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Heparan sulfatedependent ERK activation contributes to the overexpression of fibrotic proteins and enhanced contraction by scleroderma fibroblasts.

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Vol. 58, No. 2, February 2008, pp 577–585
DOI 10.1002/art.23146
© 2008, American College of Rheumatology
Heparan Sulfate–Dependent ERK Activation Contributes to
the Overexpression of Fibrotic Proteins and
Enhanced Contraction by Scleroderma Fibroblasts
Yunliang Chen,1 Andrew Leask,2 David J. Abraham,3 Daphne Pala,2 Xu Shiwen,3
Korsa Khan,3 Shangxi Liu,2 David E. Carter,4 Sarah Wilcox-Adelman,5
Paul Goetinck,5 Christopher P. Denton,3 Carol M. Black,3 Andrew A. Pitsillides,6
Catherine E. Sarraf,7 and Mark Eastwood7
itor ␤-xyloside, and soluble heparin on the overexpression of profibrotic genes were compared in fibroblasts
from lesional skin of patients with diffuse SSc and
fibroblasts from healthy control subjects. Identified
protein expressions were compared with the contractile
abilities of fibroblasts while they resided within a collagen lattice. Forces generated were measured using a
culture force monitor.
Results. Inhibiting MEK/ERK with U0126 significantly reduced expression of a cohort of proadhesive
and procontractile proteins that normally are overexpressed by scleroderma fibroblasts, including integrin
␣4 and integrin ␤1. Antagonizing heparan sulfate side
chain formation with ␤-xyloside or the addition of
soluble heparin prevented ERK activation, in addition
to reducing the expression of these proadhesive/
contractile proteins. Treatment with either U0126,
␤-xyloside, or heparin resulted in a reduction in the
overall peak contractile force generated by dermal fibroblasts. Blocking platelet-derived growth factor receptor
with Gleevec (imatinib mesylate) reduced overall contractile ability and the elevated syndecan 4 expression
and ERK activation in SSc fibroblasts.
Conclusion. The results of this study suggest that
heparan sulfate–dependent ERK activation contributes
to the enhanced contractile ability demonstrated by
dermal fibroblasts from lesional skin of patients with
scleroderma. These results are consistent with the notion that the MEK/ERK procontractile pathway is dysregulated in scleroderma dermal fibroblasts. Additionally, the results suggest that antagonizing the MEK/
ERK pathway is likely to modulate heparan sulfate
proteoglycan activity, which in turn may have a profound effect on the fibrotic response in SSc.
Objective. To investigate the contribution of heparan sulfate proteoglycan and Ras/MEK/ERK to the
overexpression of profibrotic proteins and the enhanced
contractile ability of dermal fibroblasts from patients
with systemic sclerosis (SSc; scleroderma).
Methods. The effects of the MEK/ERK inhibitor
U0126, the heparan sulfate side chain formation inhibSupported by the Scleroderma Foundation, the Scleroderma
Society, the Arthritis Research Campaign, the Raynaud’s and Scleroderma Association Trust, the British Heart Foundation, the Welton
Foundation, the Engineering and Physical Sciences Research Council,
the Canadian Institutes of Health Research, the Canada Foundation
for Innovation, and Gap B funds from the University of Western
Ontario. Dr. Leask is recipient of a New Investigator Award from the
Arthritis Society and the Scleroderma Society of Ontario and an Early
Researcher Award from the Ontario Ministry of Research and Innovation. Dr. Pala is recipient of an Ontario Government Scholarship in
Science and Technology.
Yunliang Chen, PhD: University College London and University of Westminster, London, UK; 2Andrew Leask, PhD, Daphne
Pala, BSc, Shangxi Liu, PhD: Schulich School of Dentistry, University
of Western Ontario, London, Ontario, Canada; 3David J. Abraham,
PhD, Xu Shiwen, PhD, Korsa Khan, BSc, Christopher P. Denton,
PhD, FRCP, Carol M. Black, MD, FRCP, DBE: University College
London, London, UK; 4David E. Carter, MSc: London Regional
Genomics Centre, London, Ontario, Canada; 5Sarah Wilcox-Adelman,
PhD, Paul Goetinck, PhD: Massachusetts General Hospital and
Harvard Medical School, Boston, Massachusetts; 6Andrew A. Pitsillides, PhD: Royal Veterinary Collage, London, UK; 7Catherine E.
Sarraf, PhD, FRCP, Mark Eastwood, PhD: University of Westminster,
London, UK.
Drs. Chen, Leask, Abraham, and Pala contributed equally to
this work.
Dr. Denton has received consulting fees, speaking fees,
and/or honoraria (less than $10,000 each) from Actelion and Encysive.
Dr. Black has received consulting fees, speaking fees, and/or honoraria
(less than $10,000) from Actelion.
Address correspondence and reprint requests to Mark Eastwood, PhD, School of Biosciences, University of Westminster, 115
New Cavendish Street, London W1W 6UW, UK. E-mail: eastwood@
Submitted for publication September 29, 2006; accepted in
revised form September 29, 2007.
Fibrosis is characterized by excessive deposition
and contraction of extracellular matrix (ECM) (1). Fibrosis can range in severity from a disfiguring scar to
fibrotic diseases such as liver cirrhosis, diabetic nephropathy, or systemic sclerosis (SSc; scleroderma), in which
organ dysfunction caused by excess deposition of scar
tissue can result in death due to systemic organ failure.
There is no effective therapy for fibrotic disease, in part
because the molecular basis for fibrosis is unclear.
SSc, a chronic disease of unknown etiology, is
characterized by microvascular injury, autoimmune inflammatory responses, and progressive fibrosis (2,3).
Clinically, SSc can range from limited skin sclerosis with
minimal organ involvement (limited cutaneous SSc) to
extensive fibrosis of the skin and internal organs (diffuse
cutaneous SSc [dcSSc]) (2,3). Mortality in SSc is high
and is directly related to the extent of scarring (2,3). SSc
dermal fibroblasts can be isolated and cultured readily
and will retain their enhanced expression of type I
collagen (4–6). Thus, examination of the molecular
difference that may exist between normal fibroblasts
from healthy individuals and fibroblasts from lesional
areas of patients with SSc would seem to be an ideal
system to yield valuable insights into the molecular
nature of scar tissue formation and progression in
chronic fibrotic disease in general.
An activated form of the fibroblast, termed the
myofibroblast, contributes to the increased synthesis and
contraction of ECM that are characteristic of fibrotic
disease (7,8). These fibroblasts retain their fibrotic,
myofibroblast phenotype for several passages in cell
culture and express a cohort of proadhesive and procontractile proteins (9). In a previous investigation, our
group observed that dermal fibroblasts from lesional
areas of patients with SSc show an enhanced ability to
adhere to and contract their ECM compared with nonlesional SSc and normal dermal fibroblasts. Lesional SSc
fibroblasts also show elevated constitutive ERK activation and overexpress a group of profibrotic genes, including connective tissue growth factor (CCN2) and the
heparan sulfate proteoglycans (HSPGs) syndecan 2 and
syndecan 4 (10). Syndecan 4 is required both for basal
and growth factor–induced ERK activation in normal
fibroblasts and for the enhanced activation of ERK
observed in lesional SSc fibroblasts (10). Overall, the
phenotype of lesional SSc fibroblasts appears to be a
result of a heightened activation of signals operative in
normal fibroblasts.
PGs and their polysaccharide chains, glycosaminoglycans, are components of the ECM and pericellular
matrix adjacent to the external surface of cell membranes. HSPGs are a subclass of PGs composed of a core
protein with covalently attached HS side chains. Although the protein part can determine the localization
of the PG on the cell surfaces or in the ECM, the
glycosaminoglycan component, HS, can mediate interactions with a variety of extracellular ligands such as
growth factors and adhesion molecules. Through these
interactions, HSPGs participate in many events during
cell adhesion, migration, proliferation, and differentiation. In addition, the extracellular domains of these PGs
can be shed from the cell surface, generating soluble
HSPG that can antagonize interactions at the cell surface (11,12).
Although the molecular mechanism underlying
the overexpression of profibrotic proteins in SSc is
unclear, we previously showed that the Ras/MEK/ERK
classic MAPK cascade is important for the induction of
CCN2 expression in normal mesenchymal cells (13–16).
Additionally, we have shown that the synthetic prostacyclin, iloprost, which has antifibrotic capabilities (15),
acts at least in part through the protein kinase
A–dependent inhibition of MEK/ERK signaling (13).
However, the extent to which MEK/ERK signaling
contributes to the overexpression of profibrotic proteins
in scleroderma fibroblasts is unclear.
Consequently, to begin to understand the molecular nature of scar formation in progressive fibrotic
disease, we first investigated whether the HSPG syndecan 4 contributes to the expression of profibrotic genes
in normal fibroblasts and whether this is dependent on
the activation of ERK via a MEK-dependent mechanism. Because we previously showed that the enhanced
constitutive ERK activation in lesional SSc fibroblasts is
attributable to an increase in syndecan 4 expression, we
then investigated the contribution made by the MEK/
ERK pathway and HSPGs to the overexpression of
profibrotic proteins and enhanced contractile forces in
SSc dermal fibroblasts. Our results yield new insights
into the molecular basis of ECM contraction by normal
and lesional SSc fibroblasts and suggest new methods of
combating chronic, pathologic fibrosis.
Cell culture. Briefly, cell culture was performed as
previously described (5,6). Dermal fibroblasts were isolated
from biopsy samples of affected (lesional) areas of 4 patients
with dcSSc (disease duration 12–18 months). Age-, sex, and
anatomic site–matched control biopsy specimens were obtained from 4 healthy volunteers. Informed consent was obtained from all study subjects, and ethics approval was also
obtained. All of the study subjects were women. All patients
fulfilled the American College of Rheumatology (formerly, the
American Rheumatism Association) criteria for the diagnosis
of dcSSc (17) as well as the criteria for dcSSc described by
LeRoy et al (18). None of the subjects were receiving immunosuppressive medication or corticosteroids at the time of biopsy.
The fibroblasts were maintained in Dulbecco’s modified Eagle’s medium (DMEM) (FirstLink, Birmingham, UK)
supplemented with 10% fetal bovine serum (FirstLink), 100
units/ml penicillin, and 100 mg/ml streptomycin, and were
cultured in a humidified atmosphere of 5% CO2 in air. The
fibroblasts were subcultured at a ratio of 1:4 until confluency
was reached. When appropriate, the ERK inhibitor U0126 (10
␮M) (Promega, Southampton, UK), soluble heparin (50 ␮g/
ml) (Sigma, Irvine, UK), the platelet-derived growth factor
(PDGF) receptor inhibitor imatinib mesylate (Gleevec) (2
mM) (Novartis, Surrey, UK), or the activin receptor–like
kinase 5 (ALK-5) inhibitor SB-431542 (10 ␮M) (Tocris, Ellisville, MO) was added for the durations indicated. Dermal
fibroblasts from syndecan 4⫹/⫹ and syndecan 4⫺/⫺ mice were
isolated and cultured as previously described (10).
Western blot analysis. Cells were cultured until confluence in DMEM supplemented with 10% fetal bovine serum
(FBS). Cell layers were harvested using 2% sodium dodecyl
sulfate (SDS). Proteins were quantified according to a Bradford kit (Bio-Rad, Hercules, CA), and equal amounts of
protein (25 ␮g) were subjected to SDS–polyacrylamide gel
electrophoresis using 4–12% polyacrylamide gels (Invitrogen,
Paisley, UK). Gels were blotted onto nitrocellulose, and proteins were detected using anti-CCN2 (Santa Cruz Biotechnology, Heidelberg, Germany), antimoesin, antipaxillin, antivinculin, antiezrin (Cell Signaling Technology, Beverly, MA),
anti–␣-smooth muscle actin (anti–␣-SMA) (Sigma), anti–␣4
integrin and anti–␤1 integrin (Zymed, Burlingame, CA), appropriate horseradish peroxidase (HRP)–conjugated secondary antibodies (Zymed), and an enhanced chemiluminescence
kit (Amersham, Little Chalfont, UK) (19).
Immunohistochemical analysis. Immunohistochemistry was performed using a Vectastain ABC kit (Vector,
Peterborough, UK). Serial frozen sections (6 ␮m) were cut on
a cryostat, air-dried, and then stored at ⫺80°C before being
used. Sections were fixed in ice-cold acetone and then blocked
with normal serum from appropriate species and incubated
with primary antibodies (anti–p-ERK) (Cell Signaling Technology, Hertfordshire, UK) and anti–syndecan 4 (Santa Cruz
Biotechnology) for 1 hour at room temperature. Endogenous
peroxidase was exhausted by incubation with 3% hydrogen
peroxidase at room temperature for 15 minutes in the dark.
After washing, sections were incubated with the appropriate
biotinylated secondary antibody for 30 minutes, rinsed, and
finally incubated with avidin-biotinylated HRP complex for 30
minutes. The antibody-bound peroxidase activity was subsequently disclosed by reaction of the tissue sections with
3-amino-9-ethyl-carbazole and then counterstained with hematoxylin and aqueously mounted with Crystal/Mount (Biomeda,
Foster City, CA). Cells were viewed and photographed using a
Zeiss Axiphot microscope (Zeiss, Welwyn Garden City, UK).
Images were saved as TIF files, and the intensity of antibody
staining was quantified by using the box tool of Northern
Eclipse software (Empix Imaging Institute, Mississauga, Ontario, Canada) to measure the intensity of a uniform square
region (3 independent regions per image).
PG and HS biosynthesis inhibition. For experiments
related to the inhibition of PG biosynthesis, cells were cultured
for 4 days in DMEM/10% FBS with 1 mM ␤-D-xyloside
(4-methylumbelliferyl-␤-D-xyloside) (Sigma), which blocks HS
side chain formation and HSPG biosynthesis, or, as control,
with ␣-xyloside, a related analog that does not appreciably
impact HS side chain and HSPG biosynthesis (20–22). The
cells were then washed twice in phosphate buffered saline prior
to further experimentation. Inhibitor toxicity was assessed
using a cell viability assay (MTT) as described by the manufacturer (Roche, Lewes, UK).
RNA quality assessment, probe preparation, and gene
chip hybridization and analysis. Microarrays and analysis were
performed essentially as previously described (23). All gene
chips were processed at the London Regional Genomics
Centre (Robarts Research Institute, London, Ontario, Canada; online at RNA was harvested in
TRIzol (Invitrogen, Paisley, UK) and quantified, and quality
was assessed using the Agilent 2100 Bioanalyzer (Agilent, Palo
Alto, CA) and the RNA 6000 Nano Labchip kit (Caliper Life
Sciences, Mountain View, CA). Quality data were then analyzed using the Degradometer (online at
(mean ⫾ SD degradation factor 1.99 ⫾ 0.0678). Biotinylated
complimentary RNA (cRNA) was prepared from 10 ␮g of
total RNA, according to the Affymetrix GeneChip Expression
Analysis Technical Manual (Affymetrix, Santa Clara, CA).
Double-stranded cDNA was synthesized using SuperScript II
(Invitrogen) and oligo(dT)24 primers. Biotin-labeled cRNA
was prepared by cDNA in vitro transcription using the Bizarre
High-Yield RNA Transcript Labeling kit (Enzo Brioche, New
York, NY), incorporating biotinylated uridine triphosphate
and cytidine triphosphate. Fifteen micrograms of labeled
cRNA was hybridized to Mouse Genome 430 2.0 gene chips
for 16 hours at 45°C as described in the Affymetrix GeneChip
Expression Analysis Technical Manual.
Gene chips were stained with streptavidin–
phycoerythrin, followed by an antibody solution and a second
streptavidin–phycoerythrin solution, with all liquid handling
performed with a GeneChip Fluidics Station 450 (Affymetric).
Gene chips were scanned with the Affymetrix GeneChip
Scanner 3000 (Affymetrix). Signal intensities for genes were
generated with GCOS1.2 software (Affymetrix), using default
values for the statistical expression algorithm parameters and a
target signal of 150 for all probe sets and a normalization value
of 1. Normalization was performed in GeneSpring 7.2 (Agilent). The RMA preprocessor was used to import data from
the .cel files. Data were first transformed (with measurements
less than 0.01 set to 0.01) and then normalized per chip to the
50th percentile and per gene to wild-type control samples.
Experiments were performed twice, and fold changes were
identified using the GeneSpring filter. Functional groupings
were performed with GeneSpring using Gene Ontology (GO)
criteria (part of the GO biologic process system, in which P
values less than 0.005 are considered significant and had to
have at least 10 members/functional groupings to be considered relevant). Data presented in Table 1 (available online at are an average of these independent studies.
Real-time polymerase chain reaction (PCR). Cells
were serum-starved for 24 hours and treated with heparin (50
␮g/ml) overnight. Total RNA was isolated using TRIzol (Invitrogen), and the integrity of the RNA was verified by gel
electrophoresis or with the Agilent bioanalyzer. Heparin was
removed by digestion with heparinase I (0.3 units/reaction)
(Sigma) for 1 hour at room temperature in 1⫻ One-Step
Master Mix (Applied Biosystems), prior to the addition of
primers. Total RNA (25 ng) was reverse transcribed and
amplified using TaqMan Assays-on-Demand (Applied Biosystems, Warrington, UK) in a 15-␮l reaction volume containing
2 unlabeled primers and 6-FAM–labeled TaqMan MGB
probe. Samples were combined with TaqMan One-Step Master Mix. Amplified sequences were detected using the ABI
Prism 7900HT sequence detector (PerkinElmer Cetus, Vaudreuil, Quebec, Canada) according to the manufacturer’s
instructions. Triplicate samples were run and the level of
transcript expression determined following standardization
with control 28S RNA primers as previously described, using
the difference in threshold cycle method (23). Statistical
analysis was performed by Student’s paired t-test.
Fibroblast-populated collagen lattices. Measurement
of contractile force generated within a 3-dimensional (3-D)
tethered floating fibroblast-populated collagen lattice was performed as described previously (24,25). Using 1 ⫻ 106 cells/ml
of collagen gel (FirstLink), we measured the force generated
across the collagen lattice with a culture force monitor (CFM).
This instrument is capable of measuring the minute forces
exerted by cells within a collagen lattice (26) over 24 hours as
fibroblasts attach, spread, migrate, and differentiate into myofibroblasts. Briefly, a rectangular fibroblast-seeded collagen gel
was cast and floated in medium, tethered to 2 flotation bars on
either side of the long edges, and in turn attached to a ground
point at one end and a force transducer at the other. Cellgenerated tensional forces in the collagen gel are detected by
the force transducer and logged into a personal computer.
Graphic readings are produced every 15 seconds, providing a
continuous output of force (dynes, 1 ⫻ 10⫺5N) generated (26).
The cells used in these experiments were passage matched; all
control and inhibition experiments were run in parallel.
Cell migration assay. Twelve-well plates were precoated with type I collagen (40 ␮g/ml) (FirstLink), followed by
blocking of nonspecific binding sites with 1% bovine serum
albumin (BSA; Sigma). Control and antagonist-treated normal
dermal fibroblasts were plated in triplicate in DMEM supplemented with 2% fetal calf serum. Cells adhered to the 12-well
plates and became confluent within 4 hours, after which the
monolayer was scratched with a sterile pipette tip. Following
scratching, the media was changed, either with or without
antagonists. Photomicrographs were obtained immediately after scratching (0 hours) and at 12-hour and 24-hour time
points. All experiments were performed at least 3 times.
Floating collagen gel contraction assay. Experiments
were performed essentially as described previously (10).
Briefly, 24-well tissue culture plates were precoated with BSA.
Normal fibroblasts and fibroblasts from SSc lesions were
treated with platelet-derived growth factor (PDGF; 50 ng/ml)
(R&D Systems, Abingdon, UK) and/or Gleevec (2 nM) for 24
hours. Pretreated fibroblasts were suspended in MCDB medium (Sigma) and mixed with collagen solution (1 part 0.2M
HEPES, pH 8.0, 4 parts collagen [Vitrogen 100; 3 mg/ml]
[CellTrix, Santa Clara, CA], and 5 parts MCDB ⫻ 2), yielding
a final concentration of 80,000 cells/ml and 1.2 mg/ml collagen.
Collagen/cell suspension (1 ml) was added to each well. After
polymerization, gels were detached from the wells by adding 1
ml of MCDB medium with either PDGF and/or Gleevec at the
same concentration as used in the pretreatment. Contraction
of the gel was quantified by loss of gel weight and decrease in
gel diameter over a 24-hour time period.
Requirement of syndecan 4 for expression of
profibrotic messenger RNA (mRNA) and proteins. Previously, we showed that lesional SSc fibroblasts exhibit
increased activation of the MEK/ERK signaling cascade,
which depended on increased expression of the HSPG
syndecan 4 (10). Both syndecan 4 and MEK/ERK were
required for transforming growth factor ␤ (TGF␤),
acting through the TGF␤ receptor type I (ALK-5), to
promote the ability of normal and SSc fibroblasts to
contract a floating collagen gel matrix (10). These results
collectively suggested that the phenotype of SSc fibroblasts results, at least in part, from enhanced activation
of signaling mechanisms operant in normal fibroblasts
(10). Thus, to further probe the basis for the enhanced
adhesive and contractile phenotype of the lesional SSc
fibroblast responsible for the scarring in SSc (10), we
tested the hypothesis that enhanced HSPG (syndecan
4)–dependent ERK activation is responsible for the
overexpression of profibrotic genes in SSc fibroblasts.
Because we had previously shown that the phenotype of the lesional SSc fibroblast results from enhancement of the syndecan 4–dependent signaling
present in normal fibroblasts (10), we first sought to
examine the extent to which syndecan 4 was required for
the expression of profibrotic genes in normal fibroblasts.
To address this issue, we performed genome-wide expression profiling of dermal fibroblasts from syndecan
4⫹/⫹ and syndecan 4⫺/⫺ mice.
We observed 549 mRNA whose expression was
reduced ⬎2-fold in the absence of syndecan 4. Of these,
functional cluster analysis revealed that genes involved
with growth factor activity, cell communication, and
extracellular compartment were overrepresented in
the categories of syndecan 4–dependent mRNA
(Table 1). (Approximately 900 mRNA were increased
⬎2-fold in syndecan 4⫺/⫺ fibroblasts. Cluster analysis
did not reveal that the expression of genes involved with
growth factor activity, cell communication, and extracellular compartment was increased by the loss of syndecan
4; however, the expression of several individual genes
within these categories was increased by the loss of
syndecan 4 [Table 1].) Results were confirmed by Western blotting and real-time PCR analysis and revealed
that expression of the proadhesive/contractile genes
␣-SMA, integrin ␣4 and integrin ␤1, and Fyn and the
profibrotic fibrosis–associated genes gremlin, PLOD2,
lysyl oxidase, interleukin-6, and insulin-like growth factor binding protein 1 was down-regulated in dermal fibroblasts from syndecan 4⫺/⫺ mice (Figure 1; available online
As anticipated, syndecan 4⫺/⫺ fibroblasts also showed
reduced ERK activation, as visualized by Western blot
analysis of cell extracts using an anti–p-ERK antibody
(Figure 1A). MEK inhibition using U0126 blocked the
expression of syndecan 4–dependent proteins in syndecan 4⫹/⫹ fibroblasts (Figure 1B).
To compare the expression levels of syndecan 4
and p-ERK activity between dermal tissue from patients
with SSc and healthy control dermal tissue, we performed immunohistochemical staining on cryostat sections (n ⫽ 4 each from control subjects and patients with
SSc). The expression of syndecan 4 and the activation of
ERK were strongly detected in SSc tissue relative to
healthy tissue (Figure 1C). Images were then analyzed
using Northern Eclipse software to identify the intensity
of the immunolabeling in the cells, and the mean ⫾ SD
intensity was calculated. The difference between normal
and scleroderma samples was statistically significant
(P ⬍ 0.05, by Student’s paired t-test). All of these
findings were consistent with the notion that syndecan
4–dependent ERK activation contributes to the expression of proadhesive/contractile proteins in fibroblasts.
Type I collagen was not detected as being syndecan
4–dependent in our analyses.
Requirement of enhanced ERK activation of SSc
fibroblasts for overexpression of a group of profibrotic
genes. Previous work by our group demonstrated that
fibroblasts from lesional skin of patients with SSc
showed increased activation of MEK/ERK signaling in a
syndecan 4–dependent manner (10). Thus, to extend our
studies on syndecan 4–deficient fibroblasts and to begin
to evaluate the contribution of the MEK/ERK pathway
to the expression of profibrotic proteins by SSc fibroblasts, we subjected protein lysates prepared from cells
that had been treated for 18 hours with the MEK
inhibitor, U0126 (10 ␮M), to Western blot analysis with
anti–␣-SMA, anti-CCN2, anti–integrin ␣4, anti–integrin
␤1, antipaxillin, antiezrin, antivinculin, and antimoesin
antibodies. We observed that antagonism of MEK
blocked expression of all of these proteins, with the
exception of vinculin and moesin (Figure 2A; available online at
Rheumatism/). These results suggested that ERK activation promoted expression of a cohort of profibrotic
proteins, but the overexpression of vinculin and moesin
by SSc fibroblasts occurred in a MEK/ERK-independent
manner. It is important to note that U0126, the MEK
inhibitor, also reduced expression of profibrotic proteins
in normal fibroblasts, suggesting that the MEK/ERK
pathway normally mediates expression of a cohort of
profibrotic proteins, and that pathways leading to ERK
activation are hyperinduced in SSc fibroblasts.
Role of HSPGs in overexpression of a cohort of
profibrotic genes. MEK/ERK signaling is a key component of the proadhesive cascade, being activated upon
cell adhesion and necessary for cell spreading (19).
HSPGs are also key mediators of proadhesive signaling
(19). Our previous investigation showed that blocking
the synthesis of HS side chains suppresses the ability of
fibroblasts from skin lesions of patients with SSc to
adhere to and contract ECM (10).
To assess whether MEK/ERK activation in fibroblasts is downstream of HSPGs and to clarify the effect
of antagonizing HS side chain formation and ERK
inhibition on the phenotype of SSc fibroblasts, we
examined whether uncoupling the synthesis of HS side
chains modifies the overexpression of a cohort of profibrotic proteins in fibroblasts from skin lesions of patients
with SSc. Consequently, we cultured cells for 4 days, in
the presence of ␤-xyloside to prevent HS side chain
synthesis or in the presence of ␣-xyloside as control. We
observed that pretreatment of cells with ␤-xyloside
suppressed ERK activation in fibroblasts (Figure 2B)
and also reduced the expression of an identical cohort of
proteins that we had found were targeted by blockade of
MEK-induced ERK activation with U0126 (Figure 2A).
A standard cell viability assay has been used to show that
under these assay conditions, ␤-xyloside and ␣-xyloside
were not toxic (data not shown). Our results are consistent with the notion that HSPGs mediate MEK/ERK
pathway activation in fibroblasts and are therefore required for the hyperinduction of MEK/ERK signaling in
SSc fibroblasts and for the overexpression of a cohort of
profibrotic proteins in SSc.
Role of soluble heparin in inhibiting ERK activation and overexpression of a cohort of profibrotic
genes. HSPG function can be compromised by treating
cells with heparin, which competes with HSPGs for
binding to ECM (27). To further confirm that HS side
chains are required for the ERK-mediated overexpression of fibrotic genes on SSc fibroblasts, cells were
pretreated with soluble heparin overnight. Cell lysates
were then subjected to Western blot analysis, using the
antibodies described above. We observed that heparin
pretreatment prevented ERK activation in fibroblasts
and also inhibited the expression in fibroblasts of CCN2,
integrin ␣4, integrin ␤1, and paxillin, but not vinculin
and moesin; this was very similar to the protein expression profiles observed in ␤-xyloside–treated cells, with
the exception of ␣-SMA (Figure 2C). To further elucidate whether heparin treatment alters the gene expression profile for these proteins, we performed real-time
PCR to detect the message levels of integrin ␣4 and
integrin ␤1. It transpired that 12-hour exposure to heparin
in culture inhibited the expression of genes previously demonstrated to be syndecan 4/␤-xyloside/ERK-dependent, in
both normal and SSc fibroblasts (Figure 2D).
Role of HSPGs in mediating activation of the
MEK/ERK pathway and in the enhanced contractile
phenotype of SSc dermal fibroblasts. Previously, we
showed that fibroblasts from patients with SSc have an
enhanced ability to contract ECM (10). To assess the
potential contribution of MEK/ERK to the excessive
ECM contraction of fibroblasts from lesional SSc skin,
we treated cells for 18 hours with U0126 prior to
measuring the contractile forces generated by fibroblasts, using the CFM system. Figure 3 (available online
confirms previous findings by showing that SSc fibroblasts generate greater contractile forces than normal
dermal fibroblasts and that the addition of U0126 reduces the contractile force generated by ⬃38% at the
24-hour time point for both cell types (Figure 3A).
In addition to blocking the overexpression of a
cohort of procontractile proteins mirroring those inhibited by antagonizing MEK/ERK in SSc fibroblasts, antagonizing HS side chain formation also reduces ERK
activation. To assess whether HS side chains were necessary for the contractile properties of SSc fibroblasts,
contraction studies were performed with normal and SSc
fibroblasts that had been pretreated for 4 days with
either ␣-xyloside or ␤-xyloside. The effects of treatment
with ␤-xyloside showed the cellular-derived force was
reduced by 30% for SSc dermal fibroblasts and by 47%
for normal dermal fibroblasts at the 24-hour time point
(Figure 3B). To confirm the involvement of HS side
chains in the generation of contractile forces by dermal
fibroblasts, cells were pretreated with soluble heparin
(50 ␮g/ml) for 18 hours prior to inclusion into the CFM.
Figure 3C shows that the addition of heparin resulted in
a reduction in contractile force after 24 hours (by 30%
for SSc-derived fibroblasts and 43% for normal fibroblasts).
Requirement of HSPGs/ERK for fibroblast
spreading and migration. When the forces produced by
fibroblasts over the first 4 hours were examined, U0126
and heparin treatment resulted in significantly decreased contraction compared with that of cells treated
with xyloside (Figure 3A). Upon contact with the ECM,
fibroblasts undergo a multitude of morphologic changes
including attachment, cell process spreading, and the
formation of stress fibers (28–30). Using FPCL, we
showed that these events occur during the early stage of
gel contraction and are attributable to cell attachment,
spreading, and migration on the collagen gel matrix
mediated by focal adhesions/integrins (23). To deter-
mine the effect of these antagonists on cellular morphology and migration, normal dermal fibroblasts were treated
for 24 hours with either U0126, ␤-xyloside, or heparin.
Staining cells with phalloidin to detect total cellular actin
(Figure 4A; available online at
uk/ArthritisandRheumatism/) revealed that fibroblasts
treated with either U0126 or heparin impaired spreading
on collagen, particularly at early time points (up to 4
hours), concomitant with a reduction in stress fibers and
filopodia. ␤-xyloside also impaired the ability of fibroblasts to spread and generate actin stress fibers; however, the kinetics of this impairment were delayed
relative to cells treated with heparin or U0126. These
observations are consistent with the relative abilities of
U0126, xyloside, and heparin to impede the generation
of contractile forces during the first 4 hours of the
contraction assay (Figure 3). Since cell spreading and
attachment are essential for cell motility and ECM
contraction, we compared the abilities of U0126, xyloside,
and heparin to impair cell migration using the wellestablished “scratch wound assay.” As anticipated, U0126,
␤-xyloside, and heparin significantly reduced cell migration
(Figure 4B) relative to control, vehicle-treated fibroblasts.
These results are consistent with the notion that HSPGs/
ERK-mediated cell migration contribute to the capacity
for fibroblasts to contract a 3-D collagen gel.
Contribution of PDGF-induced syndecan 4 overexpression and ERK activation to the enhanced contractile ability by lesional dermal scleroderma fibroblasts. It
has been previously demonstrated that PDGF can markedly increase cellular levels of syndecan 4 core protein in
dermal fibroblasts (31) and stimulate MEK/ERK signaling in SSc fibroblasts (32). To further investigate
whether PDGF-induced syndecan 4 overexpression and
elevated ERK activation contribute to the enhanced
contractile abilities of SSc dermal fibroblasts, fibroblasts
from normal and SSc tissue were individually pretreated
for 24 hours with either PDGF or Gleevec (a PDGF
receptor inhibitor) prior to performing a floating gel
contraction assay. PDGF significantly enhanced the
contractile ability of dermal fibroblasts within 3-D collagen gels, which, as anticipated, was completely blocked
by Gleevec. Gleevec also significantly reduced the basal
contractile ability of otherwise untreated SSc fibroblasts,
but not normal fibroblasts (Figure 5A; available online
Following floating gel contraction, the cell lysates
were assessed by Western blotting to evaluate the protein expression levels in cells within these floating collagen gels. This showed that Gleevec blocked both basal
and PDGF-induced overexpression of syndecan 4 and
also reduced ERK phosphorylation in both SSc and
normal fibroblasts (Figure 5B). Signaling through the
TGF␤ receptor type I (ALK-5) has previously been
shown to contribute to the contractile phenotype of SSc
fibroblasts (10,33,34). Intriguingly, the ALK-5 inhibitor
SB-431542 did not reduce either the elevated ERK
phosphorylation or the elevated syndecan 4 expression
in SSc fibroblasts (Figure 5B). These results are consistent with a syndecan 4–dependent ERK activation,
which depends on an autocrine PDGF signaling mechanism, that contributes to the enhanced contraction in
lesional dermal scleroderma.
Pathologic chronic fibrosis is characterized by
elevated production and contraction of ECM. There is
no effective treatment for pathologic fibrosis. Because
the systemic disease SSc affects many internal organs
and the skin, studying the molecular basis of scleroderma is likely to yield insights into the basis of fibrotic
disease in general. Isolation of fibroblasts from SSc skin
and normal skin is straightforward, and thus the contribution of the SSc fibroblast to the phenotype of fibrosis
can be readily ascertained. Our results suggest that SSc
fibroblasts show elevated ERK activation in a manner
requiring HS side chains, and that this activity is required for the overexpression of profibrotic proteins in
SSc fibroblasts, including CCN2. These results suggest
that the presence of pathologic, clinically defined scarring in chronic fibrotic disease is primarily attributable
to enhanced ECM contraction.
The HS on HSPG may directly promote adhesion
and contraction through the ability to interact with ECM
(35–37). Our recent investigations have shown that
fibroblasts from lesional skin of patients with SSc overexpress the HSPGs syndecan 2 and syndecan 4 by
contributing to the procontractile phenotype of the
lesional SSc fibroblast, because syndecan 4 is required
for TGF-induced p-ERK activation in normal fibroblasts
and the enhanced phosphorylation of ERK in SSc
lesional fibroblasts (19). In this study, antagonizing
MEK/ERK inhibited overexpression of a cohort of
proadhesive and procontractile proteins, including integrin ␣4 and integrin ␤1, which mirrored those inhibited
by antagonizing HS side chain formation and addition of
soluble heparin. Antagonizing Ras/MEK/ERK, soluble
heparin, and HS side chain formation all resulted in a
reduction in the contractile force within dermal fibroblasts. All of these results indicated that HS side chains
(of HSPGs) mediate the induction of MEK/ERK and
underpin the molecular basis of ECM contraction in SSc
dermal fibroblasts.
While this paper was in preparation, another
group of investigators reported that ERK activation was
observed in scleroderma dermal and lung fibroblasts,
and that ERK inhibition reduced collagen production in
both SSc cell types (38). In this study, we extend these
data by showing that ERK activation in SSc fibroblasts is
responsible for the enhanced expression of a different
phenotypic aspect of SSc fibroblasts, namely the overexpression of proadhesive and procontractile proteins,
including integrins, ␣-SMA, and CCN2. Our results
suggest that anti-MEK/ERK strategies might be generally useful in developing antifibrotics and are consistent
with our previous observations that the antifibrotic agent
iloprost acts, at least in part, by antagonizing MEK/ERK
signaling (13–16). Intriguingly, it appears that mechanisms similar to those operating to control profibrotic
protein expression in normal fibroblasts are also operating to control profibrotic protein expression in SSc
fibroblasts, because uncoupling of HS side chain formation or MEK blockade reduced expression of profibrotic
proteins in normal and SSc cells. These results suggest
that the fibrotic phenotype in SSc may occur through an
exaggeration of signaling cascades normally operative in
fibroblasts. For example, heightened responses to profibrotic proteins such as TGF␤ might contribute to fibrosis, perhaps not due to increased ligand or activity per se,
but rather as a result of dysregulated downstream signaling pathways or through the synergistic effects with
other profibrotic proteins (39–43). For example, downstream signaling in response to the profibrotic proteins
CCN2 and endothelin 1 involve Ras/MEK/ERK, and
CCN2 can act via HSPGs, including syndecan 4 (19,44).
In this study, we showed that blockade of the
PDGF receptor with Gleevec, but not blockade of the
endothelin or ALK-5 receptors, reduced hyperinduction
of syndecan 4 expression and ERK activation in SSc
fibroblasts. Whether the overall contribution of HS side
chains on HSPG-induced activation of ERK in fibroblasts is attributable to the ability of HSPGs to modify
adhesive signaling or to their ability to modify the
activity of growth factor signaling awaits further investigation. However, combined with our previous data
showing that antagonism of autocrine TGF␤ and endothelin signaling can alleviate some aspects of the SSc
phenotype (9,10,33,34), our current findings suggest that
PDGF, TGF␤, and endothelin all cooperate in driving
persistent fibrosis through different yet converging
In conclusion, we have shown that fibroblasts
from lesional skin of patients with SSc overexpress a
cohort of profibrotic proteins in an autonomous manner
that depends on endogenous, elevated ERK activation
mediated by HSPGs. HS-dependent ERK activation
contributes to the enhanced contractile ability of lesional dermal scleroderma fibroblasts. Fibroblast
spreading and migration are the primary mediators of
early-stage force generation during collagen gel contraction. Blocking PDGF receptor with Gleevec reduced
overall contractile ability and the elevated syndecan 4
expression and enhanced ERK activation in SSc fibroblasts. Our results suggest that targeting HS formation,
HSPG synthesis, or signaling via the MEK/ERK pathway may yield new methods of preventing scar tissue
formation in chronic fibrotic disease, including SSc.
Dr. Eastwood had full access to all of the data in the study and
takes responsibility for the integrity of the data and the accuracy of the
data analysis.
Study design. Chen, Leask, Abraham, Pala. Shiwen, Denton, Black,
Acquisition of data. Chen, Leask, Pala, Shiwen, Khan, Liu, Carter,
Denton, Sarraf.
Analysis and interpretation of data. Chen, Leask, Abraham, Pala,
Shiwen, Khan, Liu, Carter, Denton, Pitsillides, Sarraf, Eastwood.
Manuscript preparation. Chen, Leask, Abraham, Wilcox-Adelman,
Goetinck, Denton, Black, Eastwood.
Statistical analysis. Chen, Pala, Liu, Pitsillides.
Contraction studies. Eastwood.
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DOI 10.1002/art.23197
Clinical Images: Bilateral scapulothoracic bursitis
The patient, a 40-year-old woman with a rheumatoid arthritis–limited systemic sclerosis overlap syndrome, presented with a 15-day history
of rapidly increasing symmetric swelling in both parascapularis areas (A). Magnetic resonance imaging (B) revealed 2 well-demarcated
dorsal masses (maximum diameter 10 cm) situated in the subscapularis region between the serratus anterior and posterior latissimus dorsi
muscles and the thoracic wall, with characteristic signs of fluid collection. On ultrasonography, they appeared to contain anechoic fluid
with debris and thickened walls. Bursal aspiration yielded yellow fluid containing 2,200 white blood cells/mm3 with a predominance of
mononuclear cells. Culture was negative. Relief from bursitis was immediate following local steroid injection.
Franco Schiavon, MD
Roberto Ragazzi, MD
University of Padua
Padua Hospital
Padua, Italy
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