The SRF Target Gene Fhl2 Antagonizes RhoA/MALDependent Activation of SRF
The MIT Faculty has made this article openly available. Please share
how this access benefits you. Your story matters.
Citation
Philippar, Ulrike, Gerhard Schratt, Christoph Dieterich, Judith M.
Müller, Petra Galgóczy, Felix B. Engel, Mark T. Keating, et al.
“The SRF Target Gene Fhl2 Antagonizes RhoA/MAL-Dependent
Activation of SRF.” Molecular Cell 16, no. 6 (December 2004):
867-880. Copyright © 2004 Cell Press
As Published
http://dx.doi.org/10.1016/j.molcel.2004.11.039
Publisher
Elsevier
Version
Final published version
Accessed
Wed May 25 19:02:24 EDT 2016
Citable Link
http://hdl.handle.net/1721.1/83480
Terms of Use
Article is made available in accordance with the publisher's policy
and may be subject to US copyright law. Please refer to the
publisher's site for terms of use.
Detailed Terms
Molecular Cell, Vol. 16, 867–880, December 22, 2004, Copyright ©2004 by Cell Press
The SRF Target Gene Fhl2 Antagonizes
RhoA/MAL-Dependent Activation of SRF
Ulrike Philippar,1,5,6 Gerhard Schratt,1,6,7
Christoph Dieterich,2 Judith M. Müller,3
Petra Galgóczy,3 Felix B. Engel,4 Mark T. Keating,4
Frank Gertler,5 Roland Schüle,3 Martin Vingron,2
and Alfred Nordheim1,*
1
Abt. Molekularbiologie
Institut für Zellbiologie
Eberhard-Karls-Universität Tübingen
72076 Tübingen
Germany
2
Max-Planck-Institut für Molekulare Genetik
Berlin
Germany
3
Universitäts-Frauenklinik
Zentrale Klinische Forschung
Klinikum der Universität Freiburg
79106 Freiburg
Germany
4
Howard Hughes Medical Institute
Department of Cell Biology
Harvard Medical School
Children’s Hospital
Boston, Massachusetts 02115
5
Department of Biology
Massachusetts Institute of Technology
Cambridge, Massachusetts 02139
Summary
RhoA signaling regulates the activity of the transcription factor SRF (serum response factor) during muscle
differentiation. How RhoA signaling is integrated at
SRF target promoters to achieve muscle-lineage-specific expression is largely unknown. Using large-scale
expression profiling combined with bioinformatic and
biochemical approaches, we identified several SRF
target genes, including Fhl2, encoding a transcriptional cofactor that is highly expressed in the heart.
SRF binds the Fhl2 promoter in vivo and regulates
Fhl2 expression in response to RhoA activation. FHL2
protein and SRF interact physically, and FHL2 binds
the promoters of SRF-responsive smooth muscle (SM)
genes, but not the promoters of immediate-early
genes (IEGs), in response to RhoA. FHL2 antagonizes
induction of SM genes, but not IEGs or cardiac genes,
by competing with the coactivator MAL/MRTF-A for
SRF binding. Our findings identify an autoregulatory
mechanism to selectively regulate subsets of RhoAactivated SRF target genes.
Introduction
Serum response factor (SRF), a MADS-box transcription
factor, regulates the expression of immediate-early
*Correspondence: alfred.nordheim@uni-tuebingen.de
6
These authors contributed equally to this work.
7
Present address: Division of Neuroscience, Children’s Hospital and
Department of Neurobiology, Harvard Medical School, Boston, Massachusetts 02115.
genes (IEGs), cytoskeletal, and muscle-specific genes
(Miano, 2003; Treisman, 1995). Mouse embryos lacking
SRF die before gastrulation and do not form any detectable mesoderm (Arsenian et al., 1998). SRF mediates
transcriptional activation by binding to CArG box sequences in target gene promoters and by recruiting various cofactors. At IEG promoters, SRF associates with
ternary complex factors (TCFs), which bind to Ets binding sites adjacent to the CArG box, to regulate transcription downstream of MAPK signaling (Buchwalter et
al., 2004).
Recently, SRF cofactors have been described that
regulate different subsets of muscle-specific SRF target
genes. Myocardin is predominantly expressed in developing cardiac and smooth muscle and activates the
promoters of cardiac as well as smooth muscle (SM)specific SRF target genes (Wang et al., 2001). In addition,
other members of the myocardin family, MRTF-A and -B,
are expressed in muscle (Wang et al., 2002). MRTF-A/
MAL represents a potent coactivator of muscle-specific
SRF target genes. MAL translocates to the nucleus upon
RhoA-induced actin polymerization, thereby linking Rho
signaling to SRF activation (Miralles et al., 2003). The
cysteine-rich LIM-only proteins CRP1 and CRP2 serve
as bridging molecules between SRF and GATA proteins
and selectively induce the activation of SM promoters
(Chang et al., 2003). In addition to coactivators, corepressors of SRF-dependent transcription have been
identified. GATA transcription factors suppress the transcriptional activity of myocardin on a subset of SRF
target promoters (Oh et al., 2004). The cardiac-specific
homeodomain-only protein (HOP) associates with SRF
and interferes with SRF DNA binding (Chen et al., 2002).
A recent study identified the TCF Elk-1 as an antagonist
of myocardin and a repressor of SRF-mediated SM gene
transcription in response to MAPK signaling (Wang et
al., 2004).
RhoA signaling activates SRF independently of TCFs,
thereby mediating activation of muscle-specific SRF target genes (Hill et al., 1995) and promoting myogenesis
(Sordella et al., 2003).
The LIM-only protein FHL2/DRAL/Slim3 (four-and-ahalf LIM-domain protein 2) is a member of the FHL protein family and functions as a transcriptional modulator.
FHL1–3 are specifically expressed in different muscle
types, whereas FHL4 and ACT (activator of CREM in
testis) are exclusively found in testis (Fimia et al., 1999).
FHL2 is highly expressed in early cardiac precursor cells
and in the heart of adult mice (Chan et al., 1998). FHL1
is expressed in cardiac and skeletal muscle (Fimia et al.,
2000). Although FHL2-deficient mice maintain normal
cardiac function, they exhibit cardiac hypertrophy in
response to ␤-adrenergic stimulation (Kong et al., 2001).
FHL2 has been shown to act as a transcriptional coactivator of several transcription factors including androgen
receptor (Müller et al., 2000), CREB (Fimia et al., 2000),
and AP-1 (Morlon and Sassone-Corsi, 2003). In addition,
FHL2 negatively regulates MAPK signaling in cardiomyocytes (Purcell et al., 2004) and is an antagonist of
the promyelocytic leukemia zinc finger protein (PLZF)
Molecular Cell
868
(McLoughlin et al., 2002). FHL2 localizes to focal adhesions and translocates to the nucleus upon RhoA stimulation, thereby linking extracellular signals to gene control (Müller et al., 2002; Wixler et al., 2000).
Here we present a microarray approach to identify
new SRF target genes using overexpression of SRFVP16 in SRF-deficient embryonic stem (ES) cells, which
led to the identification of SRF target genes implicated
in cytoskeletal organization, apoptosis, wound healing,
and muscle differentiation. We identified Fhl2 as an SRF
target gene. FHL2 is upregulated in an SRF-dependent
manner during ES cell differentiation and in response
to RhoA activation. SRF and FHL2 interact in vitro and
in vivo and bind to the promoters of a subset of SRF
target genes following RhoA activation. FHL2 inhibits
SRF-mediated transcription and antagonizes the RhoA/
MAL-induced activation of SM promoters by competing
with MAL for SRF binding. We propose that RhoA, MAL,
SRF, and FHL2 constitute an autoregulatory feedback
mechanism regulating the expression of subsets of SRF
target genes during myogenesis.
Results
Expression Profiling to Identify SRF-Regulated
Genes in ES Cells
Despite increasing evidence for important functions of
SRF in various biological processes, further insight is
hampered by incomplete knowledge of SRF target
genes. We used a microarray approach to monitor SRFdependent gene expression at the whole-genome level.
We took advantage of the Srf⫺/⫺ ES cell system, which
allows robust induction of SRF target genes independent of signal transduction upon overexpression of the
constitutively active SRF fusion protein SRF-VP16. SRF⌬M-VP16, a mutant defective in DNA binding, served as
control (Schratt et al., 2002). To monitor gene expression
profiles of cells transfected with SRF-VP16 or SRF-⌬MVP16, mRNA from two independent transfections was
hybridized to Affymetrix microarrays (see Supplemental
Table S1 at http://www.molecule.org/cgi/content/full/
16/6/867/DC1/). Two independent Srf⫺/⫺ ES cell lines,
81 Srf⫺/⫺ and 100 Srf⫺/⫺, were used to control for cellbased variations (Weinhold et al., 2000). We considered
all genes regulated by SRF whose expression was at
least 3-fold higher in each of the samples derived from
SRF-VP16-transfected cells, compared to SRF-⌬MVP16-transfected cells. Expression of a set of 86 genes
was reproducibly induced by SRF-VP16, and Table 1
provides their functional clustering.
Several genes shown in Table 1 were reported to function in different muscle lineages, others play roles in
cell cycle/apoptosis, cytoskeletal organization, wound
healing, or cellular metabolism. Our analysis recovered
17 previously known SRF target genes, including Egr-1,
SMA (smooth muscle actin), and vinculin. However, regulation by SRF had not been demonstrated previously
for several other genes, e.g., Fhl2, tuftelin, CTGF (chondrocyte tissue growth factor), keratin 17 (KRT-17), and
endothelin-1 (ET-1). Taken together, we confirmed SRFregulation for 17 known SRF targets representing about
50% of all known SRF target genes, and identified a
putative role for SRF in the transcriptional regulation of
69 additional genes. Since our arrays covered approximately 25% of all murine genes, we anticipate additional
SRF target genes to exist. In addition, our approach
misses SRF target genes, which are activated less than
3-fold or not activated by SRF-VP16, e.g., Mcl-1 (Schratt
et al., 2004).
SRF-VP16-Inducible Genes Display Distinct
Responses to Serum Stimulation
Expression of SRF-VP16 in Srf⫺/⫺ ES cells induces morphological changes (Schratt et al., 2002) which might
affect gene expression indirectly. We therefore used
serum stimulation of ES cells as an independent way to
induce SRF target genes. Srf⫹/⫹, Srf⫺/⫺ or Srf⫺/⫺rescue ES
cells were stimulated up to 180 min by serum addition
in the presence of the protein synthesis inhibitor cycloheximide (Supplemental Table S2). Gene expression
profiles were monitored using Affymetrix microarrays.
For further analysis, we focused on the subset of 86
genes, which were also activated by SRF-VP16 (see
above). Self-organizing map (SOM) clustering of the selected 86 genes yielded four different clusters, each
representing a group of genes sharing a common serum
induction profile (Figure 1A). Only genes found in clusters b (51 genes) and d (8 genes) displayed a robust
serum induction. The remaining 27 genes in clusters a
and c, although activated by SRF-VP16, were not induced strongly by serum. Genes in cluster d displayed
the classical induction profile of IEGs and included the
known SRF-responsive IEGs Egr-1, Egr-2, JunB, Cyr61, and ␥-actin (Figure 1B). SRF-regulated cytoskeletal
genes, such as SM22␣, muscle actins and vinculin, displayed a more transient serum induction profile and
were primarily found in cluster b. In total, 14 out of
17 previously known SRF target genes were found in
clusters b and d, confirming our initial hypothesis that
the expression of most SRF target genes would be induced by both SRF-VP16 and serum stimulation. We
infer that clusters b and d are enriched for novel SRF
target genes. Therefore, scoring for independent activation by SRF-VP16 and serum represents a stringent filter
to identify novel bona fide SRF target genes.
Identification of Conserved SRF Binding Sites
by a Comparative Genomic Approach
To address whether SRF directly binds to the promoters
of genes activated by SRF-VP16, we screened the regulatory regions of the 86 identified genes for putative
SRF consensus binding sites. A 15 kb DNA sequence
upstream of the first at least partially translated exon
in the genomic sequences of mouse and human was
analyzed. To narrow down the search space, we only
considered binding sites that localized within conserved
noncoding blocks (CNBs) of mouse and human genomic
sequences (Dieterich et al., 2003). By restricting our
search space, we necessarily miss SRF binding sites
within intronic regions and upstream regions with insufficient homology for the identification of CNBs. Within
CNBs, we screened for putative SRF binding sites,
allowing one base pair deviation (CArG-like) from the
canonical SRF consensus sequence (CC(A/T)6GG) (CArG).
CArG-like sequences often constitute functional SRF
binding sites (Miano, 2003). Using these criteria, we
FHL2, a Rho-Dependent Antagonist of SRF
869
Table 1. Functional Clustering of Genes Upregulated by SRF-VP16 in Two Independent Srf⫺/⫺ ES Cell Lines, 81 Srf
⫺/⫺
and 100 Srf⫺/⫺
Fold Activation
Accession Number
Gene
81 Srf⫺/⫺ (1)
81 Srf⫺/⫺ (2)
100 Srf⫺/⫺ (1)
100 Srf⫺/⫺ (2)
SM22␣
␣-actin (smooth)
ACTA1 (skeletal)
NPPB
CRP-1
ACTC (cardiac)
cTnC
H-FABP
Fhl2
CNN-1
MLC-C
LPP
235.1
98.6
80.1
31.8
75.6
9.0
3.4
24.6
14.7
5.4
3.6
3.1
151.2
11.8
73.6
63.0
32.3
26.3
20.1
4.5
10.6
4.8
7.5
12.4
251.4
104.3
201.9
71.5
31.2
8.5
14.6
7.6
3.9
12.8
20.5
6.1
151.4
137.8
88.3
54.0
28.4
14.5
9.8
3.7
10.6
7.4
5.4
4.5
144.1
90.8
23.9
17.0
4.0
15.6
6.8
16.9
3.2
15.7
169.9
151.5
56.6
36.4
26.6
20.4
16.6
22.1
18.8
4.1
271.8
112.7
173.4
7.6
11.5
14.5
10.5
1.9
4.4
6.1
185.4
73.1
56.6
12.5
28.9
11.7
24.2
2.3
5.2
3.0
tissue factor
CTGF
endothelin-1
inhibin-betaA
Cyr61
WISP-1
MuPAR1
28.5
38.7
15.8
20.5
17.0
7.3
7.5
104.4
13.5
22.6
15.8
16.6
6.2
5.4
26.5
58.6
36.9
11.3
19.6
11.0
3.3
33.9
5.2
18.6
19.8
10.3
3.9
3.1
lipocortin 1 (2x)
keratin 17 (2x)
annexin A2
keratin 19
keratin 18
NICE-1
annexin III
vinculin
perlecan
gelsolin
APP
keratin 7
desmoplakin
keratin 8
claudin-6
tropomyosin 4
TC10
procollagen, type IV
CPE receptor
tuftelin
Fyn
␥-actin (smooth)
beta-galactoside specific lectin
Aim
keratin 14
Arhu
101.0
17.8
179.0
13.5
22.5
11.3
26.0
70.4
3.6
15.4
25.4
4.9
23.9
14.1
12.0
23.6
5.8
16.3
7.7
4.4
4.8
4.8
7.2
6.9
6.5
3.9
66.8
77
34.8
25.4
36.0
46.1
69.3
8.5
47.3
12.9
10.9
9.7
7.6
16.2
20.0
5.3
25.6
7.9
6.5
5.6
4.7
3.1
7.2
9.2
6.6
5.4
55.8
104.2
19.9
121.0
54.2
13.0
13.2
10.9
7.8
24.0
14.3
25.8
15.2
12.0
10.6
11.1
10.9
3.5
6.1
6.4
4.6
3.1
5.8
16.5
17.0
4.3
81.3
68.4
15.6
20.3
18.6
58.8
17.0
7.3
4.6
7.5
8.1
14.1
4.4
8.1
7.0
5.7
3.1
3.7
3.8
4.4
4.5
4.4
4.6
19.0
11.6
5.7
cytochrome b-558 (2x)
5.6
11.2
7.3
4.8
Muscle Specific (12)
Z68618
X13297
M12347
D16497
D88793
M15501
M29793
X14961
AF055889
U28932
AI842649
AI850370
Cell Cycle/Apoptosis (10)
X81584
X71922
M28845
AF058798
U09268
M35523
M24377
U20735
X67644
Z38110
IGFBP-6
IGF-2
Egr-1
14-3-3␴
PAC-1
CRABP-2
Egr-2
Jun B
IER3
PMP-22
Wound Healing/Angiogenesis (7)
M26071
M70642
U35233
X69619
M32490
AF100777
X62700
Cytoskeleton (28)
M69260
M13805
M14044
M36120
M22832
AI604345
AJ001633
AI462105
M77174
J04953
U82624
AA755126
AA600542
X15662
AF087824
AI835858
AW060401
M15832
AB000713
AF047704
AW046449
M21495
X15986
AA711704
AA606367
AW121294
Metabolism (9)
M31775
(continued)
Molecular Cell
870
Table 1. Continued
Fold Activation
Accession Number
Gene
81 Srf⫺/⫺ (1)
81 Srf⫺/⫺ (2)
100 Srf⫺/⫺ (1)
100 Srf⫺/⫺ (2)
malic enzyme
YSPL-1
secretin
Sid478p
lipoprotein lipase
calcyclin
cathepsinB
6.8
3.5
4.0
7.3
6.2
8.4
5.8
6.4
7.0
7.4
4.3
16.4
14.0
4.2
7.0
9.4
8.0
5.2
7.6
3.8
3.7
4.5
10.4
5.8
5.0
4.0
6.7
3.3
interferon-induced 15 kDa protein
PRNP (prion)
Stra13/Clast5
beta2 microglobulin
IGFBP-5 protease
C/EBP beta
nestin
PSD-95
calcium channel 8
TIP-1
PolI transcription related factor
cystatin
MSSP
IFI-30
protective protein for ␤-galactoside
serine protease, 43 kD
TAP-binding protein
Hes-related
unknown EST
KDT-1
25.3
27.6
27.7
32.5
23.4
12.5
11.1
5.6
6.9
3.1
7.3
6.4
5.0
3.4
4.2
7.4
7.9
9.8
8.7
3.9
11.1
23.3
27.9
7.5
17.9
8.1
7.2
9.4
7.5
5.7
6.0
5.1
5.0
6.5
5.3
6.5
11.6
4.9
6.3
5.4
93.6
10.2
6.0
15.0
4.0
16.9
10.0
7.9
6.3
7.9
3.6
3.9
3.8
3.5
3.0
14.0
8.3
7.4
9.6
4.3
21.2
9.6
4.3
3.8
4.7
2.4
4.3
6.9
3.1
5.6
3.8
3.3
4.9
3.7
3.1
8.4
6.9
2.5
4.2
5.7
Metabolism (9)
J02652
U25739
X7380
AB025408
AA726364
X66449
M65270
Others (20)
X56602
M18070
Y07836
X01838
AW125478
M61007
AW061260
AI840413
AI849587
AI842665
AI845915
U59807
AB026569
AI844520
J05261
AW228316
AI836367
AW214298
AW122893
U13371
Values represent individual activation of all genes upregulated at least 3-fold in two independent experiments, (1) and (2). Known SRF target
genes are displayed in bold.
identified a total of 28 putative SRF binding sites within
the upstream regions of the 86 genes induced by SRFVP16 (Supplemental Table S3). This frequency (32.6%) is
significantly higher (p ⫽ 1.023 ⫻ 10⫺5) than the frequency
obtained if one were to pick 86 genes randomly from
the mouse genome (1,864 in a total of 13,540 mouse
genes, resulting in a frequency of 13.8%). If the analysis
is restricted to perfect CArG consensus sequences, the
difference is more pronounced (11.6% [10 out of 86
genes] versus 1.1% [151 out of 13,540 genes]; p ⫽
4.289 ⫻ 10⫺8). This illustrates that our gene expression
profiling significantly enriched for putative SRF targets.
The majority of genes containing putative SRF binding
sites (23 out of 28) display a serum induction profile of
the clusters b and d. We found SRF binding sites in
CNBs of 10 out of 17 previously known SRF targets,
illustrating that the majority of functional SRF binding
sites can be identified with our comparative genomics
approach.
Eighteen previously unrecognized SRF target genes
were identified (Supplemental Table S3). Three of these
genes contain a conserved consensus CArG sequence
(tuftelin, Fhl2, and KRT-17). The remaining 15 genes
carry CArG-like sequences (e.g., ET-1), some of which
are only partially conserved between mouse and human
genomes (e.g., CTGF). In summary, we identified SRF
binding sites in 10 previously known and in 18 new SRF
target genes using comparative genomic sequence
analysis.
Validation of SRF-Regulated Genes by RT-PCR
and Chromatin Immunoprecipitation
We next verified SRF-dependent expression for several
genes identified in our screen using quantitative RTPCR. We focused on three genes containing a conserved consensus CArG box (tuftelin, Fhl2, and KRT17) and two genes containing CArG-like sequences
(CTGF and ET-1). In Srf⫺/⫺ ES cells, SRF-VP16 significantly upregulated the mRNA levels of tuftelin, CTGF,
KRT-17, ET-1, and Fhl2 compared to control cells (Figures 2E–2I). A similar induction was observed for the
known SRF target genes Egr-1, SM22␣, and SMA (Figures 2B–2D). Expression of the housekeeping gene
GAPDH did not change upon expression of SRF-VP16
(Figure 2A). Similar results were obtained using an independent Srf⫺/⫺ ES cell line (data not shown). Using quantitative RT-PCR, we validated that SRF-VP16 induced
the expression of at least five of the SRF-responsive
genes identified in our screen.
To determine whether SRF binds directly to the identified CArG box sequences in the promoter regions of
tuftelin, CTGF, KRT-17, ET-1, and Fhl2 in vivo, we performed ChIP assays (Figure 2J). Differentiated ES cells
were used for the ChIP assays since some SRF target
promoters, e.g., muscle-specific promoters, might not
be occupied by SRF in undifferentiated ES cells (Manabe
and Owens, 2001). SRF bound specifically to CArG box
sequences of the known SRF target genes Egr-1, Srf,
and SMA in E14 Srf⫹/⫹ ES cells (lane 6). No signal was
FHL2, a Rho-Dependent Antagonist of SRF
871
Figure 1. Genes Inducible by SRF-VP16 Display Different Serum Induction Profiles
(A) Serum induction profiles of genes in E14 Srf⫹/⫹ ES cells were clustered based on the self-organizing map (SOM) algorithm. Four different
clusters were obtained: a, weak; b, moderate and transient; c, no; and d, strong and sustained serum induction.
(B) Individual serum induction profiles of genes grouped into clusters a–d according to Figure 1A. In addition to E14 Srf⫹/⫹ ES cells, the
induction kinetics for 100 Srf⫺/⫺ and 100-2 Srf⫺/⫺rescue ES cells are shown. Degree of induction is reflected by relative color coding: blue, low
expression; red, high expression. The numbers to the right of the induction profiles represent the fold activation for serum-treated E14 Srf⫹/⫹
ES cells at the 60 min time point.
Molecular Cell
872
Figure 2. Validation of SRF-Dependent Activation of Selected Genes Identified by Microarray Profiling
(A–I) Quantitative RT-PCR was performed with mRNA from 100 Srf⫺/⫺ ES cells, transiently transfected with SRF-VP16 or SRF-⌬M-VP16, using
specific primers for GAPDH (A), Egr-1 (B), SM22␣ (C), SMA (D), tuftelin (E), CTGF (F), KRT-17 (G), ET-1 (H), and Fhl2 (I). Values represent
transcript levels relative to the endogenous housekeeping gene HPRT and are the mean of three independent experiments. Asterisk denotes
statistically significant induction (p ⬍ 0.05, Student’s t test).
(J) ChIP analysis of SRF binding to promoters of SRF target genes in vivo. Chromatin of day 8 differentiated 100 Srf⫺/⫺ and E14 Srf⫹/⫹ ES
cells was immunoprecipitated using a polyclonal anti-SRF antibody (lanes 3 and 6). Bound DNA fragments were amplified by PCR using
primers specific for the CArG-containing promoter regions of the indicated genes. Incubation without antibody was used as control (lanes 2
and 5). 1% of the isolated genomic DNA served as input (lanes 1 and 4).
observed in 100 Srf⫺/⫺ ES cells (lane 3). SRF also bound
specifically to the CArG box sequences in the tuftelin,
CTGF, KRT-17, ET-1, and Fhl2 promoters (lane 6), but
not to the CArG box-deficient ␤-globin promoter. Thus,
our results demonstrate that the promoter regions of
tuftelin, CTGF, KRT-17, ET-1, and Fhl2 are bound by SRF
in native chromatin of differentiating ES cells, identifying
these genes as direct SRF target genes.
FHL2, a Rho-Dependent Antagonist of SRF
873
Figure 3. Expression of FHL2 Is Regulated by SRF in a RhoA- and Differentiation-Dependent Manner
(A) 81 Srf⫺/⫺ and 100 Srf⫺/⫺ ES cells were transiently transfected with SRF-VP16 or SRF-⌬M-VP16, and protein extracts were prepared 72 hr
after transfection. Western blotting was performed using polyclonal anti-FHL2 or anti-␣-actinin antibody.
(B) NIH3T3 cells were transfected with luciferase reporter constructs driven by the human FHL2 promoter with an intact or mutated CArG
sequence (500 ng), in the presence or absence of RhoAV14 (100 ng). Values are given as fold activation over vector-transfected samples and
represent the mean of four independent experiments.
(C) 100 Srf⫺/⫺ and E14 Srf⫹/⫹ ES cells were differentiated by LIF removal under monolayer conditions for up to 8 days. Protein lysates were
prepared on the indicated days and subjected to Western Blotting using polyclonal anti-FHL2 or anti-␣-actinin antibody. FHL2 protein is
indicated by the arrow.
(D) 99 Srf⫺/⫹ ES cells were differentiated in the presence of retinoic acid for up to 8 days and mRNA expression was analyzed by quantitative
RT-PCR. Values represent the mean of two independent experiments ⫾ SD.
SRF Regulates the Expression of FHL2 in a RhoAand Differentiation-Dependent Manner
The Fhl2 gene was chosen for further study since FHL2
can function as both transcriptional coactivator and corepressor of several transcription factors and since FHL2
activity is regulated by the RhoA signaling pathway.
Transient transfection of SRF-VP16 specifically induced Fhl2 protein expression in Srf⫺/⫺ ES cells (Figure
3A). Therefore, SRF-VP16 is sufficient to induce expression of endogenous Fhl2 protein in Srf⫺/⫺ ES cells. SRFdependent regulation of Fhl2 was further analyzed using
luciferase reporter genes driven by the CArG box-con-
Molecular Cell
874
Figure 4. SRF and FHL2 Interact In Vitro and
In Vivo
(A) GST pull-down assays were performed
with the indicated GST fusion proteins and
in vitro-translated 35S-labeled SRF. 1% of 35Slabeled SRF was loaded as input. Coomassie
staining shows comparable expression of the
GST fusion proteins used.
(B) Mapping of the SRF interaction domain of
FHL2. GST pull-down assays were performed
as in (A).
(C) SRF Core is sufficient to interact with
FHL2. GST pull-down assay was performed
with GST-FHL2 and in vitro translated 35Slabeled SRF Core (aa 133–265). 10% of 35Slabeled SRF Core was loaded as input.
(D) Immunoprecipitation of 293T cells transfected with HA-SRF and Flag-FHL2 in the absence or presence of RhoAV14 was performed using M2 anti-Flag or an IgG control
antibody. Immunoprecipitated SRF or FHL2
protein was detected by Western blotting
with anti-SRF or anti-Flag M2 antibody, respectively. Input represents 4% of the starting protein lysate.
(E) Immunoprecipitation of rat embryonic
heart extracts was performed using anti-SRF
or an IgG control antibody. Immunoprecipitated SRF or FHL2 protein was detected by
Western blotting with anti-SRF or anti-FHL2
antibody.
(F) SRF and FHL2 bind to the promoters of
SM genes. Differentiated E14 Srf⫹/⫹ ES cells
were transfected with HA-SRF, Flag-FHL2,
and RhoAV14 as indicated. ChIP assay was
performed using an anti-SRF (lanes 3 and 7)
or anti-Flag antibody (lanes 4 and 8). No antibody incubation served as control (lanes 2
and 6). 1% of the isolated genomic DNA
served as input (lanes 1 and 5).
taining human FHL2 promoter. As a control, a CArG
box mutation abolishing SRF binding was generated. In
NIH3T3 cells, the FHL2 promoter construct was activated 12-fold by coexpression of constitutively active
RhoAV14. CArG box mutation completely prevented
RhoA-mediated activation (Figure 3B). Therefore, SRF
binding to the identified CArG box in the FHL2 promoter
is essential for its activation by RhoA.
Upon monolayer ES cell differentiation, Fhl2 protein
expression increased strongly, reaching maximum levels at day 4 (Figure 3C). This effect is SRF dependent,
since Fhl2 protein expression was not induced in differentiated Srf⫺/⫺ ES cells. Similarly, ES cell differentiation
in the presence of retinoic acid resulted in a gradual
increase of Fhl2 mRNA expression from day 2 until day 8
(Figure 3D). Fhl1 mRNA levels increased in a comparable
manner. SM22␣, SMA, and cardiac actin mRNA peaked
at day 4 and decreased afterwards (Figure 3D). Therefore, expression of FHL family members precedes expression of smooth and cardiac muscle markers in differentiating ES cells, suggesting a role for FHL proteins
in the regulation of SRF-dependent muscle genes.
SRF and FHL2 Interact In Vitro and In Vivo
Since SRF is able to interact with the LIM-only proteins
CRP1 and 2, we investigated a potential direct interac-
tion between SRF and FHL family members. We performed GST pull-down assays using in vitro-translated
35
S-labeled SRF and GST-FHL fusion proteins. FHL2 efficiently bound SRF, demonstrating that SRF and FHL2
interact directly in vitro (Figure 4A). No interaction of
SRF was observed with GST alone. SRF interacted with
FHL1 to a similar extent as with FHL2. In contrast, interaction of SRF with FHL3, FHL4, and ACT was either
weak or not detectable (Figure 4A). Therefore, SRF is
able to interact in vitro with at least two members of the
FHL protein family, FHL1 and FHL2.
To map the SRF binding domain of FHL2, we tested
different FHL2 mutants in the GST pull-down assay.
LIM3-4 bound SRF to a similar extent as full-length FHL2
(Figure 4B). In contrast, neither LIM0-2 nor LIM3/4 alone
bound SRF. This indicates that LIM domains 3 and 4 of
FHL2 are necessary and sufficient for the interaction
with SRF. An SRF mutant consisting only of the DNA
binding/dimerization domain (SRF Core) still interacts
with FHL2 (Figure 4C).
To investigate whether SRF and FHL2 interact in vivo,
we performed coimmunoprecipitation assays using
293T cells transfected with HA-SRF and Flag-FHL2 in
the absence or presence of RhoAV14. HA-SRF coimmunoprecipitated with Flag-FHL2 only in the presence of
active RhoA (Figure 4D). To explore a potential interac-
FHL2, a Rho-Dependent Antagonist of SRF
875
tion between endogenous SRF and FHL2, we performed
coimmunoprecipitation assays using rat embryonic
heart extracts. We detected FHL2 in anti-SRF immunoprecipitates, indicating that SRF and FHL2 interact in
cells of the developing heart (Figure 4E). FHL2 and SRF
colocalize in the nucleus of rat cardiomyocytes, suggesting that the FHL2/SRF interaction occurs in the nucleus (Supplemental Figure S4).
Together, these experiments show that SRF and FHL2
directly interact in vitro and form a RhoAV14-dependent
complex in vivo.
FHL2 and SRF Bind to the Promoters
of SM-Specific SRF Target Genes
We used ChIP to determine whether FHL2 and SRF bind
to the promoters of SRF target genes in differentiated
E14 Srf⫹/⫹ ES cells (d8). As expected, SRF was bound
to the promoters of the SRF target genes Egr-1, Srf,
SMA, and SM22␣ (Figure 4F, lanes 3 and 7). FHL2 specifically bound to the same SM22␣ and SMA promoter
regions recognized by SRF, and FHL2 binding was
strongly increased upon RhoA activation (lanes 4 and
8). In contrast, FHL2 binding was not detectable at the
Egr-1 and Srf promoters irrespective of RhoA activity.
Neither SRF nor FHL2 were able to bind to the CArG
box-deficient ␤-globin promoter. Our results indicate
that FHL2 is selectively recruited to the promoters of
the SM-specific SRF target genes SMA and SM22␣ in
response to RhoA signaling.
FHL2 Inhibits SRF-Dependent Transcription
of SM Genes
We next investigated the functional consequences of
FHL2 binding to SRF target promoters. FHL2 and
RhoAV14 were transiently expressed in 293T cells, and
the activity of SRF-dependent reporter genes was monitored. RhoAV14 expression induced activation of the
SM22␣ and SMA promoters, which was significantly inhibited by FHL2 in a dose-dependent manner (Figures
5A and 5B). Mutation of CArG box 1 of the SMA promoter
diminished activation by RhoAV14, whereas mutation of
CArG box 2 almost completely abolished this activation.
FHL2 was not able to further repress the RhoA nonresponsive SMA promoter constructs (Figure 5B). Although the cardiac-specific ANF promoter was efficiently activated by RhoAV14, no significant inhibition
by FHL2 was observed, suggesting promoter selectivity
by FHL2 (Figure 5C). Similarly, FHL2 failed to diminish
the activity of the c-fos or tk80 promoters (Figures 5D
and 5E). The FHL2 point mutant C7A fails to accumulate
in the nucleus upon RhoA activation (Müller et al., 2002).
In contrast to wt FHL2, expression of FHL2 C7A did not
antagonize RhoA-mediated SM22␣ promoter activation
(Figure 5F), indicating that inhibition of SRF-dependent
transcription by FHL2 requires nuclear FHL2. To address
whether SRF binding is critical for the FHL2-mediated
antagonism, we used the FHL2 mutant LIM0-2, which
is unable to bind SRF (Figure 4B), but efficiently translocates to nuclei upon RhoA activation (Müller et al.,
2002). FHL2 LIM0-2 was no longer able to antagonize
RhoA-mediated SM22␣ promoter activation in 293T cells,
demonstrating that interaction with SRF is required for
FHL2-mediated inhibition (Figure 5G). Equal expression
of full-length and mutant FHL2 proteins was confirmed
by Western blotting (data not shown). We conclude that
FHL2 antagonizes RhoA-dependent activation of specific SRF target genes.
FHL2 Antagonizes MAL
MAL is a potent SRF-dependent activator of both
smooth and cardiac muscle genes. Similar to FHL2, MAL
has been shown to accumulate in the nucleus after RhoA
activation. We investigated whether FHL2 could interfere with MAL activation of SRF target genes in response
to RhoA in 293T cells. MAL and RhoAV14 synergistically
activated the SMA, SM22␣, and ANF promoters. Coexpression of FHL2 led to a significant reduction in SM22␣
and SMA activation, but did not interfere with activation
of the ANF promoter (Figures 6A–6C). Similarly, expression of FHL2 did not affect c-fos or tk80 promoter activities (Figures 6D and 6E). This suggests that FHL2 can
interfere with MAL-mediated activation of a specific
subset of RhoA-responsive SRF target genes, including
the SM-specific genes SM22␣ and SMA.
In undifferentiated Srf⫺/⫺ ES cells, expression of SRF
and MAL led to an efficient induction of SMA and SM22␣
mRNA levels, which could be further increased by coexpression of RhoAV14. Simultaneous coexpression of
FHL2 significantly reduced RhoA/MAL-mediated activation (Figures 6F and 6G), demonstrating that FHL2 is able
to interfere with RhoA/MAL-stimulated transcription of
the endogenous SM-specific SRF target genes SMA and
SM22␣. In contrast, RNAi-mediated knockdown of Fhl2
had no effect on the expression of SM22␣, SMA, and
cardiac actin during ES cell differentiation (Supplemental Figure S5), possibly due to functional redundancy
with other FHL proteins that are expressed in ES cells
and interact with SRF (Figures 3D and 4A).
Competition between FHL2 and MAL
for Interaction with SRF
FHL2 antagonizes MAL-mediated activation of SM
genes and binds to the same region of SRF as MAL
(Figures 4C and 6A–6G). We used EMSA to test whether
FHL2 could compete with MAL for SRF binding. Consistent with a previous report, we observed an SRF-containing ternary complex when extracts expressing an
N-terminal deletion mutant, MAL⌬N, were incubated
with a CArG box oligonucleotide (Miralles et al., 2003)
(Figure 6H). The identity of the complexes was confirmed
using in vitro-translated MAL⌬N and recombinant SRF
protein (data not shown). Addition of increasing amounts
of in vitro-translated FHL2 diminished the formation of
the MAL⌬N/SRF/DNA complex in a dose-dependent
manner (Figure 6H). We were unable, however, to detect
an SRF/FHL2 complex with these EMSA conditions. This
result suggests that FHL2 and MAL compete for the
same binding site on SRF, thereby providing a possible
mechanism for the observed inhibition of MAL-mediated
activation of SRF target genes by FHL2.
Discussion
We describe here a microarray expression profiling
study that led to the identification of several SRF-regulated genes. In particular, we show that the gene encod-
Molecular Cell
876
Figure 5. FHL2 Represses SRF-Dependent SM Promoters
(A–E) 293T cells were transiently transfected with the indicated luciferase promoter constructs, increasing amounts of FHL2 expression plasmid
(100 ng, 500 ng, 1 ␮g, 1.4 ␮g), and 100 ng RhoAV14 as indicated. Values represent fold activation compared to vector control-transfected
cells and are the mean of three independent experiments. Asterisk denotes statistically significant repression (p ⬍ 0.05, Student’s t test).
(F) FHL2-mediated repression is dependent on nuclear localization. 293T cells were transfected with SM22␣-Luc and the FHL2 or C7A mutant
expression plasmid in the presence or absence of RhoAV14. Values represent percent activation relative to RhoAV14-transfected cells and
are the mean of three independent experiments.
(G) FHL2-mediated repression is dependent on interaction with SRF. 293T cells were transfected with SM22␣-Luc and the indicated FlagFHL2 expression plasmid in the presence or absence of RhoAV14. Values represent percent activation relative to RhoAV14 transfected cells
and are the mean of three independent experiments. Asterisk denotes statistically significant repression (p ⬍ 0.05, Student’s t test).
ing the LIM-only protein FHL2 is an SRF target gene
and further implicate FHL2 in the regulation of SRFdependent gene control.
Functional Clusters of SRF Target Genes
Our expression profiling approach uncovered a total of
59 genes whose expression is induced by both SRFVP16 and serum. Functional clustering of these genes
suggests a role for the transcription factor SRF in several
different processes, including cytoskeletal organization,
apoptosis, and myogenesis, processes in which SRF
has been previously implicated. Furthermore, we identify more SRF target genes involved in wound healing
(CTGF, ET-1, KRT-17) in addition to Egr-1. This suggests
that SRF regulates gene expression in response to
wounding, which is in agreement with the finding that
serum stimulation activates a transcriptional program
related to the physiology of wound repair in fibroblasts
(Iyer et al., 1999). Interestingly, several of the new SRF
target genes, including CTGF and ET-1, were also identified by Iyer et al.. Our study now implicates SRF in the
regulation of these genes. The role of SRF in wound
healing in vivo will be addressed by conditional deletion
of Srf in the mouse skin.
Regulation of FHL2 Expression
Several SRF target genes, including Fhl2, are preferentially expressed in muscle, emphasizing the pivotal role
FHL2, a Rho-Dependent Antagonist of SRF
877
Figure 6. FHL2 Antagonizes MAL/RhoA-Mediated Activation of SM Promoters and Competes with MAL for SRF Binding
(A–E) 293T cells were transiently transfected with the indicated luciferase promoter constructs, 1.4 ␮g FHL2 expression plasmid, 100 ng
RhoAV14, and 25 ng MAL as indicated. Values represent fold activation compared to vector control-transfected cells and are the mean of
three independent experiments. Asterisk denotes statistically significant repression (p ⬍ 0.05, Student’s t test).
(F and G) Undifferentiated 100 Srf⫺/⫺ ES cells were transiently transfected with SRF expression plasmid (100 ng). Cotransfection with MAL
(20 ng), FHL2 (1.4 ␮g), and RhoAV14 (100 ng) expression plasmids was performed as indicated. mRNA was prepared 48 hr after transfection
and subjected to quantitative RT-PCR analysis. Values were normalized to the average activation by MAL as deduced from four independent
experiments. Asterisk denotes statistically significant repression (p ⬍ 0.05, Student’s t test).
(H) EMSA was performed using a 32P-labeled CArG box oligonucleotide, 293T cell extracts expressing MAL⌬N, and in vitro-translated FHL2.
DNA complexes of SRF and SRF/MAL⌬N are indicated.
of SRF in muscle differentiation and function. We identified Fhl2 to be robustly induced by both SRF-VP16
and serum in ES cells. Additional experiments con-
firmed Fhl2 as a direct SRF target gene that is activated
by SRF upon RhoA signaling. Several other musclerestricted SRF target genes, as well as the Srf gene
Molecular Cell
878
itself, were shown to be regulated by the RhoA/actin
signaling (Miano, 2003; Sotiropoulos et al., 1999). Since
RhoA activity increases during myogenesis (Sordella et
al., 2003), RhoA-stimulated and SRF-dependent expression of FHL2 might represent a mechanism to ensure
FHL2 expression in cardiac muscle. However, since
RhoA/actin signaling regulates SRF target genes in different muscle lineages, additional mechanisms must exist that confer cardiac specific FHL2 expression. In this
regard, a conserved binding site for the homeodomain
transcription factor Nkx2.5, an SRF partner protein and
one of the earliest markers of the cardiac lineage, is
present in the Fhl2 promoter (Scholl et al., 2000), suggesting regulation of FHL2 expression by SRF and
Nkx2.5.
Regulation of SRF Activity by Cofactor Interactions
Interaction with transcriptional cofactors is widely used
to regulate SRF activity. The best-studied complex is the
one formed between SRF and TCFs on IEG promoters
(Buchwalter et al., 2004). In addition, increasing evidence highlights the importance of interactions between
SRF and cofactors on the promoters of muscle-specific
genes. Myocardin/MRTFs constitute a family of potent
coactivators of SRF. The ability of the SRF coactivator
MRTF-A/MAL to activate SRF-regulated transcription is
dependent upon RhoA-mediated nuclear translocation
(Miralles et al., 2003). Similarly, we demonstrated recently that RhoA regulates the transcriptional activity of
FHL2 by inducing its nuclear translocation (Müller et al.,
2002). Accordingly, activation of RhoA and subsequent
nuclear localization of FHL2 are required for FHL2-mediated inhibition of SRF target promoters and the observed antagonism of FHL2 and MAL. Together, these
findings suggest that FHL2, like MAL, links RhoA activity
to SRF-dependent gene expression. Since MAL remains
nuclear after the shutdown of target gene expression
(Miralles et al., 2003), FHL2 might participate in the downregulation of MAL activity after RhoA stimulation.
Myocardin/MRTFs are unable to discriminate between the promoters of cardiac and SM genes (Wang
et al., 2001, 2002). Therefore, mechanisms must exist
that interfere with Myocardin/MRTF-activated SM gene
expression in the heart. Given the abundant expression
of FHL2 in the heart, it is conceivable that FHL2 functions
as an inhibitor of SM genes in cardiac muscle. In addition
to FHL2, other SRF corepressors are known. GATA transcription factors repress myocardin activity at some SRF
target genes including SM22␣ and ANF (Oh et al., 2004).
HOP-mediated repression in the heart is not restricted
to SM-specific SRF targets and, therefore, likely regulates the balance between cardiomyocyte proliferation
and differentiation (Chen et al., 2002; Shin et al., 2002).
Recently, a role for the TCF Elk-1 in the repression of
SRF-dependent SM genes was reported (Wang et al.,
2004). Since Elk-1 is a target of MAPK signaling, it will
be interesting to determine how both MAPK and RhoA
signaling are integrated at different SRF-responsive promoters. Interestingly, in addition to FHL2-mediated inhibition of the RhoA pathway, FHL2 has been shown to
inhibit MAPK signaling (Purcell et al., 2004), suggesting
a regulatory contribution of FHL2 to both these signaling
pathways that target SRF.
Mechanism of FHL2-Mediated Target
Gene Repression
FHL2 inhibits the expression of a specific subset of SRF
target genes after RhoA/MAL-mediated activation. Our
results suggest that FHL2 and MAL compete for the
same binding site on SRF. Since RhoA stimulation
strongly increases FHL2 expression and nuclear localization, we hypothesize that FHL2 displaces MAL at
certain SRF-dependent promoters in response to RhoA
activation, leading to subsequent reduction in the expression of these genes. A similar mechanism has recently been described for the antagonism between Myocardin and Elk-1 (Wang et al., 2004). We found that
the HDAC inhibitor trichostatin A did not relieve FHL2mediated repression suggesting that HDAC repressor
complexes are not involved (U.P., unpublished data).
The molecular basis for the promoter selectivity of FHL2mediated repression is currently unknown. However, as
we observe this inhibition in the heterologous 293T cell
system, we speculate that DNA sequence elements
present in specific subsets of SRF-responsive promoters may mediate accessibility of SRF to FHL2. Since
direct DNA binding of FHL2 has not been reported, such
a sequence element would likely be either bound by a
factor that specifically recruits FHL2 to promoters, e.g.,
SM promoters, or by a factor that specifically inhibits
FHL2 binding to other SRF target genes, e.g., cardiac
muscle and growth-related genes. Future work is
needed to identify the components of FHL2 containing
complexes at SRF target promoters.
Taken together, our data suggest that SRF and FHL2
are components of an autoregulatory feedback loop that
antagonizes RhoA/MAL-mediated activation of smooth
muscle SRF target genes. Such mechanisms might orchestrate the lineage-specific expression of SRF target
genes in different muscle tissues.
Experimental Procedures
Plasmids
The plasmids SRF-VP16, SRF⌬M-VP16, wtSRF, RhoAV14, tk80-luc,
SRE2-luc, and c-fos-luc (Schratt et al., 2002), as well as the plasmids
pCMXPL2, pCMX-FHL2, pCMX-Flag-FHL2, pCMXGal-FHL2-C7A,
pGEX-FHL2, pGEX-FHL2 LIM0-2, pGEX LIM3-4, pGEX LIM3, and
pGEX LIM4 (Müller et al., 2000, 2002) have been described previously. pCMX-FHL2-C7A was derived from pCMXGal-FHL2-C7A.
For the GST-expression plasmids, the ORFs of mFHL1, mFHL3,
mFHL4, and mACT were cloned into pGEX4-T-1 (Amersham). FlagFHL2 and Flag-LIM0-2 were generated by cloning FHL2 or LIM0-2
into pcDNA6/Flag. The plasmids SM22␣-luc, ANF-luc, MRTF-A, HASRF, SMA-luc, SMASRE1pm-luc, and SMASRE2pm-luc have been
described (Chang et al., 2003; Wang et al., 2002). MAL⌬N was cloned
by deleting the 80 N-terminal amino acids of MRTF-A. The FHL2
promoter reporter constructs, FHL2 and FHL2mutated, were generated by PCR amplification of human genomic DNA and cloned into
pGL3basic (Promega). The CArG box was mutated to the sequence
CCATATAACT (mutated bases are shown in bold) resulting in
FHL2mutated.
Cell Culture and Transfection
The ES cell lines E14 Srf⫹/⫹, 99 Srf⫺/⫹, 81 Srf⫺/⫺, 100 Srf⫺/⫺, and 100-2
Srf⫺/⫺rescue were cultured as described previously (Weinhold et al.,
2000). Transient transfection of ES cells was performed using Lipofectamine 2000 (Invitrogen) as reported (Schratt et al., 2002). Serum
stimulation of ES cells was done as described previously (Schratt
et al., 2001), except that cycloheximide was added 30 min prior
to stimulation.
FHL2, a Rho-Dependent Antagonist of SRF
879
For monolayer ES cell differentiation, cells were washed twice
with ES media lacking LIF and cultured in the absence of LIF. For
differentiation in the presence of Retinoic Acid (RA), 5 ⫻ 10⫺7 M alltrans RA (Sigma) was added to ES media lacking LIF.
293T cells were cultured in DMEM containing 10% FCS, penicillin,
streptomycin, and L-Glutamin (GIBCO-BRL). For reporter assays,
293T cells were transfected using Lipofectamine 2000 (1:2 ratio of
DNA to Lipofectamine 2000). Luciferase activity was determined 48
hr after transfection.
NIH3T3 cells were cultured in DMEM containing 10% FCS, penicillin, streptomycin, and L-Glutamin. For reporter assays, NIH3T3 cells
were transfected using DOTAP (Roche) and luciferase activity was
assayed 24 hr after transfection.
Microarray Analysis
Total RNA was isolated from ES cells using the RNeasy Mini Kit
(Qiagen). After DNase treatment, RNA was reverse transcribed using
Oligo-dT and Superscript II (Invitrogen) followed by second-strand
synthesis. Double-stranded cDNA was transcribed in vitro using
the Ambion T7 MegaScript Kit in the presence of biotin-labeled
ribonucleotides (Enzo). Biotin-labeled cRNA was fragmented at 95⬚C
for 35 min and subsequently hybridized onto AffymetrixMurine Genome U74A Gene Arrays (12,500 probe sets). We note that the U74
release of Affymetrix Gene Arrays contained a significant proportion
of nonfunctional probe sets (ca. 2600). For further details, see http://
www.affymetrix.com/support/technical/product_updates/mgu74_
product_bulletin.affx.
The raw expression data was scaled to obtain equal mean intensities for each individual array, and absent genes (A-calls) were omitted. Scaling factors were generally less than 1.2. Data normalization
and analysis was performed using the Gene Cluster Software 2.0
(http://www.broad.mit.edu/cancer/software/genecluster2/gc2.html).
For the SRF-VP16 data, genes were only considered if they showed
at least a 3-fold change in expression in all of the four replicates performed.
Comparative Sequence Analysis of Noncoding DNA
in Orthologous Gene Loci
Orthologous genes in man and mouse were defined by protein sequence comparisons following the procedure outlined by Tatusov
et al. (1997). We computed pairwise best BLAST hits for all human
and murine protein entries of ENSEMBL release 17. Upstream regions of putative orthologs were scanned for local sequence similarities.
15 kB of DNA sequences upstream of the start of translation (ATG)
were retrieved for all human and murine transcripts. Mammalianwide repeats and low-complexity regions were excluded from the
analysis. Significant local sequence similarities were detected by
computing suboptimal local alignments. Further details can be
found elsewhere (Dieterich et al., 2003).
Subsequently, we predicted putative SRF binding sites by scanning all conserved noncoding blocks for the consensus SRF binding
motif, the CArG box CC(A/T)6GG. One mismatch to the motif consensus was allowed in either species.
Quantitative RT-PCR
Total RNA preparation, cDNA synthesis, and quantitative PCR using
the SYBR Green technology (Perkin Elmer) were performed as described previously (Weinhold et al., 2000). See Supplemental Data
for primer sequences.
Chromatin Immunoprecipitation
Chromatin immunoprecipitation (ChIP) assays were performed using a ChIP assay kit (Upstate Biotechnology Inc.) as described previously (Schratt et al., 2004). Samples were derived from E14 Srf⫹/⫹
ES cells, which were differentiated under monolayer conditions for
8 days in the absence of LIF (d8). SRF-DNA complexes were immunoprecipitated using 2 ␮l polyclonal anti-SRF antibody (Santa Cruz).
For FHL2 ChIPs, undifferentiated (d0) or differentiated (d4) E14 Srf⫹/⫹
ES cells were transiently transfected with 1.5 ␮g HA-SRF and 5 ␮g
pCMX-Flag-FHL2 in the absence or presence of 0.5 ␮g RhoAV14.
d0 ES cells were fixed 2 days after transfection. Transfected d4 ES
cells were differentiated for an additional 4 days and fixed on day
8 (d8). Flag-FHL2-DNA complexes were immunoprecipitated using
2 ␮l M2 anti-Flag antibody (Sigma). For primer sequences see Supplemental Data. Standard PCR was performed using Platinum Taq
DNA Polymerase (Invitrogen) with an annealing temperature of 65⬚C
and 35 cycles. For the SM22␣ PCR, the annealing temperature was
60⬚C. For the Fhl2, SMA, and CTGF PCRs, the Advantage-GC Genomic PCR Kit (Clontech) was used.
Western Blotting and Coimmunoprecipitation
Cells were lysed in lysis buffer A, and Western blotting was performed according to standard procedures using polyclonal antiFHL2 (Müller et al., 2002) or monoclonal anti-␣-actinin antibody
(Clone BM-75.2; Sigma).
For coimmunoprecipitation, 293T cells were transfected with 1.5
␮g HA-SRF and 5 ␮g pCMX-Flag-FHL2 in the absence or presence
of 0.5 ␮g RhoAV14. Three days after transfection, cells were lysed
in GST-Buffer. Protein extracts were precleared using goat antimouse IgG beads (Sigma), incubated with M2 anti-Flag antibody
(Sigma) or an IgG control antibody (Sigma), and immunoprecipitated
with goat anti-mouse IgG beads. Eluates were analyzed by Western
blotting using the polyclonal anti-SRF (Santa Cruz) and M2 antiFlag antibody.
Rat embryonic hearts (Wistar rats, E19–E21; Charles River) were
dissected according to standard procedures and homogenized in
lysis buffer B. Protein extracts were diluted in GST-buffer, precleared using Protein A Agarose beads (Pierce) and incubated with
polyclonal anti-SRF or rabbit IgG antibody (Sigma) crosslinked to
Protein A Agarose beads (Pierce Seize X Protein A Immunoprecipitation Kit). Proteins were eluted and analyzed by Western Blotting
using the polyclonal anti-SRF and the polyclonal anti-FHL2 antibody. See Supplemental Data for buffer compositions.
GST Pull-Down
Expression of the GST fusion proteins (Pharmacia) and the in vitro
transcription-translation reactions (Promega) were performed according to the manufacturers’ protocols. GST pull-down assays using in vitro-translated human SRF or SRF Core were performed as
described previously, except that binding reactions were at 37⬚C
(Müller et al., 2000).
Electrophoretic Mobility Shift Assays
Electrophoretic mobility shift assays (EMSAs) were performed as
described previously (Weinhold et al., 2000). A 32P-labeled DNA fragment containing the c-fos CArG box in the context of mutated TCF
site was used as DNA binding probe.
Acknowledgments
We thank E. Olson and R. Schwartz for plasmids. U.P. received a
Ph.D. scholarship by the Boehringer Ingelheim Fonds. This work was
supported by the DFG (SFB 446/B7) and the Fonds der Chemischen
Industrie to A.N. and by the DFG (SFB388/C9) and the MildredScheel-Stiftung (10-2019-BU I) to R.S. F.G. was supported by NIH
grant GM58801. We thank T. Golub and J. Staunton for help with
microarray analysis.
Received: April 28, 2004
Revised: October 18, 2004
Accepted: November 22, 2004
Published: December 21, 2004
References
Arsenian, S., Weinhold, B., Oelgeschlager, M., Ruther, U., and Nordheim, A. (1998). Serum response factor is essential for mesoderm
formation during mouse embryogenesis. EMBO J. 17, 6289–6299.
Buchwalter, G., Gross, C., and Wasylyk, B. (2004). Ets ternary complex transcription factors. Gene 324, 1–14.
Chan, K.K., Tsui, S.K., Lee, S.M., Luk, S.C., Liew, C.C., Fung, K.P.,
Waye, M.M., and Lee, C.Y. (1998). Molecular cloning and characterization of FHL2, a novel LIM domain protein preferentially expressed
in human heart. Gene 210, 345–350.
Chang, D.F., Belaguli, N.S., Iyer, D., Roberts, W.B., Wu, S.P., Dong,
Molecular Cell
880
X.R., Marx, J.G., Moore, M.S., Beckerle, M.C., Majesky, M.W., et al.
(2003). Cysteine-rich LIM-only proteins CRP1 and CRP2 are potent
smooth muscle differentiation cofactors. Dev. Cell 4, 107–118.
Chen, F., Kook, H., Milewski, R., Gitler, A.D., Lu, M.M., Li, J., Nazarian, R., Schnepp, R., Jen, K., Biben, C., et al. (2002). Hop is an
unusual homeobox gene that modulates cardiac development. Cell
110, 713–723.
cytoskeletal organization and focal adhesion assembly in embryonic
stem cells. J. Cell Biol. 156, 737–750.
Schratt, G., Philippar, U., Hockemeyer, D., Schwarz, H., Alberti, S.,
and Nordheim, A. (2004). SRF regulates Bcl-2 expression and promotes cell survival during murine embryonic development. EMBO
J. 23, 1834–1844.
Dieterich, C., Wang, H., Rateitschak, K., Luz, H., and Vingron, M.
(2003). CORG: a database for COmparative Regulatory Genomics.
Nucleic Acids Res. 31, 55–57.
Shin, C.H., Liu, Z.P., Passier, R., Zhang, C.L., Wang, D.Z., Harris,
T.M., Yamagishi, H., Richardson, J.A., Childs, G., and Olson, E.N.
(2002). Modulation of cardiac growth and development by HOP, an
unusual homeodomain protein. Cell 110, 725–735.
Fimia, G.M., De Cesare, D., and Sassone-Corsi, P. (1999). CBPindependent activation of CREM and CREB by the LIM-only protein
ACT. Nature 398, 165–169.
Sordella, R., Jiang, W., Chen, G.C., Curto, M., and Settleman, J.
(2003). Modulation of Rho GTPase signaling regulates a switch between adipogenesis and myogenesis. Cell 113, 147–158.
Fimia, G.M., De Cesare, D., and Sassone-Corsi, P. (2000). A family
of LIM-only transcriptional coactivators: tissue-specific expression
and selective activation of CREB and CREM. Mol. Cell. Biol. 20,
8613–8622.
Sotiropoulos, A., Gineitis, D., Copeland, J., and Treisman, R. (1999).
Signal-regulated activation of serum response factor is mediated
by changes in actin dynamics. Cell 98, 159–169.
Hill, C.S., Wynne, J., and Treisman, R. (1995). The Rho family
GTPases RhoA, Rac1, and CDC42Hs regulate transcriptional activation by SRF. Cell 81, 1159–1170.
Treisman, R. (1995). Journey to the surface of the cell: Fos regulation
and the SRE. EMBO J. 14, 4905–4913.
Tatusov, R.L., Koonin, E.V., and Lipman, D.J. (1997). A genomic
perspective on protein families. Science 278, 631–637.
Iyer, V.R., Eisen, M.B., Ross, D.T., Schuler, G., Moore, T., Lee, J.C.,
Trent, J.M., Staudt, L.M., Hudson, J., Jr., Boguski, M.S., et al. (1999).
The transcriptional program in the response of human fibroblasts
to serum. Science 283, 83–87.
Wang, D., Chang, P.S., Wang, Z., Sutherland, L., Richardson, J.A.,
Small, E., Krieg, P.A., and Olson, E.N. (2001). Activation of cardiac
gene expression by myocardin, a transcriptional cofactor for serum
response factor. Cell 105, 851–862.
Kong, Y., Shelton, J.M., Rothermel, B., Li, X., Richardson, J.A., Bassel-Duby, R., and Williams, R.S. (2001). Cardiac-specific LIM protein
FHL2 modifies the hypertrophic response to beta-adrenergic stimulation. Circulation 103, 2731–2738.
Wang, D.Z., Li, S., Hockemeyer, D., Sutherland, L., Wang, Z., Schratt,
G., Richardson, J.A., Nordheim, A., and Olson, E.N. (2002). Potentiation of serum response factor activity by a family of myocardinrelated transcription factors. Proc. Natl. Acad. Sci. USA 99, 14855–
14860.
Manabe, I., and Owens, G.K. (2001). CArG elements control smooth
muscle subtype-specific expression of smooth muscle myosin
in vivo. J. Clin. Invest. 107, 823–834.
McLoughlin, P., Ehler, E., Carlile, G., Licht, J.D., and Schafer, B.W.
(2002). The LIM-only protein DRAL/FHL2 interacts with and is a
corepressor for the promyelocytic leukemia zinc finger protein. J.
Biol. Chem. 277, 37045–37053.
Miano, J.M. (2003). Serum response factor: toggling between disparate programs of gene expression. J. Mol. Cell. Cardiol. 35, 577–593.
Miralles, F., Posern, G., Zaromytidou, A.I., and Treisman, R. (2003).
Actin dynamics control SRF activity by regulation of its coactivator
MAL. Cell 113, 329–342.
Morlon, A., and Sassone-Corsi, P. (2003). The LIM-only protein FHL2
is a serum-inducible transcriptional coactivator of AP-1. Proc. Natl.
Acad. Sci. USA 100, 3977–3982.
Müller, J.M., Isele, U., Metzger, E., Rempel, A., Moser, M., Pscherer,
A., Breyer, T., Holubarsch, C., Buettner, R., and Schüle, R. (2000).
FHL2, a novel tissue-specific coactivator of the androgen receptor.
EMBO J. 19, 359–369.
Müller, J.M., Metzger, E., Greschik, H., Bosserhoff, A.K., Mercep,
L., Buettner, R., and Schüle, R. (2002). The transcriptional coactivator FHL2 transmits Rho signals from the cell membrane into the
nucleus. EMBO J. 21, 736–748.
Oh, J., Wang, Z., Wang, D.Z., Lien, C.L., Xing, W., and Olson, E.N.
(2004). Target gene-specific modulation of myocardin activity by
GATA transcription factors. Mol. Cell. Biol. 24, 8519–8528.
Purcell, N.H., Darwis, D., Bueno, O.F., Müller, J.M., Schüle, R., and
Molkentin, J.D. (2004). Extracellular signal-regulated kinase 2 interacts with and is negatively regulated by the LIM-only protein FHL2
in cardiomyocytes. Mol. Cell. Biol. 24, 1081–1095.
Scholl, F.A., McLoughlin, P., Ehler, E., de Giovanni, C., and Schafer,
B.W. (2000). DRAL is a p53-responsive gene whose four and a half
LIM domain protein product induces apoptosis. J. Cell Biol. 151,
495–506.
Schratt, G., Weinhold, B., Lundberg, A.S., Schuck, S., Berger, J.,
Schwarz, H., Weinberg, R.A., Ruther, U., and Nordheim, A. (2001).
Serum response factor is required for immediate-early gene activation yet is dispensable for proliferation of embryonic stem cells.
Mol. Cell. Biol. 21, 2933–2943.
Schratt, G., Philippar, U., Berger, J., Schwarz, H., Heidenreich, O.,
and Nordheim, A. (2002). Serum response factor is crucial for actin
Wang, Z., Wang, D.Z., Hockemeyer, D., McAnally, J., Nordheim,
A., and Olson, E.N. (2004). Myocardin and ternary complex factors
compete for SRF to control smooth muscle gene expression. Nature
428, 185–189.
Weinhold, B., Schratt, G., Arsenian, S., Berger, J., Kamino, K.,
Schwarz, H., Rüther, U., and Nordheim, A. (2000). Srf(⫺/⫺) ES cells
display non-cell autonomous impairment in mesodermal differentiation. EMBO J. 19, 5835–5844.
Wixler, V., Geerts, D., Laplantine, E., Westhoff, D., Smyth, N., Aumailley, M., Sonnenberg, A., and Paulsson, M. (2000). The LIM-only
protein DRAL/FHL2 binds to the cytoplasmic domain of several
alpha and beta integrin chains and is recruited to adhesion complexes. J. Biol. Chem. 275, 33669–33678.
Accession Numbers
The GEO accession numbers for the U74A microarray data sets
reported in this paper are GSE1948 and GSE1949.