cells are somehow able to escape the negative regulatory
action of activin-A by activating production of follistatin (Dowling et al., 2000;
Rossmanith et al., 2002). In this study, activin-A was secreted in large amounts by KF
when compared with NF but KK secreted less. Interestingly, KK secreted more follistatin
than NK, suggesting that a mechanism similar to that seen in carcinoma cell types might
be active in KK, where these cells were secreting more follistatin to neutralize the
activin-A.
Previous work has demonstrated a large amount of ECM deposition below activin-A
over-expressing keratinocytes (Munz et al., 1999) suggesting that an autocrine, paracrine
and/or endocrine effect of activin-A might be active in the process of wound healing. In
the present study, we cocultured activin-A overexpressing keratinocytes (hβAHaCaT
cells) with NF. An increased proliferation in fibroblasts exposed to hβAHaCaT cells was
observed as compared to those exposed to both non-cocultured and neo HaCaT cells.
This could be due to the increased paracrine levels of activin-A in the hβAHaCaT /NF
coculture system as compared with NK cocultures.This data demonstrated that activin-A
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secreted from keratinocytes exerted a paracrine effect on the fibroblasts and that an
excess of activin-A might lead to over-proliferation of these cells, thus leading to fibrosis.
While analyzing this data one has to take in account that HaCaT cells in which activin-A
has not been overexpressed, has an endogenous level of this protein and several other
proliferative factors like IGF-1 (Insulin-like growth factor-1), TGF-β (transforming
growth factor-beta), interleukins and others. Hence one would expect them to also
increase the proliferation of the fibroblasts. Other groups have also observed such effects
when coculturing with neo\HaCaT cells. Skobe’s group (1998) observed that when
cocultured with HaCaT neo cells, fibroblasts show a 75% increase in cell numbers.
As further substantiation of a pro-proliferative effect of activin-A on human dermal
fibroblasts, exogenous activin-A increased proliferation of both NF and KF, consistent
with results from other groups (Ohga et al., 1996; Yamashita et al., 2004). Since TGF-β1
is known to be an important player in fibrotic disorders and has been demonstrated to be
highly expressed in KFs (Lee et al., 1999; Younai et al., 1994; Polo et al., 1999), it was
tested if activin-A and TGF-β1 might act synergistically to stimulate fibrotic processes.
However, co-treatment of cells with both activin-A and TGF-β1 showed no significant
synergistic effect.
ECM components are involved in many cellular processes and their patterns of
expression is different from that usually present in normal skin during different stages of
wound healing (Costa et al., 1999). Tissue fibrosis is thought to be partly the result of
excessive induction by TGF-β1, which signals fibroblasts to increase the synthesis of
ECM, decrease the production of matrix-degrading proteases and increase the production
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of the inhibitors of these proteases (Border et al., 1994). NFs and KFs, when exposed to
activin-A, increased expression of cytoplasmic collagen (procollagen). Procollagen
further undergoes post translational modifications (hydroxylation, glycosylation) and
subsequently forms a triple helix and is secreted out of the cell into the conditioned
medium. It was observed that there was an increased concentrations of secreted collagen
in the conditioned media from KFs when treated with activin-A, underscoring the
importance of activin-A in the regulation of collagen production by dermal fibroblasts in
keloid pathogenesis.
In addition, there was an increase in the expression of α-SMA, a phenotypic marker for
myofibroblasts (Yamashita et al., 2004), suggesting activin’s importance in regulating
expression and contractile activities of α-SMA. Not surprisingly, exogenous follistatin
had no effect on the expression of ECM components directly, but blocked the activin-
induced increases. Follistatin antagonizes the effect of activin-A by binding to activin
itself, rendering it inactive (Sugino et al., 1997; Phillips et al., 1998).
In summary, the present study demonstrated that activin-A was upregulated in keloid
tissues. Keratinocytes overexpressing activin-A stimulated proliferation of dermal
fibroblasts, suggesting a paracrine role to promote fibroblast proliferation, induce the
expression of α-SMA, and enhance collagen formation in KFs. However, follistatin, an
antagonist of activin-A, blocked the expression of ECM components. Thus, the findings
strongly suggest that activin-A is a potent inducer of fibroblast activation and involved in
the pathogenesis of keloids. They also emphasize the importance of follistatin in
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regulating activin-A bioactivity and suggest a possible therapeutic potential of follistatin
in the treatment and prevention of keloids (Fig 16).
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Figure 16: Activin-A expression increased by epithelial-mesenchymal interactions resulting in a fibrotic phenotype. KK expressed elevated levels of activin-A which has a paracrine effect on underlying fibroblasts thus increasing their proliferative potential and ability to produce ECM components which are charecteristics of a fibrotic phenotype. Follistatin can bind to activin-a and render it inactive thus having a potential as a possible therapeutic agent.
FOLLISTATIN THERAPY
KELOID SCAR
Proliferation
Collagen
Fibronectin
α-SMA
KELOID KERATINOCYTE
KELOID FIBROBLAST
+ +
Activin-A
Activin-A receptor
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5 ROLE OF PROTEOGLYCANS IN KELOID PATHOGENESIS
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5.1 INTRODUCTION
The TGF-β family of growth factors has been widely studied and is thought to play a
central role in tissue regeneration by regulating fibroblast proliferation, differentiation
and matrix production (Derynck et al., 2003; Kim et al., 2000; Niessen et al., 1999; Tuan
et al., 1998). However various other soluble and insoluble effectors are equally important
in wound repair and may thus play a part in keloid formation. One such effector group
comprises cell surface proteoglycans, namely the syndecans, which are a family of
transmembrane heparan sulphate proteoglycans that are differentially expressed during
development and wound repair (Carey et al., 1997; Gotte et al., Lories et al., 1992).
Owing to their unique function of integrating signaling from circulating growth factors
and ECM proteins with other cellular receptors like the integrins, they have often been
referred to as “tuners of transmembrane signaling” (Zimmermann et al., 1999). They
exist as four isoforms: syndecan-1, syndecan-2, syndecan-3 and syndecan-4, each having
characteristic cell- and tissue-specific distributions throughout development (Bernfield et
al., 1993; Kim et al., 1994).
Syndecan-2, also called fibroglycan, is especially abundant in mesenchymal cells (Gallo
et al., 2000) and has been shown to play an important role in diverse biological processes.
Mouse cells of mesenchymal origin in kidney, lung, and stomach and also in cells that
form cartilage and bone (David et al., 1993) are known to express high levels of
syndecan-2. Syndecan-2 modulates cell behavior by affecting cell adhesion and signaling
(Essner et al., 2006). Chen and his co-workers (2004) demonstrated that syndecan-2
regulates TGF-β signaling by binding to it through the core protein of its ectodomain. It
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has been shown to interact with various other cytokines and growth factors, like FGF-2
and VEGF, and mediate their effects on promoting mesenchymal tissue growth and
angiogenesis (Burgess et al., 1989; Leung et al., 1989; Besner et al., 1990; Clasper et al.,
1999).
Syndecan-2 has also been implicated in various types of cancers, with increased
expression demonstrated in human ovarian carcinoma biopsies (Davies et al., 2004) and
other cancer cell lines (Gulyas et al., 2004). It has also been shown to modulate colon
cancer cell adhesion, motility and proliferation (Park et al., 2002; Han et al., 2004).
Owing to these properties, syndecan-2 has recently become a subject of widespread
research. Its role in carcinogenesis, embryogenesis, wound healing and other clinical and
physiological conditions is being thoroughly explored.
In addition to heparan sulphate proteoglycans, soluble chondroitin and dermatan sulphate
proteglycans like decorin have also been shown to play a role in cell-matrix interactions.
Decorin is an abundant component of bone and skin ECM and associates with type I, II
and VI collagen fibrils as well as fibronectin in vivo (Laine et al., 2000; Lozzo et al.,
1996; 1998). Decorin is expressed abundantly in NF in which its expression has been
shown to be enhanced by IL-1 and dexamethasone and inhibited by TGF-β (Laine et al.,
2000; Heino et al., 1988; Kähäri et al., 1995; 1991). Unlike the syndecans, decorin
inhibits the biological activity of TGF-β by binding with the core protein in various cell
types (Hilderbrand et al., 1994) and thus has been explored as a potential antifibrotic
agent. Its antifibrotic effect has been demonstrated in experimental kidney, lung, cerebral
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and muscular fibrosis (Border et al., 1992; Giri et al., 1997; Logan et al., 1999;
Fukushima et al., 2001)
In addition to modulating cell-matrix interaction, decorin has also been shown to have an
antiproliferative effect on various carcinoma cells. The antiproliferative effect of decorin
was demonstrated to be mediated by either the upregulation of cyclin-dependent kinase
inhibitors like p21 and p27 or by interacting with various growth factors (De Luca et al.,
1996; Fischer et al., 2001). It was seen that malignant transformation of cells resulted in
loss of decorin expression. However when decorin expression was restored, it caused
malignant cell growth arrest, highlighting its protective effect on cellular dynamics
(Lozzo et al., 1999; Santra et al., 1995).
In the present study, intrinsic syndecan-2, FGF-2 and decorin expressions were studied in
keloid tissues by performing immunohistochemical analysis and western blot assays. The
role of serum on the expression profile of syndecan-2 and decorin was further
investigated by stimulating the NF and KF with 10% fetal calf serum. The role of FGF-2
in shedding of the syndecan-2 ectodomain and its implications in modulating cell
behavior was studied by treating KF cultures with recombinant FGF-2 and
immunoblotting the conditioned media for the expression of syndecan-2. The role of
epithelial-mesenchymal interactions in modulating syndecan-2, FGF-2 and decorin
expression dynamics was investigated using an established two-chamber serum-free
coculture model. In addition, the antifibrotic effect of decorin was studied by
investigating its effect on the expression of ECM components such as collagen,
fibronectin and α-SMA.
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5.2 RESULTS • Keloid scar intrinsically expressed higher levels of syndecan-2 and FGF-2 and
lower levels of decorin
The expression profiles of syndecan-2, FGF-2 and decorin in keloids and normal skin
tissues were studied using a western blot assay. Normal skin expressed low levels of
syndecan-2 (Fig. 17) and FGF-2 (Fig. 22) however decorin was abundantly expressed
(Fig. 25). In contrast, keloid scar samples had abundant levels of syndecan-2 (Fig. 17),
FGF-2 (Fig. 22) and low levels of decorin (Fig. 25). Although decorin was
downregulated in keloid tissues a 32kda band was observed in this sample which was
completely missing in normal tissue.
• Increased localization of FGF-2 seen in the basal epidermis and dermis of
keloids compared to normal skin
Paraffin sections of normal skin and keloid scar were subjected to immunohistochemical
analysis for the expression of FGF-2. Increased levels of FGF-2 were observed in the
dermis and basal epidermis of keloid scar tissues compared to normal skin (Fig. 21ab).
FGF-2 also stained strongly in keloid dermis whereas the dermis of normal skin
demonstrated significantly diminished or negative staining (Fig. 21c d).
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• Serum growth factors upregulated syndecan-2 and downregulated decorin
expression in KF
Stimulation with 10% FCS is interpreted by fibroblasts as a wound healing signal, thus
NF and KF were treated with 10% FCS to investigate if syndecan-2 and decorin were
involved in the initial phases of wound healing. Both NFs and KFs showed increased
levels of syndecan-2 (Fig. 18) on stimulation with serum growth factors, suggesting a
possible role of syndecan-2 in the initial wound healing mechanism. However, decorin
expression was downregulated on stimulation with serum (Fig. 26).
• Coculture of KK with KF increased levels of syndecan-2 in the conditioned
media
To investigate if epithelial-mesenchymal interactions modulate syndecan-2 expression, a
coculture model was set up wherein NK and KK were cocultured with NF and KF
respectively. Fibroblasts were subsequently lysed and total cell extracts were subjected to
western blot. Although no difference was observed in cellular levels of syndecan-2 in
both NFs AND KFs cocultured with their respective keratinocytes (Fig. 19a), an
increased level of syndecan-2 was observed in the conditioned media of KK/KF
cocultures compared to KFs in monoculture (Fig. 19b) suggesting the possible cleavage
of syndecan-2 in KFs upon coculture with KKs.
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• Cocultured keratinocytes and fibroblasts demonstrated increased expression of
FGF-2 compared to cells in monoculture
In order to investigate the effect of epithelial-mesenchymal interactions on the dynamics
of FGF-2 expression, NK and KK were cocultured with NF and KF respectively, with
cell lysates collected from both groups and blotted for FGF-2 expression. An increase in
FGF-2 expression was observed in both cocultured keratinocytes (Fig. 23a) and
fibroblasts (Fig. 24) as compared to those in monoculture, highlighting a possible
autocrine effect of FGF-2 on keratinocytes as well as a paracrine effect on the underlying
fibroblasts. In addition, cocultured KKs expressed higher levels of FGF-2 as compared to
cocultured NKs (Fig. 23b) suggesting a more active mechanism within KKs compared to
NK.
• Increased levels of decorin found only in the conditioned media from NK/NF
cocultures as compared to monocultures
Although conditioned media obtained from the coculture of NF with normal NK
demonstrated higher levels of decorin compared to normal fibroblast monocultures (Fig
27a), no significant difference in decorin levels was observed in the conditioned media of
cocultured KFs as compared to KFs in monoculture (Fig 27b) suggesting a loss KK
control of underlying KF decorin expression.
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• rhFGF-2 increased syndecan-2 cleavage and thus its detectable levels in KF
conditioned media
The role of FGF-2 in the cleavage of fibroblast syndecan-2 resulting in its increased
levels in conditioned media was investigated by treating KFs with exogenous FGF-2 and
analyzing the conditioned media for expression of syndecan-2 by western blot. A
significant increase in the levels of syndecan-2 was observed compared to untreated KFs
(Fig. 20).
• Decorin decreased the expression of ECM components in both NF and KF
To investigate the effect of decorin on the expression of ECM components decorin at
various concentrations (250 ng/ml, 500 ng/ml, 1000 ng/ml and 2000 ng/ml) was added to
both NF and KF after which cell lysates and conditioned media were analyzed for
collagen, fibronectin and α-smooth muscle actin expressions. A decrease in the
expressions of the ECM components was observed both in the cell lysate (Fig. 28a b) and
conditioned media (Fig. 28c).
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Figure 17: Elevated levels of syndecan-2 in tissue extracts obtained from keloid tissue. Expression of syndecan-2 in normal and keloid skin tissues were analyzed by western blot assay. Study groups comprised of normal skin and keloids. Tissue specimens were sonicated in lysis buffer and 100 µg of total protein extracts were subjected to SDS-PAGE and western blotting. Blots were hybridized with anti-syndecan-2 polyclonal antibody Lane 1-5: Keloid scar (K50, K51, K52, K53, and K54). Lane 6-8: Normal skin (N31, N32, N33). Equal loading was confirmed by blotting with β-actin.
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Figure 18: Serum growth factors upregulated syndecan-2 expressions in NF and KF. Expression of syndecan-2 by fibroblasts after exposure to 10% FCS was detected by western blot assay. Fibroblasts were seeded in 6-well plates at a density of 1 × 104 cells/ml in DMEM/10% FCS for 24 hrs and subsequently in serum-free medium for another 48 hr. The fibroblasts were subject to either DMEM/10% FCS or serum-free DMEM for 24 hr. Fibroblasts were lysed and 100µg total protein cell lysates were subjected to SDS-PAGE and western blotting. Blots were hybridized with anti-syndecan-2 polyclonal antibody. Lane 1: NF4+DMEM, Lane 2: NF4+DMEM/10% FCS, Lane 3: NF14+DMEM, Lane 4: NF14+DMEM/10%FCS, Lane 5: KF45+DMEM, Lane 6: KF45+DMEM/10%FCS, Lane 7: KF49+DMEM, Lane 8: KF49+DMEM/10% FCS. Equal loading was confirmed by blotting with β-actin for cell extracts
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Figure 19: Increased ectodomain shedding of syndecan-2 is observed in keloid cocultures as compared to monocultures. a) Cell lysate obtained from NFs and KFs cocultured with NKs and KKs was assayed by western blot for syndecan-2 expression. Cell lysates from NF and KF in monoculture were used as controls. No difference in the expression of syndecan-2 was observed in cocultured fibroblast cell extracts compared to fibroblasts in monoculture. Lane 1: NF1 non cc, Lane 2: NK1/NF1, Lane 3: NF2 non cc, Lane 4: NK3/NF2, Lane 5: KF 45 non cc, Lane 6: KK45/KF45, Lane 7: KF 48 non cc, Lane 8: KK48/KF48. b) Conditioned media from KF cocultures were assayed by western blot for syndecan-2 expression. Conditioned media from KFs in monoculture were used as controls. Equal volumes of conditioned media were subjected to SDS-PAGE. Conditioned media from KF cocultures shed more syndecan-2 into the conditioned media as compared to the KF monocultures. Lane 1: KF48 non cc, Lane 2: KK48/KF48, Lane 3: KF45 non cc, Lane 4: KK45/KF45, Lane 5: KF24, Lane 6: KK24/KF24.
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Figure 20: Exogenous rhFGF-2 stimulated shedding of syndecan-2 from fibroblast cell surface into the conditioned media. rhFGF-2 (10 ng/ml) was added to the fibroblast cultures and conditioned media assayed for the expression of syndecan-2. Conditioned media from untreated KFs were used as controls. Equal volumes of conditioned media were subjected to SDS-PAGE. KFs treated with FGF-2 shed more syndecan-2 as compared to untreated fibroblasts. Lane 1: KF43 non cc, Lane 2: KF43+FGF-2, Lane 3: KF29 non cc, Lane 4: KF29+FGF-2.
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Figure 21: Increased localization of FGF-2 in the basal layer of epidermis and dermis in keloid tissues. Paraffin sections of normal (a, c) and keloid (b, d) tissues were prepared and labeled with antibodies specific for FGF-2 and observed with microscopy (40x). In each panel, the insert shows the same tissue labeled with a non-immune rabbit antibody of the appropriate immunoglobulin isotype instead of the specific FGF-2 antibody. Keloid tissues had more intense labeling in the basal layer of the epithelium and the dermis compared with normal skin. The localization of FGF-2 (or its lack in the negative control inserts) is shown by an arrow. The dermis and epidermis are represented by ‘D’ and ‘E’, respectively.
10 µm 10 µm
10 µm 10 µm
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Figure 22: Elevated levels of FGF-2 in tissue extracts obtained from keloid tissue. Expression of FGF-2 in normal and keloid skin tissues were analyzed by western blot assay. Study groups comprised of normal skin and keloids. Tissue specimens were sonicated in lysis buffer and 100 µg of total protein extracts were subjected to SDS-PAGE and western blotting. Blots were hybridized with anti-FGF-2 polyclonal antibody Lane 1-8: Keloids (K50, K51, K52, K53, K54, K55, K56 and K57). Lane 9-13: Normal skin (N31, N32, N33, N34, N35). Equal loading was confirmed by blotting with β-actin.
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Figure 23: Cocultured NK and KK expressed increased levels of FGF-2 as compared to monocultured keratinocytes. Cell lysates from monocultured NK and KK were extracted and analyzed by western blot for the expression of FGF-2. Similarly, NKs and KKs which had been cocultured with NFs and KFs respectively were lysed and cell extracts were analyzed by western blot for the expression of FGF-2. a) Lane 1: NK1, Lane 2: NK1/NF1, Lane 3: KK48, Lane 4: KK48/KF48, b) Lane 1: NK1/NF1, Lane 2: NK2/NF2, Lane 3: NK3/NF3, Lane 4: KK48/KF48, Lane 5: KK45/KF45, Lane 6: KK49/KF49. Equal loading was confirmed by blotting with β-actin for cell extracts.
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Figure 24: Cocultured NF and KF expressed increased levels of FGF-2 as compared to cells in monoculture. Cell lysates from monocultured NFs and KFs were extracted and analyzed by western blot for the expression of FGF-2. Similarly, NFs and KFs which had been cocultured with NKs and KKs respectively were lysed and cell extracts were analyzed by western blot for the expression of FGF-2. Cocultured fibroblasts expressed increased levels of FGF-2 compared to moncoultures. Equal loading was confirmed by blotting with β-actin for cell extracts Lane 1: NF1, Lane 2: NK1/NF1, Lane 3: NF2, Lane 4: NK2/NF2, Lane 5: KF48, Lane 6: KK48/KF48, Lane 7: KF49, Lane 6: KK49/KF49
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Figure 25: Downregulation of decorin in tissue extracts obtained from keloid tissue. Expression of decorin in normal and keloid skin tissues were analyzed by western blot assay. Study groups comprised of normal skin and keloids. Tissue specimens were sonicated in lysis buffer and 100µg of total protein extracts were subjected to SDS-PAGE and western blotting. Blots were hybridized with anti-decorin monoclonal antibody Lane 1-3: Keloids (K51, K52 and K53). Lane 4-6: Normal skin (N31, N32, N33). Equal loading was confirmed by blotting with β-actin.
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Figure 26: Serum growth factors downregulated decorin expressions in NF and KF. Expressions of decorin by fibroblasts after exposure to 10% FCS was detected by western blot assay. Fibroblasts were seeded in 6-well plates at a density of 1 × 104 cells/ml in DMEM/10% FCS for 24 hrs and subsequently in serum-free medium for another 48 hrs. The fibroblasts were subject to either DMEM/10% FCS or serum-free DMEM for 24 hr. Fibroblasts were lysed and 100 µg total protein cell lysates were subjected to SDS-PAGE and western blotting. Blots were hybridized with anti-decorin monoclonal antibody. Lane 1: NF4+DMEM, Lane 2: NF4+DMEM/10% FCS, Lane 3: NF14+DMEM, Lane 4: NF14+DMEM/10%FCS, Lane 5: KF45+DMEM, Lane 6: KF45+DMEM/10%FCS, Lane 7: KF49+DMEM, Lane 8: KF49+DMEM/10% FCS. Equal loading was confirmed by blotting with β-actin for cell extracts.
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Figure 27: Increased secretory decorin in cocultured NFs and not KFs. NF and KF were cocultured with NK and KK, respectively and conditioned media were collected and analyzed for the expressions of decorin. a) Lane 1: NF4+DMEM, Lane 2: NF1+DMEM, Lane 3: NF14+DMEM, Lane 4: NF3+DMEM, Lane 5: NK1/NF1, Lane 6: NK2/NF2, Lane 7: NK14/NF14, Lane 8: NK5/NF5, b) Lane 1: KF29+DMEM, Lane 2: KF48+DMEM, Lane 3: KF49+DMEM, Lane 4: KF25+DMEM, Lane 5: KK48/KF48, Lane 6: KK45/KF45, Lane 7: KK25/KF25, Lane 8: KK49/KF49
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Figure 28: Effect of decorin on collagen, fibronectin and α-SMA expressions. NF or keloid fibroblasts KF were cultured in the absence or presence of different concentrations decorin. Cells were harvested, lysed and both cell lysates and conditioned media collected for western blot analysis for collagen, fibronectin or α-SMA. a) Lane 1: NF1+DMEM, Lane 2: NF1+Decorin (250 ng/ml), Lane 3: NF1+Decorin (500 ng/ml), Lane 4: NF1+Decorin (1000 ng/ml), Lane 5: NF1+Decorin (2000 ng/ml), b) Lane 1: KF48+DMEM, Lane 2: KF48+Decorin (250 ng/ml), Lane 3: KF48+Decorin (500 ng/ml), Lane 4: KF48+Decorin (1000 ng/ml), c) Lane 1: KF48+DMEM, Lane 2: KF48+Decorin (250 ng/ml), Lane 3: KF48+Decorin (500 ng/ml), Lane 4: KF48+Decorin (1000 ng/ml).
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5.3 DISCUSSION
The healing of any skin injury involves a spontaneous cascade of events comprising
communication between cells and the release of mediators as growth factors, and
cytokines, leading to inflammation, cell proliferation, collagen-ECM deposition, scar
formation, maturation and ultimately remodeling. Any abnormality or disruption of this
sequence of events may lead to formation of abnormal scarring which may include keloid
scars, characterized by excessive collagen-ECM deposition and often, the formation of
contractures.
Recent advances in this field have seen the role of growth factors being extensively
investigated. Interestingly, researchers have found most of the growth factors and
cytokines involved in the wound healing process to be immobilized at the cell surface and
ECM through binding with proteoglycans (Rouslahti et al., 1991). These proteoglycans in
turn are known to play important roles in various physiological processes. Besides
providing mechanical strength by filling in the space between the collagen and elastin
fibres by absorbing water, they also influence collagen formation, cell proliferation, cell
migration and cell adhesion during wound healing (Pratibha et al., 2000).
Syndecan-2, a cell surface proteoglycan, plays an important role in the transmodulation
of the ECM assembly. Since keloid scars are characterized by an aberrant organization of
the ECM assembly, we investigated if syndecan-2 was being abnormally expressed in
keloid tissues as compared to normal skin tissues. Western blot analysis of the tissue
extracts demonstrated that syndecan-2 was indeed upregulated in keloid tissues and that it
could be one of the factors responsible for the excess deposition of ECM components.
95
In order to understand the underlying biology of syndecan-2 in wound repair and scar
formations, the expression profiles of syndecan-2 at different stages of the wound healing
process was investigated using in-vitro models. Upon cutaneous injury, the coagulation
cascade is activated and an inflammatory response sets in as the first stage of wound
healing. The active platelet plug sequesters serum in the immediate vicinity providing a
rich source of cytokines/growth factors with paracrine effects on neighbouring cells. Such
exposure of fibroblasts to serum is interpreted to be a physiological wounding signal
(Iyer et al., 1999). It was observed that when NF and KF when exposed to serum growth
factors expressed increased amounts of syndecan-2 suggesting its possible role in the
early stages of the healing process and in the modulation of the ensuing events by
interaction with some of the macrophage-derived growth factors like FGF-2, VEGF, EGF
(Gotte et al., 2003). The hypothesis that syndecan-2 might be an important player in the
early phase of the healing process is further strengthened by numerous studies reporting
syndecan-2 to be induced by proinflamatory cytokines like IL-1α and IL-1β and TGF- β
which are abundant in the early wound environment (Clasper et al., 1999; Worapamorn
et al., 2001; Sebestyen et al., 2000).
Strikingly enough, it has been observed that during development, syndecan-2 is localized
at sites where intensive epithelial-mesenchymal interactions shape and transform the
epithelia and the mesenchyme into morphologically and functionally differentiated
tissues suggesting a possible syndecan-2’s role in modulating epithelial-mesenchymal
interactions (David et al., 1993). Previous findings have shown that epithelial-
mesenchymal interactions play an important role in keloid pathogenesis with KK
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modulating NF and KF growth and proliferation (Lim et al., 2001), influencing collagen
expression by both NF and KF, inducing NF to secrete collagen in a keloid-like manner
(Lim et al., 2002), and utilizing the IGF system of mitogens as inductive ligands for this
process. Thus to investigate if these interactions would in anyway affect the expressions
of syndecan-2, NF and KF were cocultured with NK and KK, respectively, and the cell
lysates and conditioned media blotted. Surprisingly, although no difference in the
expressions of syndecan-2 from cell lysateswas observed, a definite increase in syndecan-
2 levels was seen in the conditioned media obtained from KFs cocultured with KK. This
strongly suggests that syndecan-2 was being shed from the KF cell surface into the
conditioned media under the influence of the overlying keratinocytes. Syndecan-1 and -4
have been previously shown to be shed from the surface of mouse epithelia and SVEC4-
10 cells in vitro, and factors released during stress, injury and cancer have been
demonstrated to upregulate this process (Fitzgerald et al., 2000; Elenius et al., 2004;
Bayer et al., 2001; Seidel et al., 2000; Subramanian et al., 1997). Previous studies by
Fears’s group (Fears et al., 2006) highlighted the role of syndecan-2 shedding induced by
various factors, to promote angiogenesis. Of the various factors studied, FGF-2 was
observed to stimulate syndecan-2 shedding from microvascular endothelial cells. When
exogenous rhFGF-2 was added to the KF cultures, syndecan-2 shedding was also
observed, with increased amounts of secreted syndecan-2 found in the conditioned media.
The production and secretion of heparin binding growth factors such as FGF have been
implicated in various macrophage-mediated processes, including tumor angiogenesis,
arteriogenesis and capillary sprouting, wound healing, and inflammation (Clasper et al.,
1999). It would appear that the syndecan-FGF system could also be an important player
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in initiating aberrant wound healing processes leading to fibrosis. As rhFGF-2 increased
syndecan-2 shedding in KFs, it was investigated if FGF-2 was intrinsically upregulated in
keloid tissues compared to normal skin and, more importantly, in KK/KF cocultures
where increased levels of syndecan-2 were observed in conditioned media. Increased
expressions and localizations of FGF-2 in keloid epidermis and dermis was found as
compared to normal skin sections. In addition, fibroblasts cocultured with NK and KK
expressed increased levels of FGF-2 as compared to fibroblasts in single cell culture.
Interestingly KK/KF cocultures expressed increased levels of FGF-2 as compared
NK/NF. One arm of keloidogenesis resulting from epithelial-mesenchymal interactions
would thus appear to involve increased FGF-2 levels which in turn stimulate the shedding
of syndecan-2 from fibroblasts. This shed syndecan-2 would not only modulate the
activity of FGF-2 but also other profibrotic growth factors (Han et al., 2004).
FGF-2 has also been demonstrated to decrease decorin mRNA expression in
osteosarcoma cells. The reduced expression of decorin corresponded with increased
collagen production by these cell types (Bodo et al., 2002; Wegrowski et al., 2000).
Since keloids are marked by increased production of type 1 collagen the expression of
decorin in keloid tissues was investigated. As expected a decreased expression of decorin
was observed in keloid tissues as compared to normal skin. Interestingly a 32 kda band
was observed in keloid tissues probably due to the degradation of decorin to an inactive
fragment by various MMP’S known to be present in keloids (Fujiwara et al., 2005).
Previous studies have shown MMP’s to degrade decorin to an inactive fragment which is
then unable to further bind to TGF-β, releasing it from the ECM with resultant increased
activity in vivo (Imai et al., 1997). Further studies will be required to assess if this
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mechanism plays a role in keloid pathogenesis. When stimulated with 10% serum, a
reduction in decorin was observed in both NF and KF. This may be due to negative
feedback from increased levels of TGF-β and other growth factors in the serum. Previous
results by Zandvoort’s group (Zandvoort et al., 2006) demonstrated TGF-ß1 stimulation
to decrease decorin production in isolated pulmonary parenchymal fibroblasts of COPD
patients.
The effect of epithelial-mesenchymal interactions in modulating the expression of
decorin in a coculture model was investigated and it was observed that although NK/NF
cocultures expressed increased amounts of decorin compared to monocultures, KK/KF
cocultures did not show any significant difference in decorin levels as compared to
monocultures. This would suggest that the overlying keratinocytes might have directly or
indirectly blocked the mechanisms responsible for modulating fibroblast decorin
expression. FGF-2 demonstrated above to be upregulated in cocultures might well be one
of the factors responsible, and further studies will be required to confirm this hypothesis.
Finally, it was investigated if exogenous administration of decorin could be used as an
anti- fibrotic agent. NF and KF were treated with decorin at different concentrations and
expressions of different ECM components were investigated. It was observed that
collagen was downregulated on treatment in a dose-dependent manner both in the cell
lysates and the conditioned media. In addition α-SMA, a phenotypic marker for
myofibroblasts, and fibronectin were also downregulated on treatment with decorin,
which would suggest the potential utility of decorin against excess fibrosis.
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Overall, it was shown that syndecan-2 and FGF-2 were not only overexpressed in keloid
tissues but could also interact with each other resulting in the shedding of syndecan-2
which could in turn activate a whole cascade of events resulting in keloidogenesis (Fig
29). In addition, decorin appeared to be downregulated in keloid tissues, and this protein
with strong antifibrotic effects could have the potential to be used as a therapeutic agent
for keloids.
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Figure 29: Ectodomain shedding of syndecan-2 by FGF-2 stimulated by epithelial-mesenchymal interactions resulting in a fibrotic phenotype One arm of keloidogenesis results from epithelial-mesenchymal interactions which appear to involve increased FGF-2 levels which in turn stimulate the shedding of syndecan-2 from fibroblasts. This shed syndecan-2 not only modulates the activity of FGF-2 but also other profibrotic growth factors leading to keloid scars.
Keloid Fibroblasts
Stimulates Release of FGF-2
FGF-2
FGF-2 receptor
Syndecan -2
Keloid Keratinocytes
+ +
ECM
Cleaved syndecan -2 binds to profibrotic factors
KELOID SCAR
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6 ROLE OF STEM CELL FACTOR and c-KIT SYSTEM IN KELOID PATHOGENESIS
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6.1 INTRODUCTION Stem cell factor (SCF), also known as mast cell growth factor, steel factor or KIT ligand
is a hematopoietic and tissue growth factor that serves as a ligand for the c-KIT oncogene
(Gabrilove et al., 1994). It is widely expressed throughout the body by stromal cells,
fibroblasts and endothelial cells (Ashman et al., 1999) and is detectable at a low level in
the circulation (Broudy et al., 1997; Lyman et al., 1998). KIT (CD117) on the other
hand, coded by proto-oncogene c-KIT, is a 145–160 kDa transmembrane tyrosine kinase
belonging to the type III family of receptor kinases (Yarden et al., 1987). KIT is
expressed in a variety of normal cells and tissues (Natalie et al., 1992; Lammie et al.,
1994) often with its ligand (Williams et al., 1990). Previous studies have demonstrated
expression of c-KIT mRNA or protein by mast cells (Mayrhofer et al., 1987; Majumdar
et al., 1988; Nocka et al., 1989), melanocytes (Nocka et al., 1989), testis (Majumdar et
al., 1988), and in bone marrow (Wang et al., 1989). In addition c-KIT protein expression
has been found in non-haemopoietic cell types, including vascular endothelial cells
(Broudy et al., 1994), astrocytes, renal tubules, breast glandular epithelial cells and sweat
glands (Natalie et al., 1992; Lammie et al., 1994). Binding of SCF to KIT results in
dimerisation of the receptors followed by activation of its intrinsic tyrosine kinase
activity (Blume-Jensen et al., 1991). The interaction between c-KIT and SCF ligand
plays an important role in hematopoiesis, in mast cell development and function,
embryogenesis, proliferation and migration of primordial germ cells and melanoblasts
(Huang et al., 1990; Zsebo et al., 1990).
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SCF and its receptor c-KIT also play important roles in the process of wound healing.
SCF-KIT interactions in mast cells trigger degranulation at the site of wounding. The
released mediators increased fibroblast proliferation and collagen synthesis, contributing
to abnormal tissue growth leading to fibrosis in various organs (Garbuzenko et al., 2002;
Fireman et al., 1999; El Kossi et al., 2003). In addition to their role in fibrosis, autocrine
or juxtacrine cycles involving c-KIT and SCF have been demonstrated to be important in
some solid tumors, for example, small cell lung cancer (Krystal et al., 1996; Hibi et al.,
1991).
In the present study, owing to its function in wound healing, we hypothesized a role for
SCF/c-KIT system in keloid scar pathogenesis. Moreover, its upregulation in other
fibrotic disorders compels its investigation as a possible factor responsible for a keloidic
phenotype.The results in this study demonstrated that both SCF and c-KIT were
upregulated in keloid scar tissues in vivo thus corroborating our hypothesis. They were
also upregulated in cultured fibroblasts on stimulation with serum highlighting their
importance in the initial phase of wound healing. In addition it was demonstrated that
epithelial-mesenchymal interactions, mimicked by coculture of keratinocytes and
fibroblasts in vitro, not only stimulated secretion of soluble form of SCF in keloid
cocultures but also brought about shedding of the extracellular domain of c-KIT perhaps
by upregulation of TACE. TACE was also shown to be upregulated in keloid scars in
vivo and keloid cocultures in vitro. In addition the KIT receptor was activated, as
demonstrated by increase in phospho c-KIT, in keloid cocultures as compared to normal
cocultures.
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6.2 RESULTS • Elevated expression of SCF, c-KIT, phosphorylated c-KIT (Tyr 703, Tyr 721)
and TACE in keloid scar tissues as compared to normal skin tissues
Stem cell factor (SCF) (Fig. 30) and both mature (145kDa) and soluble (95kDa) forms of
its receptor c-KIT (Fig. 33a) were found to be overexpressed in keloid scar tissue extracts
as compared to normal. In addition phosphorylated forms of c-KIT (tyr 703, tyr 721)
were also over expressed in keloid scar tissues though the difference was significant only
in tyr 721 (Fig. 33b). TACE or ADAM17, a known sheddase of many membrane proteins
was also shown to be expressed at higher levels in keloid scar tissues compared to normal
skin tissues (Fig. 37).
• Increased localization of c-KIT in the keloid epidermis compared to normal skin
Immunohistochemical studies were carried out to investigate the expression of c-KIT and
TACE in normal skin and keloid scar tissues. An increased level of c-KIT expression was
observed in the basal epidermis (rete-ridge) of keloid scar. The normal skin epidermis
demonstrated significantly less or negative staining (Fig. 34a b).
• Serum growth factors upregulated SCF levels in KF
Stimulation with 10% FCS is interpreted by fibroblasts as a wound healing signal thus
NF and KF were treated with 10% FCS to investigate if SCF was involved in the initial
phases of wound healing. Both NF (Fig 31a) and KF (Fig 31b) demonstrated an increase
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in levels of SCF on stimulation with serum growth factors suggesting a possible role of
SCF in the initial wound healing mechanism.
• Coculture of KK with KF increased levels of SCF in the conditioned media
To investigate if epithelial-mesenchymal interactions modulate SCF expression, a
coculture model was set up wherein KK were cocultured with KF. Fibroblasts were
subsequently lysed and total cell extracts were subjected to western blot. Although no
difference was observed in cellular levels of SCF in KF cocultured with KK (Fig. 32a),
an elevated level of SCF was observed in the conditioned media of KK/KF cocultures
compared to fibroblasts in monoculture (Fig. 32b).
• Upregulation of c-KIT in KKs as compared to Nks
Normal and keloid cells were cultured for 24 hrs followed by serum starvation for 48 hrs.
The cells were subsequently treated with growth medium for 24 hrs and subjected to flow
cytometry. KKs expressed higher levels of c-KIT as compared to NKs (Fig. 35ab).
• Increased levels of c-KIT in cocultured keratinocyte/fibroblast conditioned
media as compared to monocultures
NKs and KKs were cocultured with NFs and KFs respectively and the cell lysates from
both the cell types were collected and blotted for c-KIT. A downregulation of c-KIT
expression was observed in both cocultured keratinocyte (Fig 36a) and fibroblast cell
lysates (Fig 36b) as compared to monocultures. Phospho c-KIT (tyr 721) was however
upregulated in keloid cocultured cell lysates as compared to normal coculture lysates
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indicating activation of c-KIT in coculture conditions (Fig 36e). The conditioned media
obtained from the cocultured fibroblasts demonstrated elevated levels of c-KIT
suggesting a possible cleavage of c-KIT into the conditioned media upon coculture (Fig
36c). In addition higher cleaved c-KIT levels were observed in conditioned media
obtained from keloid cocultures as compared to normal cocultures suggesting an
enhanced shedding mechanism prevalent in keloid cocultures (Fig 36d).
• Elevated expression of TACE in cocultured KFs as compared to cocultured NFs
An increased expression of mature TACE was observed in total cell lysates obtained
from KFs coculutred with KKs as compared to NFs cocultured with NKs suggesting an
activation of this enzyme system in keloids (Fig. 38).
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Figure 30: Elevated levels of SCF in tissue extracts obtained from keloid tissues Expression of SCF in normal and keloid skin tissues were analyzed by western blot assay. Study groups comprised of normal skin and keloids. Tissue specimens were sonicated in lysis buffer and 100 µg of total protein extracts were subjected to SDS-PAGE and western blotting. Blots were hybridized with anti SCF polyclonal antibody Lane 1-5: Keloid scar (K50, K51, K52, K53). Lane 6-8: Normal skin (N31, N32, N33). Equal loading was confirmed by blotting with β-actin
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Figure 31: Serum growth factors upregulated SCF expression in normal and keloid fibroblasts. Expression of SCF by fibroblast after exposure to 10% FCS was detected by Western blot assay. Fibroblasts were seeded in 6-well plates at a density of 1 × 104 cells/ml in DMEM/10% FCS for 24 hrs and subsequently in serum-free medium for another 48 hrs. The fibroblasts were subject to either DMEM/10% FCS or serum-free DMEM for 24 hrs. Fibroblasts were lysed and 100µg total protein cell lysates were subjected to SDS-PAGE and western blotting. Blots were hybridized with anti-SCF polyclonal antibody. a) Lane 1: NF4+DMEM, Lane 2: NF4+DMEM/10% FCS, Lane 3: NF14+DMEM, Lane 4: NF14+DMEM/10%FCS. b) Lane 1: KF45+DMEM, Lane 2: KF45+DMEM/10%FCS, Lane 3: KF49+DMEM, Lane 4: KF49+DMEM/10% FCS. Equal loading was confirmed by blotting with β-actin for cell extracts.
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Figure 32: Increased levels of SCF in conditioned media obtained from keloid cocultures as compared to monocultures a) Cell lysates obtained from KFs cocultured with KKs was assayed by western blot for SCF expression. Cell lysate from keloid fibroblasts in monoculture were used as controls. No difference in the expression of SCF was observed in cocultured fibroblast cell extracts compared to fibroblasts in monoculture. Lane 1: KF 49 non cc, Lane 2: KK49/KF49, Lane 3: KF 48 non cc, Lane 4: KK48/KF48. b) Conditioned media from KF cocultures were assayed by western blot for SCF expression. Conditioned media from KF in monoculture were used as controls. Equal volumes of conditioned media were subjected to SDS-PAGE. Conditioned media from KF cocultures demonstrated higher levels of SCF as compared to the KF monocultures. Lane 1: KF48 non cc, Lane 2: KK48/KF48, Lane 3: KF49 non cc, Lane 4: KK49/KF49
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Figure 33: Elevated levels of c-KIT and Phospho c-KIT (tyr 703, tyr 721) in tissue extracts obtained from keloid tissues. Expression of c-KIT and Phospho c-KIT at sites tyr 703 and tyr 721 in normal and keloid skin tissues were analyzed by western blot assay. Study groups comprised of normal skin and keloid scars. Tissue specimens were sonicated in lysis buffer and 100 µg of total protein extracts were subjected to SDS-PAGE and western blotting. Blots were hybridized with anti-c-KIT polyclonal, Phospho c-KIT tyr 703, Phospho c-KIT tyr 721 monoclonal antibodies. a) Lane 1-5: Normal skin (N31, N32, N33, N34, N35) Lane 6-13: Keloids (K50, K51, K52, K53, K54, K55, K56, K57). b) Lane 1-2: Normal skin (N31, N32) Lane 3-5: (K50, K51, K52)
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Figure 34: Increased localization of c-KIT in the basal layer of epidermis of keloid tissues. Paraffin sections of normal (a) and keloid (b) tissues were prepared and labeled with antibodies specific for c-KIT and observed with microscopy (40x). In each panel, the insert shows the same tissue labeled with a non-immune mouse antibody of the appropriate immunoglobulin isotype instead of the specific c-KIT antibody. Keloid tissues had more intense labeling in the basal layer of the epithelium compared with normal skin. The localization of c-KIT (or its lack in the negative control inserts) is shown by an arrow. The dermis and epidermis are represented by ‘D’ and ‘E’, respectively.
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Figure 35: c-KIT upregulated in KK as compared to NK. NK and KK were cultured, trypsinised and then probed with c-KIT monoclonal antibody followed by anti FITC conjugated secondary antibody. c-KIT was upregulated in keloid keratinocytes as compared to NK. a) NK 103, b) KK48
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Figure 36: Increased ectodomain shedding of c-KIT in KK/KF cocultures. Cell lysates from monocultured NK and KK were extracted and analyzed by western blot for the expression of c-KIT. Similarly, NK and KK which had been cocultured with NF and KF respectively were lysed and cell extracts were analyzed by western blot for the expression of c-KIT. Similarly NF and KF in monoculture and coculture were probed with anti-c-KIT and anti phospho c-KIT (tyr 721) antibody and subjected to western blot. Conditioned media from normal and keloid cocultures were collected and analyzed for the expression of c-KIT. Conditioned media from monocultures was used as control. a) Lane 1: NK1 Day 1, Lane 2: NK1/NF1 Day 1, Lane 3: NK1 Day 2, Lane 4: NK1/NF1 Day 2, Lane 5: NK1 Day 3, Lane 6: NK1/NF1 Day3, Lane 7: NK1 Day 4, Lane 8: NK1/NF1 Day 4, Lane 9: NK1 Day 5, Lane 10: NK1/NF1 Day 5, Lane 11: KK 48 Day 1, Lane 12: KK48/KF48 Day 1, Lane 13: KK48 Day 2, Lane 14: KK48/KF48 Day 2, Lane 15: KK48 Day 3, Lane 16: KK48/KF48 Day 3, Lane 17: KK48 Day 4, Lane 18: KK48/KF48 Day 4, Lane 19: KK48 Day 5, Lane 20: KK48/KF48 Day 5. b) Lane 1: NF1, Lane 2: NK1/NF1, Lane 3: NF2, Lane 4: NK2/NF2, Lane 5: KF48, Lane 6: KK48/KF48, Lane 7: KF49, Lane 8: KK49/KF49. Equal loading was confirmed by blotting with β-actin for cell extracts. c) Lane 1: KF 48, Lane 2: KK48/KF48, Lane 3: KF 49, Lane 4: KK49/KF49. d) Lane 1: NK1/NF1, Lane 2: NK2/NF2, Lane 3: NK3/NF3, Lane 4: NK8/NF8, Lane 5: KK48/KF48, Lane 6: KK49/KF49, Lane 7: KK45/KF45, Lane 8: KK 44/KF44. e) Lane 1: NK3/NF3, Lane 2: NK8/NF8, Lane 3: KK48/KF48, Lane 4: KK49/KF49
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Figure 37: TACE overexpressed in keloid scar compared to normal skin Expressions of TACE in normal and keloid skin tissues were analyzed by western blot assay. Study groups comprised of normal skin and keloid scars. Tissue specimens were sonicated in lysis buffer and 100 µg of total protein extracts were subjected to SDS-PAGE and western blotting. Blots were hybridized with anti-TACE polyclonal antibody. Lane 1-6: Keloids (K50, K51, K52, K53, K54, K55, K56, K57), Lane 7-11: Normal skin (N31, N32, N33, N34)
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Figure 38: Upregulation of TACE in KF cocultures as compared to NF cocultures Expression of TACE in cocultured NF and KF were analyzed by western blot assay. Lane 1: NK1/NF1, Lane 2: NK2/NF2, Lane 3: NK3/NF3, Lane 4: NK8/NF8, Lane 5: KK48/KF48, Lane 6: KK49/KF49, Lane 7: KK45/KF45, Lane 8: KK 44/KF44.
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6.3 DISCUSSION The keloid is a dermal form of FPD characterized by increased proliferation of
fibroblasts, as well as, increased production of collagen in vitro (Cohen et al., 1990;
Ladin et al., 1995). Apoptosis-related genes and proteins are believed to be
downregulated or mutated in keloid tissues and fibroblasts (Sayah et al., 1999; Chodon et
al., 2000). In this study, the role of SCF and its receptor c-KIT was investigated in keloid
pathogenesis both in normal skin and keloid tissue sections, as well as in keratinocyte and
fibroblast cultures. A tyrosine kinsase inhibitor, gleevec was explored as a possible
therapeutic agent for keloids.
In order to determine the overall expressions of SCF in keloid and normal skin tissues, an
immunoblot analysis was performed on the tissue extract obtained from keloid and
normal skin. It was observed that SCF (50kda and 25kda forms) were highly expressed in
keloids compared to normal tissues, supporting the hypothesis that SCF might play a
crucial role in keloid pathogenesis. In addition, both mature (145kDa) and soluble
(95kDa) forms of KIT were found to be upregulated in keloid tissues suggesting a
possible aberration in receptor-ligand interaction. The tyr703 and tyr721 phosphorylation
sites have been previously demonstrated to be responsible for the activation of Ras/Erk
and PI3-Kinase pathways, respectively. A number of studies have implicated the critical
importance of the Ras/Erk pathway in cell division and survival (Lewis et al., 1998).
Adaptor proteins like Grb2 can directly associate with phosphorylated tyr703 in KIT
which may lead to activation of subsequent downstream pathways (Thommes et al.,
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1999). Activation of PI3-Kinase via KIT has been linked to mitogenesis, differentiation,
survival, adhesion, secretion and actin cytoskeleton reorganization (Yee et al., 1993;
Timokhina et al., 1998; Serve et al., 1995; Kubota et al., 1998; Vosseller et al., 1997). It
was found that tyr721 when phosphorylated, interacts directly with PI3-Kinase (Serve et
al., 1995). In this study, both tyr 703 and tyr 721 phosphorylation sites of c-KIT were
found to be upregulated in keloid scar tissues compared to normal skin tissues, although
the difference was significant only in case of tyr 721.
In order to study the localization of the KIT protein in the skin, immunohistochemistry
was performed to investigate the relative expressions of KIT protein in both keloid and
normal skin. An increased staining at the basal layer of keloid epidermis was observed
compared to normal skin. Thus, as the basal layer consists of mitotically active
keratinocytes, KIT could be playing a major role in modulating the dynamics of these
active and undifferentiated keratinocytes. These results were confirmed by flow
cytometry which demonstrated that KKs expressed increased levels of c-KIT as
compared to NKs.
When fibroblasts are exposed to serum, they interpret it to be a physiological wounding
signal (Iyer et al., 1999). Hence, exposure to serum reflects how fibroblasts would react
in vivo in the early phase of wounding. The increase in SCF observed upon exposure to
serum in both NFs and KFs suggested that SCF might have a role in early wound healing
process. This is consistent with previous studies, which found that SCF peaked on the
first day of wounding. The increase in SCF production increased the activation of mast
cells and triggered their degranulation, leading to the release of factors such as IL-1α and
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TNF-α that lead to inflammation, which is an early response in wound healing (Huttunen
et al., 2002).
Previous findings have shown that epithelial-mesenchymal interactions play an important
role in keloid pathogenesis with KK modulating the growth and proliferation of NF and
KF (Lim et al., 2001), influencing collagen expression by both NF and KF, inducing NF
to secrete collagen in a keloid-like manner (Lim et al., 2002), and utilizing the IGF
system of mitogens as inductive ligands for this process. Thus to investigate if these
interactions in anyway affected expression of SCF and its receptor c-KIT, NF and KF
were cocultured with NK and KK and blotted. No significant differences in SCF
expression between the fibroblast cell lysates derived from coculture and monoculture
was observed. However an increase in SCF levels was detected in the conditioned media
derived from the cocultures. ADAM19 and ADAM33 in COS and HEK293 cells
(Chesneau et al., 2003; Zou et al., 2004), MMP- 9 in hematopoetic stem cells (Heissig et
al., 2002), IL1-β produced by keratinocytes (Da silva et al., 2002) have all been shown to
promote cleavage and secretion of SCF. The soluble SCF can then act as a proliferative
agent (Ren et al., 2003). The mechanism by which epithelial-mesenchymal interactions
brought about an increase in secreted levels of SCF is not know and further studies are
needed to be unravel to delineate the mechanism. The role of the interactions between the
mesenchyme and the epithelium on the expression of c-KIT was further investigated.
Although the phosphorylated levels of c-KIT (tyr 721) were elevated in keloid cocultures
as compared to normal cocultures, both the keratinocytes and fibroblasts which had been
cocultured expressed decreased amounts of c-KIT compared to that from monoculture.
However the conditioned media obtained from these cocultures demonstrated elevated
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levels of the receptor suggesting a shedding of c-KIT. Keloid cocultures demonstrated
increased levels of soluble c-KIT as compared to normal cocultures. To investigate the
mechanism by which c-KIT was cleaved, a sheddase TACE, which was previously
demonstrated to be critical for shedding of c-KIT (Cruz et al., 2004) was explored. Not
only an increase of mature active TACE in keloid tissue extracts compared to normal
tissue extracts was observed but also found it to be upregulated in cocultured KF cell
lysates compared to cocultured NF confirming the prevalence of a TACE mediated
shedding mechanism. Although the presence of an ectodomain shedding exists in keloid
cocultures, the role of soluble c-KIT in keloid pathogenesis is still unclear. It has been
previously reported that binding of SCF results in rapid kinase activity-dependent
receptor endocytosis, ubiquitination and degradation (Yee et al., 1993; 1994; Miyazawa
et al., 1994). Activation of the receptor also results in cleavage and release of the soluble
extracellular domain (Yee et al., 1993; 1994; Miyazawa et al., 1994; Brizzi et al., 1994)
which then competes for the ligand with the membrane bound form (Broudy et al., 1997).
These mechanisms are activated as a negative feedback response to regulate ligand
receptor induced signal transduction. A similar mechanism may exist in keloid cocultures
where increased activation of c-KIT, as seen by increased level of phospho c-KIT, results
in ectodomain shedding by proteolytic cleavage. There have been numerous studies
demonstrating presence of soluble c-KIT, as a negative feedback response, in serum in
various disorders (Tajima et al., 1998; Kawakita et al., 1995). Alternative to the negative
feedback hypothesis, the soluble c-KIT bound to the free SCF ligands in the ECM, may
also help to increase the retention of the ligand in the extracellular environment, resulting
in a prolongation of the half-life due to the increased size and stability of the complex
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(Broudy et al., 1997; Peters et al., 1996). By prolonging the overall half-life of the SCF
ligand, the soluble KIT receptors may then serve to stabilize free SCF levels between
peaks and valleys of SCF secretion, leading to the sustained activation of the SCF-KIT
pathways (Heaney et al., 1995). Thus to confirm the exact role of ectodomain shedding
observed, more mechanistic studies are warranted to ascertain the behavior and effect of
the soluble KIT receptor-SCF complexes in vivo.
In summary, it has been shown for the first time that SCF and its receptor c-KIT were
both upregulated and activated in keloid tissues. In addition, epithelial–mesenchymal
interactions seemed to modulate both the dynamics of SCF and c-KIT resulting in their
shedding into the conditioned media, a process mediated probably by TACE (Fig 39a,
39b). Given these results the SCF/c-KIT system might be a target for therapeutic
intervention for keloid scars.
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Figure 39: c-KIT system in keloid pathogenesis a) SCF loop: Under the influence of the overlying keratinocytes, fibroblasts produce soluble SCF which not only has an autocrine effect but also a paracrine effect on the keratinocytes thus forming a short positive loop system between the two.. b) c-KIT shedding: Epithelial-meenchymal interactions result in mature KIT (145kDa) to be cleaved by proteolytic enzymes (TACE) thus releasing soluble KIT (95kDa) into the ECM. The soluble c-KIT then binds to SCF. The soluble KIT can be either a result of a negative feedback mechanism to regulate the overactivated scf/c-KIT system in keloidic cells or can be a mechanism in itself to activate the KIT system by increasing the half life and stability of SCF by binding to it and resulting in its sustained effect.
SCF C-KIT TACE
a)
KELOID KERATINOCYTES
KELOID FIBROBLAST
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b)
Keloid Fibroblast
c-Kit ectodomain
Shed c-Kit either binds to SCF
Shed kit as bio marker
Keloid Keratinocytes
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7 INVESTIGATION OF GLEEVEC AS A THERAPEUTIC AGENT FOR KELOID SCARS
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7.1 INTRODUCTION In the past, a wide variety of treatments has been used for keloids, including intralesional
corticosteroids, surgery, cryosurgery, silicone gel sheeting, radiotherapy, pressure
therapy, and laser therapy like flashlamp pulsed-dye laser treatment. Newer therapies like
interferon α2b, bleomycin injections and intralesional 5-fluorouracil (5FU) have also
been explored. Unfortunately, various side-effects like severe pain, ulceration, necrosis
and hyperpigmentation limit the utility of all these treatments (Nanda et al., 2004, Mustoe
et al., 2002). Neither of the mentioned therapies proved to be effective for patients,
underscoring the need to discover alternatives.
Having identified SCF and its tyrosine kinase receptor c-KIT to be upregulated and
activated in keloid pathogenesis, it was explored if a tyrosine kinase inhibitor could be
used as a possible therapeutic agent for keloid scars. In recent years, biopharmaceutical
research has actively targeted protein kinases for the treatment of several neoplastic
diseases. Two FDA approved examples are Gleevec™ (Imitanib Mesynate – Novartis
AG) for the treatment of chronic myeloid leukemia and gastrointestinal stromal tumors
and Iressa™ (Gefitinib - AstraZeneca) for the treatment of metastatic non-small cell lung
cancer by inhibiting intracellular tyrosine kinase phosphrylation.
Gleevec essentially inhibits four kinases: Bcr-abl, c-abl, c-KIT and PDGF (Capdeville et
al., 2002). As imatinib is an inhibitor of the receptor tyrosine kinases for PDGF and SCF,
it is able to block PDGF and SCF-mediated cellular events (Lefevre et al., 2004).
Gleevec in particular has been previously demonstrated to be effective for the treatment
of chronic myeloid leukemia (CML) by blocking cell proliferation and inducing
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apoptosis in Bcr-abl- expressing CML. It has been effective against gastrointestinal
stroma tumor (GIST) owing to its inhibitory action on KIT (Sattler et al., 2004). It has
also been demonstrated to have antifibrotic effects in organ fibrosis like lung fibrosis
(Aono et al., 2005) and renal fibrosis (Wang et al., 2005).
In the present study, the effects of various concentrations of gleevec on the proliferation
of KK and KF was explored by MTT assay. Studies were performed to further investigate
if gleevec was able to inhibit the c-KIT pathway by blotting with antibodies against
phospho c-KIT and the ensuing molecules in the PI-3 kinase pathway. The expression of
the ECM components like collagen, fibronectin and α-smooth muscle actin and the
expressions of profibrotic and pro-angiogenic growth factors like VEGF, TGF-β, HDGF,
FGF-2, SCF on treatment with gleevec were also monitored. To investigate if gleevec
could reduce scar contracture, an in-vitro model which consisted of a FPCL was used and
its contraction on treatment with gleevec studied. Finally the effect of gleevec on the
bioenergetics of the cell was investigated by studying the intracellular ATP and the rate
of intracellular ATP synthesis in KF on treatment with gleevec.
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7.2 RESULTS • No significant difference in the proliferative potential of KK and KF on
treatment with gleevec
Gleevec was added in varying concentrations (0.5, 1, 2.5, 5, 10, and 20 µg/ml) in
presence of 5% v/v serum to KK and KF and MTT assay was performed. No significant
difference was observed in the proliferative potential of both keratinocytes and fibroblast
on treatment with gleevec (Fig. 40).
• Gleevec inhibited the SCF/c-KIT pathway by dowregulating the
phosphorylation of c-KIT at tyr 721, Akt and mTOR
KFs were treated with 5 µg/ml of gleevec in DMEM containing 5% v/v serum for 15
min, 1 hr and 24 hrs.Total extracts from cells were probed with antibodies against
phosphorylated forms of c-KIT (Tyr 721), Akt and mTOR. A decrease in the expression
of phospho c-KIT, phospho Akt and phospho mTOR was observed within 15 minutes of
treatment with gleevec (Fig. 41).
• Gleevec downregulated the expressions of ECM and profibrotic factors in KFs
in a dose-dependent manner
Gleevec in presence of 5% v/v serum was added in varying concentrations (2.5, 5, and
10µg/ml) to KFs and the total cell extracts subjected to western blot. Significant decrease
in levels of collagen, fibronectin and α-SMA were observed on treatment with gleevec in
a dose-dependent manner (Fig. 42a). A significant decrease in expression levels of
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profibrotic factors like VEGF, HDGF, TGF-β1, SCF, and FGF-2 were also observed
(Fig. 42b).
• Gleevec decreased the contraction of collagen lattice populated by KF in a dose -
dependent manner.
FPCL were prepared as previously described and treated with various conentrations of
gleevec in a medium containing 5% v/v serum for 48 hrs. The medium was subsequently
removed and the lattices stained by rhodamine solution to give it a pink colour. The
lattices were then photographed and the area calculated as previously mentioned for side-
by-side analysis. It was observed that collagen lattices populated with Keloid-derived
fibroblasts demonstrated significantly lesser contraction when treated with gleevec as
compared to collagen lattices treated with medium containing 5% v/v serum which was
used a control (Fig 43). Lattices in serum-free media were used a negative controls.
• Gleevec reduced the rate of ATP synthesis and intracellular ATP levels in KFs.
KFs were treated with gleevec (5 µg/ml) for 24 hrs and subjected to luciferin-luciferase
assay. The rate of ATP synthesis in KF was decreased on treatment with gleevec at a
concentration of 5 µg/ml. In addition the intracellular ATP levels in KFs were also
significantly reduced at similar concentrations of gleevec (Fig 44).
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Figure 40: No significant difference in the proliferative potential of KK and KF. KKs and KFs were treated with different concentrations of gleevec in 5% serum (0.5, 1, 5, 10, 20 µg/ml). Cells were subjected to MTT assay after 24hrs of treatment. Untreated fibroblasts in 5% serum were used as control. Fibroblasts in DMEM only were used as a negative control. No significant difference in proliferation was observed.a) KK48. b) KF48.
130
Figure 41: Gleevec downregulated expression of Phospho Akt, Phospho mTOR, Phospho c-KIT (tyr 721). KFs were treated with gleevec (5 µg/ml) and the cell lysates were collected after 15 min, 1 hr, and 24 hrs of treatment. The lysates were blotted with antibodies against Phospho Akt, Phospho mTOR, Phospho c-KIT (tyr 721). Gleevec downregulated the expression of phosphor Akt, Phospho mTOR and Phospho c-KIT (tyr 721) after 15 min. KFs treated with 5% serum were used as control. Lane 1: KF48 +DMEM with 5% serum after 15 min, Lane 2: KF48 + gleevec (5 µg/ml) in 5% serum after 15 min, Lane 3: KF48 +DMEM with 5% serum after 1 hr, Lane 4: KF48 + gleevec (5 µg/ml) in 5% serum after 1 hr, Lane 5: KF48 +DMEM with 5% serum after 24 hrs, Lane 6: KF48 + gleevec (5 µg/ml) in 5% serum after 24 hrs. Equal loading was confirmed by blotting with β-actin.
131
Figure 42: Gleevec downregulated expression of collagen, fibronectin, α-SMA, VEGF, HDGF, TGF-β1, SCF, FGF-2. Keloid fibroblasts were treated with different concentrations of gleevec in 5% serum (2.5µg/ml, 5µg/ml). After treatment for 24hrs the cells were lysed and blotted with antibodies against extracellular matrix components like a) collagen, fibronectin, α-SMA and b) profibrotic factors like TGF-β1, SCF, VEGF, HDGF, FGF-2. Untreated fibroblasts in 5% serum were used as controls. Lane 1: KF48 +DMEM with 5% serum, Lane 2: KF48+ 2.5µg/ml gleevec in 5% serum, Lane 3: KF48+ 5µg/ml gleevec in 5% serum. Equal loading was confirmed by blotting with β-actin.
132
Figure 43: Gleevec significantly reduced the contraction of collagen gel lattice incorporated with keloid fibroblasts. The fibroblast populated collagen lattices were treated with various concentrations of gleevec (0.5µg/ml, 2.5µg/ml, 5µg/ml, 10µg/ml) for 48hr. The lattices were then collected and stained with Rhodamin B and photographed. a) KF 48+DMEM, b) KF48 +DMEM with 5% serum, c) KF48+ 0.5µg/ml gleevec in 5% serum, d) KF48+ 2.5µg/ml gleevec in 5% serum, e) KF48+ 5µg/ml gleevec in 5% serum, f) KF48+ 10µg/ml gleevec in 5% serum.
133
Figure 44: Gleevec significantly reduced the intracellular ATP and the rate of ATP synthesis in KF. KFs were treated with gleevec (5 µg/ml) for 24 hrs and analysed for the levels of intracellular ATP. The rate of ATP synthesis was also calculated in the treated samples with untreated samples as control. Open bar represents untreated KFs and solid bars represent treated KFs.
134
7.3 DISCUSSION In this study, no significant difference in the proliferative potential of KK and KF on
treatment with various concentrations of gleevec was observed. This was in accordance
with previous studies by Distler and his co-workers (2006) in which they reported that
gleevec showed no effect on the proliferative potential of dermal fibroblasts. However
gleevec was able to reduce the expression of ECM components, like collagen,
fibronectin, α-SMA significantly in KF in a dose-dependent manner highlighting its
importance as a potential antifibrotic agent. Gleevec also downregulated TGF-β and
VEGF which have been previously shown to be upregulated in KF (Kim et al., 2000; Xia
et al., 2004; Ong et al., 2007). It was dowregulated expression of HDGF which was
previously reported to be involved in organ development and modeling (Mori et al.,
2004) and recently demonstrated to be upregulated in keloid tissues (unpublished). FGF-2
and SCF, growth factors implicated in various fibrotic disorders (Yu et al., 2003;
Garbuzenko et al., 2002), were also downregulated upon treatment with gleevec in a
concentration-dependent manner. the effects of gleevec on FPCL, an established 3-d
wound contraction and scar contracture model (Bell et al., 1979) was further tested.
Gleevec was significantly reduced the contraction of the lattices thus highlighting its role
in modulating scar contracture.
Having demonstrated an antifibrotic effect of gleevec, it was explored if gleevec owed its
inhibitory activity to c-KIT receptor inactivation. Activation of PI-3 kinase via KIT has
been linked to mitogenesis, differentiation, survival, adhesion, secretion and actin
cytoskeleton reorganization (Yee et al., 1993; Timokhina et al., 1998; Serve et al., 1995;
135
Kubota et al., 1998; Vosseller et al., 1997). It was found that tyr721 when
phosphorylated, interacts directly with PI-3 kinase (Serve et al., 1995). Within 15 min of
treatment with gleevec, the expression of phospho c-KIT (tyr 721) in KF was reduced. In
addition, downstream signaling molecules like phospho Akt and phospho mTOR were
also downregulated in these KF suggesting an inhibition of the P13 kinase pathway in
these fibroblasts on treatment with gleevec.
Ueda’s team (1999) previously demonstrated a high level of ATP for a longer time in KF.
Thus we investigated if gleevec could reduce the intracellular ATP levels as well as the
rate of ATP production in KF. It was observed that gleevec could significantly decrease
both the rate of ATP production, as well as total intracellular ATP levels in KFs.
However the mechanism by which it has such an effect needs to be explored.
To summarize, this study demonstrated that a selective tyrosine kinase inhibitor, gleevec,
was able to downregulate the expression of ECM components, as well as profibrotic
factors in KFs via inhibition of the c-KIT mediated PI3 kinase signaling pathway. It was
also able to make the KFs less bioenergetic by reducing the ATP levels. Thus given its
effects, it could well be used as a potential therapeutic agent for keloids.
136
GENERAL CONCLUSION
The achievement of this project is the identification and elucidation of novel molecular
pathways in the pathogenesis of keloid scars. Using a large database of clinical tissue
samples from keloid and normal patients and various in vitro models of wound repair, it
was demonstrated for the first time that activin-A, stem cell factor (SCF) and its tyrosine
kinase receptor c-KIT, fibroblast growth factor-2 (FGF-2) and proteoglycans like
syndecan-2 and decorin play important roles in keloid pathogenesis and could be possible
targets for therapeutic intervention. Development of specific antibodies that antagonize
these growth factors or block its receptor could result in reduced collagen accumulation,
predicated on the results of this study. Similarly, based on these results, there is a scope
for development of peptide antagonists or receptor decoy molecules based on molecular
modeling of the receptor–ligand binding characteristics. Another approach would be to
screen for small molecules, potentially identifying compounds that antagonize the
receptors and its downstream signaling.
In addition to identifying these new targets, this study also underscores the importance of
epithelial-mesenchymal interactions in keloid pathogenesis. The interaction between the
dermis and the overlying epidermis was demonstrated to be critical in scar contracture
and in modulating the biology of activin-A, SCF/c-KIT, FGF-2, syndecan-2 and decorin
in keloid scars. This observation may have immense clinical relevance. Currently
available therapy involves the intralesional corticosteroid injection which is very painful
for the patients. Having demonstrated the importance of the epithelium, topical
application of chemical agents and antagonists of profibrotic growth factors could be
137
used as novel delivery systems thus making the therapy more patient compliant and
friendly.
Another interesting data which sprouted from this study was the shedding of the c-KIT
receptor in keloidic conditions which had been previously shown to occur in malignant
disorders. Keloids, although ‘benign’, have always been acknowledged to possess growth
autonomy to a certain degree resembling malignant tissues, resulting in the invasion and
involvement of adjacent skin in the course of its growth, and thus the clinical description
of the keloid as a scar that exceeds the boundaries for the original wound or incision. The
finding of c-KIT receptor shedding might well reflect growth autonomy in keloid
fibroblasts resembling transformed cells.
This study went on to explore new pharmacological interventions. This resulted in the
identification of growth factors, like follistatin and decorin, and chemical agents like
gleevec, as potential therapeutic agents for keloid scars. Follistatin is a natural antagonist
of activin-A and having identified activin-A as a player in keloids, it could be used as a
potential therapy. In this study follistatin was shown to downregulate the expression of
ECM components like collagen, fibronectin and α-SMA. Another growth factor which
was shown to be effective in downregulating ECM significantly was decorin. The
observation that decorin is downregulated in keloid scars and exuberantly expressed in
normal skin underscores its importance in normal wound repair. Finally having identified
a tyrosine kinase receptor namely c-KIT to be important in keloid pathogenesis, a well
known FDA-approved tyrosine kinase inhibitor, gleevec, was explored. Using in vitro
models for anti-scar research experimentation, it was found that gleevec could reduce the
138
expression of ECM components, as well as key profibrotic and pro-angiogenic growth
factors like VEGF, TGF-β, and HDGF. In addition it was also able to downregulate SCF
and FGF-2, which were also identified in this study, as key regulators of keloidogenesis.
Gleevec reduced contractile s of KF and also rendered the fibroblast less bioenergetic. At
the molecular levels, gleevec was shown to effectively block the PI-3 kinase mediated
SCF/c-KIT pathway. Figure 45 summarizes the growth factors involved in keloid
pathogenesis and the various possible targets for therapeutic intervention elucidated by
our research.
139
Figure 45: Summary of growth factors involved in keloid pathogenesis and possible therapeutic targets.
I
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Appendix 1
List of patients
NORMAL NF/NK1 NF2 NF/NK4 NF8 NF/NK9 NF/NK5 NF/NK3 NS31 NS32 NS33 NS34 NS35 KELOID KF6 KF46 KF3 KS/KF/KK48 KF/KK31 KF/KK32 KF/KK43 KF/KK16 KS/KF/KK 25 KF/KK30 KF/KK45 KF/KK24 KF/KK29 KS50 KS51 KS52 KS53 KS54 KS55 KS56 KS57
F M M F F F F F M M F M F M F F F M M F M F F M F M F M M F M F
Chinese Indian Chinese Malay Chinese Chinese Chinese Malay Chinese Chinese Chinese Malay Chinese Chinese Malay Indian Chinese Indian Chinese Chinese Indian Chinese Malay Malay Indian Malay Chinese Indian Malay Malay Chinese Chinese
27 years 32 years 3 years 28 years 24 years 35 years 12 years 13 years 20 years 29 years 35 years 33 years 16 years 36 years 20 years 23 years 17 years 28 years 34 years 19 years 25 years 30 years 23 years 21 years 18 years 10 years 18 years 21 years 28 years 16 years 25 years 22 years
Breast Abdominal Earlobe Breast Breast Breast Groin Groin Forearm Thigh Breast Forearm Earlobe Forearm Earlobe Earlobe Earlobe Chest Elbow Earlobe Forearm Face Forearm Chest Earlobe Face Earlobe Elbow Face Forearm Earlobe Earlobe
- - - - - - - - - - - - 1 1 1.5 1.5 0.5 2 1 1.5 1 1 1.5 2 1 1 1.5 1 1.5 1 2 1
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