© 1999 American Heart Association, Inc.
Vascular Biology |
Similarities and Differences in Smooth Muscle 
-Actin Induction by TGF-ß in Smooth Muscle Versus Non–Smooth Muscle Cells
From the Department of Molecular Physiology and Biological Physics, University of Virginia Health Sciences Center, Charlottesville (P.J.A., G.K.O.); the Franz-Volhard Clinic, Charité at the Humboldt University of Berlin, Berlin, Germany (M.B.H.); and the Department of Medicine, University of Cambridge, Addenbrookes Hospital, Cambridge, UK (P.J.A.).
Correspondence to Paul J. Adam or Gary K. Owens, Department of Molecular Physiology and Biological Physics, Box 449, University of Virginia Health Sciences Center, Charlottesville, VA 22908. E-mail gko{at}virginia.edu
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Abstract |
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Abstract—Transforming growth factor-ß (TGF-ß) has been shown to stimulate smooth muscle (SM)
-actin expression in smooth muscle cells (SMCs) and non-SMCs. We
previously demonstrated that the 2 CArG boxes A and B and a novel
TGF-ß control element (TCE) located within the first 125 bp of the SM
-actin promoter were required for TGF-ß inducibility of SM
-actin in SMCs. The aims of the present study were (1) to
determine whether the TCE exhibits SMC specificity or contributes to
TGF-ß induction of SM -actin expression in non-SMCs (ie,
endothelial cells and fibroblasts) and (2) to determine
whether TGF-ß can induce expression of multiple TCE-containing SMC
differentiation marker genes, such as SM22, h1 calponin,
and SM myosin heavy chain (SM MHC) in non-SMCs. Results of transient
transfection assays demonstrated that mutation of CArG A, CArG B, or
the TCE within a 125-bp promoter context completely abolished TGF-ß
inducibility of SM -actin in endothelial cells and
fibroblasts. However, in contrast to observations in SMCs, inclusion of
regions upstream from -155 completely repressed TGF-ß responsiveness
in non-SMCs. Electrophoretic mobility shift assays showed that TGF-ß
enhanced binding of a serum response factor to the CArG elements and
the binding of an as-yet-unidentified factor to the TCE in
endothelial cells and fibroblasts, but to a much lesser
extent compared with SMCs. TGF-ß also stimulated expression of the
SMC differentiation marker SM22 in non-SMCs. However, in contrast to
SMCs, TGF-ß did not induce expression of h1 calponin and
SM MHC in non-SMCs. In summary, these results suggest a conserved role
for CArG A, CArG B, and the TCE in TGF-ß–induced expression of SM
-actin in SMCs and non-SMCs that is modified by a complex interplay
of positive- and negative-acting cis elements in a
cell-specific manner. Furthermore, observations that TGF-ß stimulated
expression of several early but not late differentiation markers in
non-SMCs indicate that TGF-ß alone is not sufficient to induce
transdifferentiation of non-SMCs into SMCs.
Key Words: smooth muscle -actin • transforming growth factor-ß • smooth muscle cells • non–smooth muscle cells
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Introduction |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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The multifunctional growth factor transforming growth factor (TGF)-ß has been implicated in a number of different cellular functions, including cell differentiation.1 2 3 For example, a recent study from our laboratory provided evidence that TGF-ß might act as a positive differentiation factor for smooth muscle cells (SMCs).3 TGF-ß coordinately upregulated several SMC differentiation markers, including SM
-actin, SM myosin
heavy chain (MHC), and h1 calponin. However,
induction of SM -actin by TGF-ß is not restricted to SMCs. A study
by Shah et al4 demonstrated that TGF-ß stimulated
differentiation of neural crest cells into SMCs or SMC-like cells based
on morphological criteria and the induction of SM -actin and
h1 calponin. Of particular interest, Hirschi et
al5 recently demonstrated that coculture of multipotential
10T1/2 cells with endothelial cells (ECs) resulted in
TGF-ß–dependent induction of multiple SMC differentiation markers,
including SM -actin, SM22, SM MHC, and h1
calponin. TGF-ß has also been shown to induce expression of SMC
differentiation marker genes in differentiated non-SMCs. For example,
Arciniegas et al6 reported that TGF-ß induced SM
-actin in adult bovine ECs. Induction of SM -actin was
accompanied by a loss of factor VIII–related antigen and
transdifferentiation into an SMC-like cell. Of interest, a recent study
by DeRuiter et al7 provided in vivo evidence that
embryonic ECs may transdifferentiate into
subendothelial mesenchymal cells that express SM
-actin and become a potential source of SMCs. Whether TGF-ß might
play a role in this process has yet to be determined.
TGF-ß has also been shown to stimulate SM -actin in fibroblasts
derived from a variety of different tissues.8 9 10
Subcutaneous administration of TGF-ß in rats resulted in formation of
granulation tissue that was rich in myofibroblasts expressing SM
-actin.10 Myofibroblasts play a central role in tissue
repair by closing an open wound through contraction and
production of extracellular matrix.1 In a porcine
model of balloon overstretch coronary artery injury,
adventitial fibroblasts coexpressed TGF-ß and SM -actin,
suggesting that TGF-ß might play a role in SM -actin induction in
these cells.11 Despite clear evidence that TGF-ß can
induce SM -actin expression in a variety of non-SMCs, none of the
studies cited above investigated the underlying molecular mechanisms
for this effect. Moreover, with the exception of studies in
multipotential embryonic cells,4 5 previous studies have
not examined whether TGF-ß induces expression of SM differentiation
markers other than SM -actin in non-SMCs, a key question in
assessing its potential role as a transdifferentiation factor.
SM -actin expression has been shown to be governed by a complex
interplay of both positive- and negative-acting cis elements
that vary in different cell types.12 For example, a
125-bp SM -actin promoter construct had high activity in SMCs and
ECs but was inactive in fibroblasts and L6 myotubes, even though the
latter express the endogenous SM -actin gene. The first
125 bp of the SM -actin promoter contain 2 conserved CArG-like
elements that were shown to be required for SM -actin expression in
SMCs but were not required for basal expression of the 125-bp promoter
in ECs.12 Of particular relevance, a study from our
laboratory demonstrated that the first 125 bp of the SM -actin
promoter were sufficient to confer TGF-ß responsiveness in cultured
SMCs.3 TGF-ß inducibility of SM -actin was shown to
be dependent on 3 cis elements located within the first 125
bp: 2 highly conserved CArG elements (A and B) and a novel TGF-ß
control element (TCE). Results of electrophoretic mobility shift assays
(EMSAs) demonstrated that TGF-ß markedly increased the binding of a
serum response factor to the CArG elements and the binding of an
as-yet-unidentified factor to the TCE.
The aims of the present study were to address the following
questions: (1) What are the molecular mechanisms whereby TGF-ß
induces expression of SM -actin in non-SMC types such as ECs and
fibroblasts that do not normally express their endogenous
SM -actin gene? (2) Do cis elements shown to be important
for TGF-ß responsiveness of SM -actin in SMCs also confer TGF-ß
responsiveness in non-SMCs, or are other cis elements
involved? (3) Does TGF-ß also induce expression of other SM
differentiation markers in non-SMCs, including SM MHC, SM22, and
h1 calponin, whose promoters are known to contain
conserved TCEs and CArG elements?
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Methods |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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Construction of Promoter–Chloramphenicol Acetyltransferase (CAT) Expression Plasmids
The generation of various truncated SM
-actin promoter/CAT
reporter constructs, including the CArG A and B mutants and mutation of
the TCE-like sequence, has been previously reported.3 12
All promoter-CAT plasmid DNAs used for transfections were prepared by
using an alkaline lysis procedure13 followed by banding on
2 successive ethidium bromide/CsCl gradients. Multiple independent
plasmid preparations were tested for each construct.
Cell Culture, Transient Transfections, and Reporter Gene
Assays
SMCs from rat aorta and bovine aortic ECs (BAECs) were isolated
and cultured as previously described.12 We chose to use
BAECs instead of rat aortic ECs (RAECs) for our experiments for the
following reasons: (1) In a previous study from our
laboratory,12 we demonstrated that the expression pattern
of SM -actin promoter constructs was the same in BAECs and RAECs,
suggesting that the mechanisms and trans factors that
regulate SM -actin transcription in BAECs and RAECs are likely to be
the same. (2) BAECs were easier to transfect and less costly to grow.
Rat 1 fibroblasts were a gift of Dr Weber (University of Virginia,
Charlottesville) and were cultured identically as SMCs. For transient
transfection assays, SMCs were seeded in 6-well plates at a density of
3x103/cm2, BAECs at a
density of 8x103/cm2, and
fibroblasts at a density of
2x103/cm2. Cells were
grown to confluence in 10% serum-containing medium and then
growth-arrested for 4 days (fibroblasts for 3 days) in serum-free
medium14 before stimulation with TGF-ß (2.5 ng/mL
human TGF-ß1 from R&D Systems) diluted with
vehicle (4 mmol/L HCl, 1 mg/mL BSA). Control cultures were treated
with vehicle only.
Transfection of the CAT–reporter gene constructs was performed in triplicate (4 µg of DNA per well). For SMCs and fibroblasts, transfection reagent DOTAP (Boehringer Mannheim) was used according to the manufacturer's recommendations (6.7 µL/µg of DNA). BAECs were transfected with the use of the transfection reagent Transfectam (Promega) according to the manufacturer's instructions because transfection efficiency in these cells was lower with DOTAP (M.H. and G.O., unpublished observations, 1997). No differences in transfection efficiencies were observed between DOTAP and Transfectam in other cell types. Cells were exposed to the DNA/DOTAP or DNA/Transfectam mixture for 9 to 12 hours. The medium was then replaced with fresh serum-free medium, and TGF-ß (2.5 ng/mL) or vehicle was added. SMCs and BAECs were harvested 72 hours (fibroblasts at 48 hours) later by scraping into ice-cold buffer A (15 mmol/L Tris [pH 8.0], 60 mmol/L KCl, 15 mmol/L NaCl, 2 mmol/L EDTA, 0.15 mmol/L spermine tetrahydrochloride, and 1 mmol/LDTT).15 Cell lysates were prepared by 4 freeze-thaw cycles, followed by a 10-minute heat inactivation at 65°C; 95-µL aliquots of each cell extract were assayed for CAT activity by enzymatic butyrylation of tritiated chloramphenicol (DuPont-NEN).16 CAT activities were normalized as described previously.3 12 Experiments were repeated 2 to 4 times, and relative CAT activity data were expressed as the mean±SE unless otherwise noted.
RNA Isolation, Probe Synthesis, and Northern Blot Analysis
Cultured vascular SMCs, BAECs, and fibroblasts were isolated
from the surface of culture dishes by 2 washes in 1x PBS followed by
incubation in 2 mL of 1x trypsin/EDTA solution for 3 minutes at
37°C. Cells were then washed twice in ice-cold 1x PBS and collected
by centrifugation at 900 rpm. Total cytoplasmic RNA was
isolated by lysis in 150 mmol/L NaCl, 10 mmol/L Tris (pH
7.4), 1 mmol/L MgCl2, and 0.5% (vol/vol)
Nonidet P-40. Nuclei and cell debris were pelleted at 3000 rpm for 5
minutes, and the supernatant was made to 1.5% with SDS before being
extracted twice with citrate-buffered phenol (pH 4.5) and
centrifuged at 3000 rpm. RNA was precipitated overnight in a
1/10 volume of 3 mol/L sodium acetate (pH 4) and 2.5 volumes of
100% ethanol, collected by centrifugation at 14 000
rpm for 15 minutes, and resuspended in an appropriate volume of
RNAase-free water.
Ten micrograms (10 µL) of total cytoplasmic RNA was denatured in 30
µL of denaturation buffer (19 µL formamide, 7 µL formaldehyde,
and 4 µL 10x MOPS), heated at 60°C for 5 minutes, and then placed
on ice. Five microliters of loading buffer was added, and the samples
were electrophoresed at 160 mA in 1.5% agarose gels containing 2.2
mol/L formaldehyde, 20 mmol/L MOPS, and 1 mmol/L EDTA. RNA
was transferred from the gel to a nylon membrane (Hybond N+, Amersham)
by capillary blotting and cross-linked by using 254-nm UV radiation at
1.5 J/cm2.
[-32P]dCTP–radiolabeled cDNA probes were
generated by random hexamer priming and Klenow extension (Prime It,
Stratagene). SM -actin expression was detected with a 512-bp
EcoR1 fragment that encoded amino acids 202 to 374 of human
skeletal -actin cDNA (a gift from Drs Gunning and Kedes, Veterans
Administration Medical Center, Palo Alto, Calif). The SM22 probe was
a 1-kb rat SM22 cDNA (a gift from Dr C.M. Shanahan, University of
Cambridge, UK). h1 Calponin expression was
identified with a cDNA probe corresponding to the coding region
spanning nucleotides 144 to 715 (a gift from Dr M.
Parmacek, University of Chicago, Chicago, Ill). Probes were hybridized
to blots overnight at 65°C and then washed 3 times in 0.1x SSC/0.1%
SDS before being exposed to x-ray film (Kodak AR) for 4 to 24 hours.
Standardization of RNA loading and transfer was achieved by reprobing
the Northern blots with a 5.8-kb EcoR1 cDNA fragment for 18S
rRNA.
RNase Protection Analysis
A 380-bp cRNA fragment of rat SM MHC, containing 80 bp that is
alternatively spliced to give the SM1 and SM2 isoforms, was cloned into
pGEM 4Z (Promega Corp). This was used to synthesize a
[-32P]UTP–labeled single-stranded (ss) RNA
probe with SP6 polymerase by using the MAXIscript in vitro
transcription kit (Ambion). RNase protection analysis of the SM
MHC fragments representing SM1 and SM2 (see Figure 9B
) was conducted on 10 µg of SMCs or fibroblast (with or
without TGF-ß) RNA by using the HybeSpeed RPA kit (Ambion) according
to the manufacturer's instructions.
[-32P]UTP–labeled RNA size markers (RNA
Century, Ambion) were prepared according to the manufacturer's
instructions.
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Western Blot Analysis
Cell lysates were prepared from confluent, growth-arrested BAEC
and fibroblast cultures stimulated with TGF-ß (2.5 ng/mL) or vehicle
for 8 hours. In brief, cells were rinsed with PBS, scraped into 0.6 mL
of ice-cold RIPA buffer (PBS, 1% NP-40, 0.5% sodium deoxycholate, and
0.1% SDS) plus protease inhibitors (10 mg/mL PMSF, 30
µg/mL aprotinin, and 100 mmol/L sodium orthovanadate), and
passed through a 21-gauge needle several times. Cell lysates were then
incubated on ice for 30 minutes and microfuged for 20 minutes at 4°C
(protocol provided by Santa Cruz). Sample loading was normalized to DNA
content as determined with a DNA fluorometer (Hoefer Scientific). Four
hundred nanograms of DNA was loaded per well on a 7.5%
SDS–polyacrylamide gel electrophoresis (PAGE) Mini-Protean gel
(Bio Rad). The proteins were transferred onto a polyvinylidene
difluoride membrane at 100 V for 1.5 hours. Blocking of the
membrane and probing with appropriate antibodies were performed
according to the enhanced chemiluminescence Western blotting protocol
from Amersham Life Science. Affinity-purified rabbit polyclonal serum
response factor (SRF) antibodies (Santa Cruz), raised against a peptide
corresponding to SRF amino acids 486 to 505, were used as primary
antibodies at a concentration of 1 µg/mL.
Preparation of Nuclear Extracts and EMSAs
Crude nuclear extracts were prepared by the method of Dignam et
al17 by using confluent, growth-arrested BAECs and
fibroblasts stimulated with TGF-ß or vehicle for 7 hours. Protein
concentrations were measured by the Bradford assay (Bio Rad). Probes
for EMSAs were obtained by end-labeling 20 µmol/L ss
oligonucleotides with 150 µCi of
[
-32P]ATP (6000 Ci/mmol) and T4
polynucleotide kinase. Labeled ss
oligonucleotides were annealed, and unincorporated
nucleotides were removed by using Nuc Trap Push columns
(Stratagene) as recommended by the manufacturer.
Nucleotides used either as a probe or as cold competitors
have been described previously.3 When TCE binding was
determined, the 20-µL binding reaction contained 
20 000 counts
per minute of labeled probe, 5 µg nuclear extracts in Dignam buffer
D, 20 mmol/L HEPES [pH 7.9], 50 mmol/L KCl, 4 mmol/L
MgCl2, 0.2 mmol/L EDTA, 0.5 mmol/L DTT,
15% glycerol, 2.5% NP-40, 0.5 µg poly(dA-dT) (Sigma), and cold
competitor oligonucleotides where indicated. Different
binding conditions were chosen to optimize SRF binding: The 20-µL
binding reaction contained 20 000 cpm of labeled CArG A or B
oligonucleotides, 5 µg nuclear extracts in Dignam
buffer D, 100 mmol/L KCl, 5 mmol/L HEPES (pH 7.9), 1
mmol/L EDTA, 35 mmol/L Tris (pH 7.5), 1.125 mmol/L DTT, 10%
glycerol, and 0.125 µg of poly(dI-dC) as a nonspecific competitor.
Specific antibodies against SRF (2 µg per reaction, Santa Cruz) were
added where indicated. The radiolabeled DNA was subsequently added to
the binding reaction and incubated for an additional 20 minutes at room
temperature. Protein-DNA complexes were resolved on a 4.5%
polyacrylamide gel (30:1,
acrylamide/bisacrylamide; Bio-Rad) and
electrophoresed at 170 V in 0.5x TBE (45 mmol/L Tris borate,
1 mmol/L EDTA). The gels were then dried and subjected to
autoradiography at -70°C.
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Results |
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TGF-ß Stimulated SM
-Actin mRNA Expression in Rat 1
Fibroblasts and BAECsTo elucidate the molecular mechanisms whereby TGF-ß regulates SM
-actin expression in non-SMCs, we examined the effect of TGF-ß on
SM -actin mRNA expression in fibroblasts and BAECs. Fibroblasts and
ECs were grown to confluence, growth-arrested in serum-free medium, and
stimulated with 2.5 ng/mL TGF-ß or vehicle. Total RNA was extracted
from these cultures at the times indicated and used for Northern blot
analysis to detect SM -actin gene expression (Figure 1A and 1B). Results demonstrated that
TGF-ß markedly increased SM -actin mRNA expression in both
fibroblasts (Figure 1A) and BAECs (Figure 1B). SM
-actin mRNA levels in fibroblasts were markedly enhanced after 4 and
8 hours but returned to control levels 24 hours after TGF-ß
stimulation (Figure 1A). In BAECs, SM -actin mRNA levels were
elevated at 8 and 24 hours (Figure 1B). The different temporal
pattern of SM -actin stimulation in fibroblasts versus BAECs
suggests that different signal transducing pathways may be involved.
For example, the delayed and sustained upregulation of SM -actin
mRNA expression in BAECs after TGF-ß stimulation could be due to
secondary, de novo synthesized factors. TGF-ß was also found to
increase the expression of nonmuscle ß-actin to almost the same
extent as SM -actin in both fibroblasts and BAECs. In contrast,
results of previous studies showed that TGF-ß treatment selectively
increased SM -actin in SMCs.3
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TGF-ß–Induced Stimulation of SM -Actin in Fibroblasts and ECs
Was Dependent on CArG Elements A and B and a TCE Element
Previous studies from our laboratory have shown that
TGF-ß–induced stimulation of SM -actin in SMCs was due at least
in part to increased transcription.3 Transient
transfection analyses demonstrated that the first 125 bp of the
SM -actin promoter were sufficient to confer TGF-ß responsiveness
in SMCs.3 On the basis of site-directed mutagenesis, 3
cis regulatory elements within these 125 bp were identified
to be essential for TGF-ß responsiveness: 2 CArG elements, A (-62)
and B (-122), and a novel TCE.3
To determine whether increases in SM -actin transcription
contributed to TGF-ß–induced increases in SM -actin mRNA
expression in fibroblasts and BAECs, transient transfection assays with
a construct containing the first 125 bp of the SM -actin promoter
linked to a promoterless CAT reporter gene (p125/CAT) were performed.
Results demonstrated that TGF-ß induced an 3-fold increase in
reporter activity in fibroblasts (Figure 2A) and an 6-fold increase in BAECs
(Figure 2B) above vehicle-treated control levels, suggesting
that TGF-ß–induced increases in SM -actin mRNA were due at least
in part to increased SM -actin gene transcription. We then tested
whether cis regulatory elements important for TGF-ß
induction of SM -actin in SMCs were also required for
TGF-ß–mediated stimulation of SM -actin in non-SMCs. A p125/CAT
wild-type reporter construct and constructs containing mutations of
either CArG A or B, alone or in combination, were transiently
transfected in fibroblasts and BAECs. Mutating both CArG's completely
abolished the TGF-ß–induced increases in CAT activity in fibroblasts
(Figure 2A) and BAECs (Figure 2B). The latter observation
is of particular interest, since we previously demonstrated that both
CArG elements were not required for basal SM -actin expression in
BAECs, indicating that both CArG-dependent and -independent mechanisms
contribute to regulation of the 125-bp SM -actin promoter in BAECs.
To determine the importance of the TCE, 3 different previously
characterized mutations of the TCE3 were tested
within the context of a p125/CAT promoter construct for their effects
on SM -actin expression in fibroblasts and BAECs stimulated with
TGF-ß. Results showed that each mutation of the TCE completely
abolished reporter activity in fibroblasts (Figure 2A) compared
with the p125/CAT wild-type construct. Likewise, TCE mutations markedly
reduced CAT activities in BAECs (Figure 2B). TCE mutations
reduced not only TGF-ß–induced increases in reporter activities but
also basal transcriptional activities, suggesting that BAECs and
fibroblasts may produce TGF-ß in an autocrine fashion, similar to
SMCs, under our culture conditions.3 These results
indicate that TGF-ß–induced increases in SM -actin in non-SMCs
require the same cis regulatory elements necessary in SMCs,
at least when tested in the context of the p125 promoter.
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TGF-ß Treatment Increased SRF Binding to the SM -Actin CArG's
in Fibroblasts and BAECs
Because transfection data provided evidence for the functional
importance of the SM -actin CArG elements for TGF-ß inducibility
of SM -actin in fibroblasts and BAECs, we tested whether TGF-ß
affected SRF binding to the SM -actin CArG's. Previous studies had
shown that TGF-ß markedly enhanced SRF binding to CArG A and B in
SMCs.3 EMSAs were performed with labeled 20-bp CArG B
oligonucleotides and nuclear extracts from SMCs,
fibroblasts, and BAECs treated with TGF-ß or vehicle (Figure 3). TGF-ß treatment of fibroblasts and
BAECs increased binding activity to CArG B (lanes 10 and 6) compared
with vehicle-treated controls. However, when compared with SMCs (lane
2), TGF-ß–induced enhancement of binding activity in fibroblasts and
BAECs was much less. To determine whether binding to CArG B was
specific, cold competitor oligonucleotides were added
to the binding reactions. Cold competitor
oligonucleotides completely abolished complex
formation, indicating that binding to CArG B was specific (lanes 4, 8,
and 12). When polyclonal SRF antibodies were added to the binding
reactions, complexes that formed with CArG B were supershifted in all 3
cell types, indicating that SRF was part of the complex. Similar
experiments were performed using CArG A as a probe. Results
demonstrated that TGF-ß markedly increased SRF binding to CArG A in
SMCs and modestly in fibroblasts but did not affect SRF binding in
BAECs (data not shown). The observation that TGF-ß increases SRF
binding to CArG A in SMCs and fibroblasts but not in BAECs suggests
that TGF-ß-dependent mechanisms controlling SRF binding are regulated
in a cell-specific manner. For example, although TGF-ß increases SRF
binding to the relatively weak SRF binding site within CArG
A18 in SMCs and fibroblasts, this mechanism may not be
present or sufficient to increase SRF binding to CArG A in BAECs.
Consistent with this finding, the failure of TGF-ß to
increase SRF protein expression in BAECs could contribute to the
failure of TGF-ß to increase SRF binding to CArG A in BAECs (see
Figure 4B). Furthermore, we have shown previously that basal SM
-actin expression in BAECs is CArG independent, whereas CArG boxes
are absolutely required for SM -actin expression in
SMCs.12 Taken together, these data indicate that
TGF-ß–induced increases in SRF binding to CArG A and B were much
greater in SMCs than non-SMCs and that SRF binding activities are
regulated in a cell-specific manner.
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TGF-ß Increased SRF Protein Expression in SMCs and Fibroblasts
but Not in BAECs
We have previously demonstrated that TGF-ß–induced increases in
SRF binding in SMCs were due at least in part to enhanced SRF
expression.3 To test whether TGF-ß–induced increases in
SRF expression also contributed to enhanced SRF binding in non-SMCs, we
performed Western blot analyses on lysates obtained from
TGF-ß–treated fibroblasts and BAECs (Figure 4
). Results demonstrated that TGF-ß
markedly increased immunoreactive SRF expression in fibroblasts
compared with vehicle-treated controls (Figure 4A). A similar
increase in SRF expression was observed when fibroblasts were
stimulated with 10% FBS. Moreover, both TGF-ß and 10% FBS induced
not only high-molecular-weight (67-kDa) but also
low-molecular-weight (63- to 64-kDa) SRFs compared with vehicle
controls. Misra et al19 demonstrated that mature, highly
phosphorylated SRF has a molecular weight of 67 kDa,
whereas newly synthesized SRF has a lower molecular weight. Thus, our
results suggest that SRF had been synthesized de novo. In contrast,
TGF-ß treatment did not significantly increase SRF protein expression
in BAECs (Figure 4B). In summary, our results indicate that
increased SRF protein expression contributed at least partially to the
enhanced SRF binding in fibroblasts. However, other mechanisms,
including posttranscriptional modification of SRF,19 20
might contribute to the increased SRF binding to CArG B in BAECs.
TGF-ß Increased TCE Binding Activities in Fibroblasts and
BAECs
Previous results from our laboratory demonstrated that TGF-ß
markedly increased binding activity of an as-yet-unidentified factor to
the TCE in SMCs.3 The functional importance of the TCE for
TGF-ß inducibility of SM -actin in non-SMCs prompted us to test
whether TGF-ß also affects TCE binding activities in non-SMCs. EMSAs
were performed with a 19-bp probe containing the TCE and nuclear
extracts from TGF-ß or vehicle-treated fibroblasts and BAECs and
compared with SMCs (Figure 5). Results
showed enhanced binding activities with nuclear extracts derived from
all TGF-ß–treated cell types compared with their respective
controls. Of interest, TGF-ß–induced TCE binding activities were
greatest in SMCs (lane 2), intermediate in fibroblasts (lane 6), and
lowest in BAECs (lane 4). Addition of cold TCE competitor
oligonucleotides (100x molar excess) inhibited shift
band formation (lanes 7 through 9), indicating that the shift band
formed with the TCE probe represented a sequence-specific
protein-DNA complex. No shift band was formed when the mutant TCE m2
oligonucleotide was used as a labeled probe and
incubated with nuclear extracts from TGF-ß–treated cells (lanes 10
through 12). These results clearly indicate that TCE binding activity
is not restricted to SMCs. Moreover, the similarity in mobility of the
TCE shift complex in each cell type suggests that a similar binding
factor (or factors) is expressed in each cell type.
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SM -Actin Promoter Regions Upstream From -125 bp Repressed
TGF-ß Inducibility of SM -Actin in Non–SMCs but Not in
SMCs
There is extensive evidence showing that SM -actin expression
is regulated in a cell type–specific manner.12 21 22 23 For
example, in BAECs, a 125-bp promoter construct had high transcriptional
activity, but inclusion of regions upstream from -125 bp up to -2.8
kb (designated pProm) completely repressed expression of SM -actin
in BAECs.12 Promoter regions upstream from -125 also
reduced, to some extent, the basal transcriptional activity in SMCs but
did not affect TGF-ß responsiveness of SM -actin.3 To
test whether negative regulatory elements affected TGF-ß–induced SM
-actin expression in non-SMCs, we transfected a construct containing
2.8 kb of the SM -actin promoter in fibroblasts and BAECs (Figure 6). Transfection data demonstrated that
TGF-ß induced a 2-fold increase in pProm CAT activity in BAECs, but
no increase in reporter activity was observed in fibroblasts in
response to TGF-ß. By comparison, TGF-ß stimulated an 8-fold
increase in pProm CAT activity in SMCs. Thus, there appeared to be cell
type–specific repressor elements upstream from -125 bp that markedly
inhibited TGF-ß responsiveness of the core p125 promoter in
non-SMCs.
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As a first step to further identify negative regulatory elements within
pProm that repressed TGF-ß inducibility of SM -actin in non-SMCs,
several deletion mutants were tested for their ability to confer
TGF-ß responsiveness. Results of transient transfection assays
demonstrated that both basal and TGF-ß–inducible reporter activity
of a p208/CAT construct was significantly reduced in BAECs (Figure 7A) compared with constructs containing
125 or 155 bp of the promoter. A further reduction in basal and
TGF-ß–inducible reporter activity was observed in constructs
containing 371 or 547 bp of the SM -actin promoter. Previous studies
from our laboratory identified 2 muscle (M)-CAT elements located at
-184 and -320 of the rat SM -actin promoter that bind the
transcription enhancer factor-1 family of transcription
factors.24 Because M-CAT elements have been shown to act
as positive or negative regulatory elements depending on the cell type
studied,24 25 26 we tested the effects of mutations of the
M-CAT elements on the TGF-ß inducibility of SM -actin in BAECs.
Mutation of either M-CAT, alone or in combination, did not restore
TGF-ß inducibility of the p371/CAT construct (data not shown),
suggesting that the M-CAT elements were not responsible for repression
of TGF-ß responsiveness in BAECs.
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Deletion mutants of pProm were also tested in fibroblasts. Results
demonstrated that inclusion of a promoter region between -155 and
-208 completely abolished TGF-ß inducibility of SM -actin
transcription in fibroblasts (Figure 7B). This finding is
consistent with previous reports27 that provided
evidence for a potent, highly conserved repressor element located at
-191 and -224 within the mouse SM -actin promoter that subdued SM
-actin expression in fibroblasts, in that deletion of this region
resulted in transcriptional activation of SM -actin in response to
serum in fibroblasts. The p208/CAT promoter construct tested in the
present studies contained only the first half of this repressor
region, suggesting that this region is sufficient for trans
factor binding and inhibition of TGF-ß responsiveness of SM -actin
in fibroblasts. In the region between -155 and -208 bp, there is also
an M-CAT element.24 However, it seems unlikely that
this element would be responsible for the repressor activity associated
with this region, since the M-CAT element situated at -176 to -181
within the mouse SM -actin promoter is required for high-level
expression of SM -actin in fibroblasts.26 Furthermore,
studies from our laboratory demonstrated that mutation of this element
had no effect on SM -actin transcription in
fibroblasts.24 In summary, results indicate that (1)
negative-acting elements upstream from -155 restrict TGF-ß
responsiveness of the SM -actin gene in non-SMCs but not in SMCs and
that (2) in non-SMCs, TGF-ß inducibility of SM -actin requires
positive-acting cis elements located outside pProm that
presumably act in concert with the CArG elements A and B, as well as
with the TCE.
TGF-ß–Stimulated SM22 but Not h1 Calponin or SM
MHC mRNA Expression in Non-SMCs
Studies of aortic vascular SMCs have demonstrated the ability of
TGF-ß to increase the expression of a number of SM differentiation
marker genes, including SM -actin, SM MHC, and
h1 calponin.3 These results
suggested that TGF-ß might act as a positive differentiation factor
for SMCs. To determine whether TGF-ß enhanced expression of other SM
differentiation marker genes in addition to SM -actin in non-SMCs,
the levels of SM22, h1 calponin, and SM MHC
mRNA expression were measured in fibroblasts and BAECs after TGF-ß
treatment (Figures 8 and 9). Cells were cultured as described in
the legend to Figure 1A and 1B. Northern blot analysis
demonstrated that TGF-ß strongly upregulated SM22 mRNA levels in
fibroblasts 4 and 8 hours after TGF-ß treatment, with expression
returning to vehicle control levels after 24 hours (Figure 8A).
SM22 mRNA expression was also markedly increased in BAECs 8 and 24
hours after the addition of TGF-ß (Figure 8B). Interestingly,
in contrast to SMCs, h1 calponin mRNA expression
was not detected in either control or TGF-ß–treated fibroblasts or
BAECs (fibroblast data are shown in Figure 9A). This result
suggests that TGF-ß may selectively upregulate the expression of
early SMC differentiation marker genes (SM -actin and SM 22) but
not of later-stage marker genes (h1 calponin) in
non-SMCs. In support of this concept, we performed RNase protection
analysis experiments, which showed that TGF-ß also failed to
increase SM MHC expression in fibroblasts (Figure 9B).
|
|
Discussion |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
The goals of the present study were to investigate the molecular mechanisms whereby TGF-ß stimulates SM
-actin expression
in non-SMCs and to determine whether TGF-ß also induces expression of
other SM differentiation markers in non-SMCs, including SM MHC,
SM22, and h1 calponin. Transfection
analyses demonstrated that expression of a 125-bp SM -actin
promoter/CAT construct in BAECs and fibroblasts was similar to that in
SMCs and was dependent on both CArG elements A and B and a previously
described TCE.3 However, in contrast to SMCs, inclusion of
regions upstream from -155 completely restricted SM -actin
expression in non-SMCs. In addition to SM -actin, TGF-ß stimulated
SM22 in non-SMCs but failed to induce late-stage differentiation
markers, including SM MHC and h1 calponin.
Results suggest that TGF-ß is able to upregulate a subset of SM
differentiation marker genes in non-SMCs but not transdifferentiation
of ECs or fibroblasts into SMCs.
Despite the functional dependence of TGF-ß–mediated stimulation of
SM -actin transcription on the 2 CArG elements and the TCE in both
SMCs and non-SMCs, results of the present studies also provide
clear evidence for differences in TGF-ß–induced stimulation of SM
-actin in SMCs versus non-SMCs. For example, although no qualitative
differences were observed in SRF and TCE factor binding in SMCs versus
non-SMCs, the magnitude of changes in TGF-ß–induced increases in SRF
and TCE factor binding was much greater in SMCs than in non-SMCs. There
were also cell type–dependent differences in mechanisms that
contributed to increased SRF binding. For example, TGF-ß enhanced SRF
expression in fibroblasts (Figure 4A) and SMCs3 but
not in ECs (Figure 4B). Moreover, TGF-ß–induced increases in
SRF binding to the CArG elements in BAECs are unlikely to be mediated
by the homeodomain-containing protein, MHox, as shown previously in
SMCs,29 because it is not expressed in
BAECs.30 Although MHox is likely to be expressed in
fibroblasts,30 it appears to lack functional importance
for SM -actin transcription, because mutation of the highly
conserved MHox binding site in the mouse SM -actin promoter did not
affect SM -actin transcription in fibroblasts.21 The
as-yet-unidentified TCE binding factor may also play a role in
regulating TGF-ß effects on SRF binding. TGF-ß–induced increases
in TCE binding activities in SMCs far exceeded those in non-SMCs and
coincided with markedly enhanced SRF binding. This result suggests that
these factors act in a cooperative fashion in SMCs. However, because
neither SRF nor the TCE binding factor is SMC-specific, our studies
suggest that these 2 factors contribute to quantitative rather than
qualitative differences in SM -actin expression in SMCs versus
non-SMCs.
Another striking difference between TGF-ß–mediated upregulation of
SM -actin in SMCs versus non-SMCs was that potent,
negative-regulatory elements located between -155 and -547 bp
restricted SM -actin expression in non-SMCs. Previous studies from
our laboratory have shown that inclusion of the promoter region from
-155 bp to -2.8 kb also reduced basal expression of SM -actin in
SMCs to some degree but to a much lesser extent than in BAECs. This
suggests that negative-regulatory elements associated with this region
may repress expression of SM -actin in non-SMCs under normal
circumstances. However, relatively little is known about potential
negative-acting trans factors that may interact with this
region. Recently, Sun et al26 identified 2 ssDNA
binding proteins in fibroblasts that bound to a highly conserved
repressor element from -165 to -195 bp within the mouse SM -actin
promoter. These ssDNA binding proteins were shown to stabilize a local
ss conformation within the promoter and to preclude binding of the
essential activator protein, transcription enhancer
factor-1, to the M-CAT element (-181 and -176). Consistent
with these outcomes, our results showed that inclusion of the region
from -155 to -208 bp completely abolished basal and TGF-ß–induced
SM -actin transcription in fibroblasts. Whether the ssDNA binding
proteins are also responsible for the marked reduction in activity of
the p208/CAT construct in BAECs needs to be determined. However, in
contrast to non-SMCs, upstream regions of the SM -actin promoter did
not affect TGF-ß responsiveness of SM -actin in SMCs. This finding
suggests that the trans factors that interact with these
negative-regulatory elements are either not present in SMCs or that
TGF-ß downregulates their expression or binding activity.
The presence of negative cis elements within the 2.8-kb SM
-actin promoter that potently repress the relatively high activity
of a shorter p125 SM -actin promoter fragment in non-SMCs
presents a paradox, in that the endogenous SM -actin
gene can be stimulated by TGF-ß in fibroblasts and BAECs in vitro and
in vivo (Figure 1A and 1B).6 10 11 This
observation suggests that there are TGF-ß–dependent mechanisms that
overcome the negative-regulatory effects of the repressor regions. One
possibility is that there are additional positive-acting
TGF-ß–responsive elements outside the 2.8-kb promoter region that
act in concert with the CArG and the TCE, thereby overriding the effect
of the repressor elements located between -155 bp and -2.8 kb.
However, the regions of the SM -actin gene that might be required to
override repressor effects in non-SMCs have not been identified.
Another mechanism whereby the effects of negative-regulatory elements
could be overcome by TGF-ß is through induction of a change in
chromatin structure. Consistent with this possibility, results
of a study by Alevizopoulos et al31 demonstrated that
TGF-ß participates in chromatin-regulated transcriptional control. In
brief, they demonstrated that TGF-ß–induced activation of the
collagen 2(1) promoter is mediated via interaction of the
transcription factor C-terminal fragment-1 with histone H3.
Finally, a third mechanism whereby TGF-ß could increase SM -actin
expression in non-SMCs is through some posttranscriptional control
mechanisms, including stabilization of SM -actin
mRNA.22 32 33 Whether these mechanisms contribute to
TGF-ß induction of SM -actin in fibroblasts and ECs requires
further investigation.
Our observation that TGF-ß–stimulated expression of the early (SM
-actin and SM22) but not late (h1 calponin
and SM MHC) differentiation marker genes in non-SMCs also implies that
there are some fundamental differences in regulation of these SMC
marker genes. Although SM -actin, SM22, h1
calponin, and SM MHC are coordinately expressed at high levels in the
mature SMC, there is a temporal dissociation of expression of these
markers during development.22 SM -actin and SM22 are
the earliest markers of SMC lineage,22 34 35 whereas
h1 calponin and SM MHC are expressed at later
stages during development.22 36 37 In addition, SM
-actin and SM22 display a similar pattern of tissue distribution
in early development. Both markers are expressed in the cardiac,
skeletal, and SMC lineages in the embryo before becoming restricted to
SMCs late in embryogenesis.22 35 In contrast, expression
of SM MHC and h1 calponin is more highly
restricted to SMCs during development. SM MHC is exclusively expressed
in SMC lineages throughout development.37
h1 Calponin transcripts have been found in SMCs
and in the heart tube, but surprisingly, no h1
calponin protein was detected in the heart.38 The
relatively restricted expression of SM MHC and h1
calponin in SMCs suggests that in non-SMCs, it may be regulated by
positive-acting transcription factors unique to SMCs. Alternatively,
expression may be strongly inhibited by repressors or
posttranscriptional mechanisms in non-SMCs.38 Of interest,
Madsen et al39 identified an Sp1 binding site adjacent to
CArG box 2 in the SM MHC promoter that acts as a repressor in SMCs and
that Sp1 levels were increased in myointimal SMCs that express lower
levels of SM MHC.39 Interestingly, the
h1 calponin promoter also contains 2 Sp1 binding
sites within the first -500 bp.28 Whether Sp1 sites of
either SM MHC or h1calponin act as
transcriptional repressor sites in non-SMCs remains to be determined.
Taken together, our results suggest that, although there are a number
of common cis regulatory elements, such as CArG's and the
TCE, that appear to contribute to the expression of each of these
differentiation marker genes in SMCs, there are clearly unique
regulatory elements and mechanisms required for cell type– and
gene-specific regulation.
Results of the present studies provide clear evidence that TGF-ß upregulates only a subset of SMC differentiation marker genes in non-SMCs. In contrast, several recent studies have shown that TGF-ß can stimulate the late-stage differentiation markers h1 calponin and SM MHC in multipotential neural crest cells and 10T1/2 cells.4 5 The failure of TGF-ß to induce the SMC differentiation program in fibroblasts or BAECs may reflect the differential developmental potential of these 2 cell types and/or additional humoral factors present in the neural crest and 10T1/2 cell cultures but not in our BAEC or fibroblast culture systems. Taken together, the current results indicate that TGF-ß cannot by itself stimulate transdifferentiation of somatic cells such as BAECs or fibroblasts into SMCs, although it appears to be sufficient to stimulate differentiation of several multipotential embryonic cell lines to an SMC-like lineage.
In summary, results of the present study provide clear evidence
that TGF-ß–induced upregulation of SM -actin in SMCs and non-SMCs
requires common elements, including CArG's and the TCE. However, these
elements function differentially in SMCs versus non-SMCs through a
complex interplay of positive- and negative-acting cis
elements that act in a cell-specific manner. Furthermore, we found that
TGF-ß was not sufficient to induce transdifferentiation of somatic
cells, including BAECs and fibroblasts, into SMCs. Further studies are
required to identify regulatory mechanisms that drive TGF-ß–induced
expression of SM -actin in non-SMCs and how these override the
potent repressor activity that normally suppresses expression of SM
-actin in non-SMCs.
|
Acknowledgments |
|---|
This study was supported by grants RO1 HL-38854 and PO1 HL-19242 from the National Institutes of Health, Bethesda, Md (to Dr Owens), Fellowship Grant VA-94-F-14 (to Dr Hautmann) from the American Heart Association, Virginia Affiliate, Inc, and by Junior Research Fellowship FS-96072 (to Dr Adam) from the British Heart Foundation. We gratefully acknowledge the expert technical assistance of Diane Raines and Andrea Tanner.
Received September 21, 1998; accepted January 26, 1999.
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A. Masszi, C. Di Ciano, G. Sirokmany, W. T. Arthur, O. D. Rotstein, J. Wang, C. A. G. McCulloch, L. Rosivall, I. Mucsi, and A. Kapus Central role for Rho in TGF-beta 1-induced alpha -smooth muscle actin expression during epithelial-mesenchymal transition Am J Physiol Renal Physiol, May 1, 2003; 284(5): F911 - F924. [Abstract] [Full Text] [PDF]
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R. C. Chambers, P. Leoni, N. Kaminski, G. J. Laurent, and R. A. Heller Global Expression Profiling of Fibroblast Responses to Transforming Growth Factor-{beta}1 Reveals the Induction of Inhibitor of Differentiation-1 and Provides Evidence of Smooth Muscle Cell Phenotypic Switching Am. J. Pathol., February 1, 2003; 162(2): 533 - 546. [Abstract] [Full Text] [PDF]
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I. Manabe, T. Shindo, and R. Nagai Gene Expression in Fibroblasts and Fibrosis: Involvement in Cardiac Hypertrophy Circ. Res., December 13, 2002; 91(12): 1103 - 1113. [Abstract] [Full Text] [PDF]
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A. Zalewski, Y. Shi, and A. G. Johnson Diverse Origin of Intimal Cells: Smooth Muscle Cells, Myofibroblasts, Fibroblasts, and Beyond? Circ. Res., October 18, 2002; 91(8): 652 - 655. [Full Text] [PDF]
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J. G. Cogan, S. V. Subramanian, J. A. Polikandriotis, R. J. Kelm Jr., and A. R. Strauch Vascular Smooth Muscle alpha -Actin Gene Transcription during Myofibroblast Differentiation Requires Sp1/3 Protein Binding Proximal to the MCAT Enhancer J. Biol. Chem., September 20, 2002; 277(39): 36433 - 36442. [Abstract] [Full Text] [PDF]
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M. G. Frid, V. A. Kale, and K. R. Stenmark Mature Vascular Endothelium Can Give Rise to Smooth Muscle Cells via Endothelial-Mesenchymal Transdifferentiation: In Vitro Analysis Circ. Res., June 14, 2002; 90(11): 1189 - 1196. [Abstract] [Full Text] [PDF]
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P. Qiu and L. Li Histone Acetylation and Recruitment of Serum Responsive Factor and CREB-Binding Protein Onto SM22 Promoter During SM22 Gene Expression Circ. Res., May 3, 2002; 90(8): 858 - 865. [Abstract] [Full Text] [PDF]
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J. E. Pitera, V. V. Smith, A. S. Woolf, and P. J. Milla Embryonic Gut Anomalies in a Mouse Model of Retinoic Acid-Induced Caudal Regression Syndrome : Delayed Gut Looping, Rudimentary Cecum, and Anorectal Anomalies Am. J. Pathol., December 1, 2001; 159(6): 2321 - 2329. [Abstract] [Full Text] [PDF]
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A. M. Hoggatt, A. M. Kriegel, A. F. Smith, and B. P. Herring Hepatocyte Nuclear Factor-3 Homologue 1 (HFH-1) Represses Transcription of Smooth Muscle-specific Genes J. Biol. Chem., September 29, 2000; 275(40): 31162 - 31170. [Abstract] [Full Text] [PDF]
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P. J. Adam, C. P. Regan, M. B. Hautmann, and G. K. Owens Positive- and Negative-acting Kruppel-like Transcription Factors Bind a Transforming Growth Factor beta Control Element Required for Expression of the Smooth Muscle Cell Differentiation Marker SM22alpha in Vivo J. Biol. Chem., November 22, 2000; 275(48): 37798 - 37806. [Abstract] [Full Text] [PDF]
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P. Qiu and L. Li Histone Acetylation and Recruitment of Serum Responsive Factor and CREB-Binding Protein Onto SM22 Promoter During SM22 Gene Expression Circ. Res., May 3, 2002; 90(8): 858 - 865. [Abstract] [Full Text] [PDF]
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