(Received for publication, May 6, 1997)
From the Department of Cell Biology, Baylor College of Medicine, Houston, Texas 77030
Serum response factor (SRF), a member of an
ancient family of DNA-binding proteins, is generally assumed to be a
ubiquitous transcription factor involved in regulating growth
factor-responsive genes. However, avian SRF was recently shown
(Croissant, J. D., Kim, J.-H., Eichele, G., Goering, L., Lough, J.,
Prywes, R., and Schwartz, R. J. (1996) Dev. Biol. 177, 250-264) to be preferentially expressed in myogenic lineages and is
required for regulating post-replicative muscle gene expression. Given
the central importance of SRF for the muscle tissue-restricted
expression of the striated -actin gene family, we wanted to
determine how SRF might contribute to this muscle-restricted
expression. Here we have characterized the murine SRF genomic locus,
which has seven exons interrupted by six introns, with the entire locus
spanning 11 kilobases. Murine SRF transcripts were processed to two
3
-untranslated region polyadenylation signals, yielding 4.5- and
2.5-kilobase mRNA species. Murine SRF mRNA levels were the
highest in adult skeletal and cardiac muscle, but barely detected in
liver, lung, and spleen tissues. During early mouse development,
in situ hybridization analysis revealed enrichment of SRF
transcripts in the myotomal portion of somites, the myocardium of the
heart, and the smooth muscle media of vessels of mouse embryos.
Likewise, murine SRF promoter activity was tissue-restricted, being
80-fold greater in primary skeletal myoblasts than in liver-derived HepG2 cells. In addition, SRF promoter activity increased 6-fold when
myoblasts withdrew from the cell cycle and fused into differentiated myotubes. A 310-base pair promoter fragment depended upon multiple intact serum response elements in combination with Sp1 sites for maximal myogenic restricted activity. Furthermore, cotransfected SRF
expression vector stimulated SRF promoter transcription, whereas dominant-negative SRF mutants blocked SRF promoter activity,
demonstrating a positive role for an SRF-dependent
autoregulatory loop.
Serum response factor (SRF),1 a member of an ancient family of DNA-binding proteins, contains a highly conserved DNA-binding/dimerization domain of 90 amino acids, termed the MADS box (1). The structure of the MADS box domain, recently elucidated by Pellegrini et al. (2), was assembled before the divergence of plants and animals. The identical MADS box structures were present in yeast transcription factors MCM1 and ARG80, a large number of homeotic like plant proteins, and invertebrate and vertebrate SRFs (reviewed in Ref. 3). All of these transcription factors, through their common MADS boxes, virtually bind to the same DNA sequences (1) and interact with similar kinds of co-accessory regulatory factors (reviewed in Refs. 4 and 5). Molecular dissection of human SRF revealed phosphorylation sites in the N-terminal domain that influence DNA binding, whereas sequences downstream of the MADS box contain the C-terminal transcription activation domain (6, 7). These extra MADS box sequences are well conserved in vertebrate SRF species, but have diverged from lower animal and plant species.
Earlier studies (8-10) have demonstrated that SRF-binding sites,
termed serum response elements (SRE; GG(A/T)6CC), play a primary role in regulating early response genes such as
c-fos and egr-1 (reviewed in Ref. 11). Growth
factor signaling through the c-fos SRE appeared to be
mediated through the formation of ternary complexes with an accessory
factor, p62TCF, and through protein-DNA interactions with a
purine-rich sequence at the 5-end of the c-fos SRE (12,
13). The ETS domain proteins Elk-1 (14) and SAP-1 (15) possess
biochemical activities that are characteristic of p62TCF
(16) and interact with the C-terminal portion of the MADS box (17).
Furthermore, another MADS box accessory factor, a paired-like homeodomain protein, Phox1 (18), has been shown to facilitate the DNA
binding activity of SRF on the c-fos SRE. In addition, Phox1
and Elk-1 potentiate the ability of SRF to transcriptionally activate
the c-fos promoter in response to growth-mediated events (18).
SRF plays an obligatory role in regulating post-replicative muscle gene
expression. The multiple SREs in the promoters of vertebrate striated
-actin genes are required for myogenic expression (19-22).
Site-directed mutagenesis of the SREs of the avian striated
-actin
promoters revealed that these SREs acted in combination with each other
and were necessary for transcription (22, 23). We (24) and others (7,
25) have shown that SREs are not equivalent in function due to the
contextural sequences embedding each SRE. For example, the skeletal
-actin proximal SRE allowed a minimal promoter to be activated
during muscle differentiation (26). Indeed, SRF could activate the
avian skeletal
-actin promoter in transient transfection experiments
by competing against a negative-acting YY1 factor for binding on the
proximal SRE (7, 25). Avian SRF mRNA and protein dramatically
increase as primary myoblasts withdraw from the cell cycle and fuse. In
addition, SRF becomes localized to the nucleus in differentiated
myotubes (27). Thus, the increase in SRF mRNA appeared prior to the
up-regulation of
-actin gene activity during myogenesis (28).
Vandromme et al. (29) demonstrated that microinjection of
SRF antibodies prevented the progression of myogenic differentiation,
implying an early dependence on SRF. In addition, Croissant et
al. (27) demonstrated that a dominant-negative SRF mutant,
defective in DNA binding but capable of heterodimerizing with other SRF
monomers, inhibited the transcriptional activity of the skeletal
-actin gene promoter in myogenic cultures and also blocked terminal
differentiation. Thus, SRF has a requisite role in
-actin gene
transcription during terminal skeletal muscle differentiation.
Recently, Spencer and Misra (30) demonstrated that a 322-bp promoter
region of the murine SRF gene was responsive to serum stimulation in
NIH3T3 fibroblasts and that the SREs and Sp1-binding sites present
within this promoter region were responsible. Since it is generally
assumed that SRF is a ubiquitous transcription factor, no attempt has
so far been made to understand the basis for the muscle tissue enriched
expression of SRF. Given the central importance of SRF for the muscle
tissue-restricted expression of the sarcomeric actin gene family, we
wanted to determine how SRF might contribute to this muscle-restricted
expression. Here we have characterized the SRF genomic locus. Murine
SRF gene activity was reminiscent of the expression pattern of another
MADS box-containing factor, MEF-2 (31), being primarily restricted to
cell types derived from embryonic mesoderm such as skeletal, cardiac,
and smooth muscles and, to a lesser extent, to cell types of
neuroectodermal origins. Also, SRF was virtually absent in
endoderm-derived tissues such as the liver, lung, and spleen. To
understand the mechanisms responsible for the tissue-regulated
expression of SRF, we also analyzed the cis-acting elements
in the SRF promoter region. Our results indicated the 310 bp of the SRF
promoter region upstream of the cap site as the 5-regulatory boundary
required for muscle-restricted expression. Furthermore,
dominant-negative SRF mutants blocked SRF promoter activity in muscle
cells. SRF gene activity appeared to be under an autoregulatory loop,
in which two high affinity SREs in the core promoter were required for
the SRF expression in skeletal muscle cells.
A fragment of mouse SRF
cDNA corresponding to nucleotides 834-1520 of human SRF (a gift
from Dr. E. Olson) was used to screen a mouse heart cDNA library
(Stratagene). One million plaques were initially screened with the
above-mentioned mouse SRF cDNA fragment as the probe, and five
clones were isolated. Restriction mapping and limited sequencing
indicated that four of these clones were identical. The PstI
fragment from the 3-end of one of these clones corresponding to
nucleotides 1747-1985 was used to rescreen the same library, and seven
more clones were isolated. The remaining 3
-untranslated region
sequences were isolated by screening 5 × 105 plaques
with the NheI-BglII fragment of the mouse SRF
genomic DNA, which corresponded to nucleotides 3354-4201 of human SRF cDNA. The mSRF cDNA restriction fragments were subcloned and
sequenced using the Sequenase Version 2.0 sequencing system (U. S.
Bioscience, Inc.).
The mouse strain 129 genomic library constructed in the EMBL2 -vector was a gift from Dr.
Philip Soriano. Three clones, each containing ~14 kbp of insert DNA,
were isolated by screening one million plaques with the mSRF cDNA
fragment corresponding to nucleotides 834-1520 of the human SRF
cDNA. Restriction mapping and Southern analysis of these clones
with various probes derived from the murine SRF cDNA indicated that
clone
-5 contained the complete SRF gene. Appropriate fragments from
this clone were subcloned and sequenced. The exon/intron borders were
assigned by aligning the cDNA and genomic DNA sequences.
Total RNA was isolated from
adult mouse tissues according to Chomczynski and Sacchi (32). Thirty
µg of total RNA was resolved on formaldehyde-containing 1% agarose
gel and then blotted onto GeneScreen membrane (NEN Life Science
Products). The prehybridization, hybridization, and washings were
according to the manufacturer's recommendations. The coding region
(nucleotides 939-1747) and the post I poly(A) region probes were
hybridized overnight in 50% formamide at 42 °C and washed at
65 °C. The 23-mer primer used for primer extension analysis was
complementary to nucleotides 82-104 of mSRF cDNA. The primer was
end-labeled with T4 polynucleotide kinase and
[-32P]dATP, and 100,000 cpm of the labeled primer was
hybridized overnight at 30 °C with 50 µg of skeletal muscle total
RNA in 80% formamide, 100 mM sodium citrate, pH 6.4, 300 mM sodium acetate, pH 6.4, and 1 mM EDTA. The
primer-RNA hybrid was precipitated and extended with 200 units of
SuperScript II reverse transcriptase (Life Technologies, Inc.) in 50 mM Tris-HCl, pH 8.3, 75 mM KCl, 3 mM MgCl2, 0.01 M dithiothreitol,
and 0.5 mM dNTPs for 1 h at 45 °C. The reaction was
treated with 1 µg of RNase A; extracted once with phenol/chloroform; and analyzed on 8 M urea, 6% denaturing polyacrylamide
gel. A dideoxy sequencing ladder generated using the same primer was used as size markers.
In
situ hybridization was performed on 7-mm sections of day 11.5 mouse embryos as described by Croissant et al. (27). A murine SRF probe, corresponding to the 3-untranslated region subcloned
into pBluescript, was linearized with the appropriate restriction
enzymes to produce antisense 35S-labeled copy riboprobes.
Sections were hybridized overnight at 58 °C and washed at 64 °C.
Sections were processed for emulsion autoradiography, post-stained with
Hoechst 33258, and visualized by epifluorescence and dark-field
microscopy.
The wild-type p456 plasmid
was constructed by ligating the PCR-amplified 479-bp fragment
containing 456-bp 5
- and 23-bp 3
-sequences relative to the
transcriptional start site of the SRF gene and inserted between the
NheI and XbaI sites of the luciferase reporter
plasmid pGL2. To construct plasmids p
310, p
292, and p
227, p
456
was digested with KpnI and briefly digested with Bal-31
exonuclease. After inactivation of Bal-31 exonuclease, the 5
-overhangs
were removed by T4 DNA polymerase digestion. The remaining DNA was
trimmed with EcoRI, which cuts once within the luciferase
vector. The SRF promoter region and part of the luciferase vector were
gel-purified and ligated into the
SmaI/EcoRI-digested fragment of pGL2. Plasmids
p
136, p
57, and p
37 were derived from intermediate constructs
containing a BglII site introduced by site-directed
mutagenesis into plasmid p
456. The smaller BglII fragments
from these intermediate plasmids containing 136, 57, and 37 bp of the
SRF promoter were cloned at the BglII site of pGL2-basic to
construct p
136, p
57, and p
37, respectively.
Site-directed mutations were introduced into the 456
background by PCR. Polymerase chain reactions were performed in a
50-µl volume of Pfu polymerase buffer (100 pmol each of
the primer pairs, 0.2 mM dNTPs, and 5 units of
Pfu polymerase (Stratagene)). The upstream and downstream
wild-type primers were designed with NheI and
XbaI sites, respectively, at their 5
-ends. Mutagenic
primers contained either BglII (for SRE mutation) or
EcoRI (for Sp1 site mutation). The conditions for PCR were
as follows: initial denaturation at 95 °C for 10 min and an
additional 30 cycles of denaturation at 95 °C for 1 min, annealing
at 55 °C for 1 min, and extension at 72 °C for 2 min. The final
extension was at 72 °C for 20 min. The products of the secondary PCR
containing the desired mutations were digested with NheI and
XbaI and ligated at the NheI site of pGL2. All
mutations were confirmed by sequencing.
Chicken embryo
primary skeletal myoblasts were isolated from day 11 embryonic breast
muscle tissue as described previously (33). After transfection, cells
were placed in minimum Eagle's medium containing 10% horse serum and
2% chicken embryo extract. CV1 and HepG2 cells were maintained in
Dulbecco's modified Eagle's medium containing 10% fetal bovine serum
both before and after transfection. Cells were transfected with 1 µg
of the indicated plasmid DNA along with 0.2 µg of
pCMV--galactosidase by LipofectAMINE (Life Technologies, Inc.)
according to the manufacturer's recommendations. Transactivation
assays were performed with SRF promoter-reporter constructs (1 µg
cotransfected with 150 ng of SRF expression vector or empty vector).
SRF promoter inhibition assays were performed with 150 ng of a SRF
dominant-negative mutant (pCGNSRFpm1 or pCGNSRF
C). Cells were
harvested 48 h post-transfection, and luciferase activity was
measured according to standard methods in a luminometer. Luciferase activity was normalized for transfection efficiency using the
-galactosidase values. For the promoter transactivation experiments, luciferase activity was normalized to the total protein.
Nuclear extracts were prepared according to Bohinski et al. (34). The protein concentration of extracts was estimated by the Bio-Rad protein assay reagent. Electrophoretic mobility shift assays used 5-10 µg of the nuclear extract prepared from myotubes. The nuclear extract was first incubated for 10 min at room temperature with 1 µg of poly(dG-dC) in 1 × binding buffer (50 mM NaCl, 20 mM HEPES-KOH, pH 7.5, 0.1 mM EDTA, 0.5 mM dithiothreitol, and 10% glycerol). Specific and nonspecific double-stranded oligonucleotides and antibodies were included in the reaction for competition and supershift assays, respectively. Subsequently, 0.01 pmol of the indicated end-labeled probe was added and incubated for a further 10 min. DNA-protein complexes were resolved on a 5% polyacrylamide gel cast and run in 0.5 × Tris/glycine buffer. The gel was dried and autoradiographed.
DNase I Footprinting of the SRF Gene PromoterThe SRF
promoter fragment from 253 to +23, relative to the transcriptional
start site, was end-labeled at +23 and used for DNase I footprinting.
One µg of poly(dI-dC) was incubated at room temperature for 5 min
without or with increasing amounts of GST-SRF in 20 µl of 1 × binding buffer. The reaction was incubated at room temperature for an
additional 10 min after adding 5 × 105 cpm of probe.
The MgCl2 and CaCl2 concentrations were
adjusted to 5 and 1 mM, respectively, and the probe was
digested with 0.004 units of pancreatic DNase I at room temperature for
90 s. The reaction was stopped by the addition of 100 µl of stop
buffer (100 mM NaCl, 100 mM Tris-HCl, pH 7.6, 15 mM EDTA, 0.375% SDS, 50 mg/ml sonicated salmon sperm
DNA, and 100 mg/ml proteinase K) and incubated at 37 °C for 20 min.
After phenol/chloroform extraction and ethanol precipitation, the
samples were dissolved in loading buffer. Approximately 20,000 cpm of
the denatured samples was resolved on urea-6% polyacrylamide gel,
dried, and autoradiographed.
A mouse 129 genomic library was screened with the human SRF
cDNA (35) corresponding to nucleotides 834-1520, overlapping the
conserved MADS box region. Three overlapping clones, each containing
~14 kbp of mouse genomic DNA, were isolated, and one of these clones
(SRF 5) contained the complete SRF locus. The SRF structural gene
extends over 10 kbp and consists of seven exons (Fig.
1A), encoding the SRF protein of 504 amino
acids (Figs. 1B and 2). The sequence at the
exon/intron borders conformed to the GT-AG consensus sequence as shown
in Table I (36). The size of the exons ranged from 77 to
848 bp. However, based on which of the two polyadenylation signal
sequences are used, the last exon was either 833 or 2337 bp in size.
The methionine start codon (ATG) was located in the first exon, 347 bp
downstream of the major cap site. The first exon contained 347 bp of
GC-rich 5
-untranslated region and 501 bp of the coding region. The
conserved MADS box region was split by the first intron and bordered by the second intron. The transcription activation domain was spread over
exons 4-7. The stop codon (TGA) was located in the seventh and last
exon.
|
Comparison of the mouse SRF amino acid sequence with the human, chicken, and Xenopus sequences revealed a very high degree of sequence conservation during evolution (Fig. 2). The 90-amino acid MADS box region (Fig. 2, boldface) was identical in all three vertebrate SRF species. Mouse SRF is more closely related to human SRF than to Xenopus SRF. The coding region of mSRF is 94 and 95% similar to the nucleic acid and amino acid sequences of human SRF, respectively. The length of the coding region was also conserved for these two mammalian species. However, in comparison with more ancient relatives, such as the Drosophila SRF pruned, homologous sequences were limited only to the MADS box. Examination of the N-terminal domain revealed the presence of a conserved 36-amino acid insert in the amino-terminal region in human and mouse SRFs, which was absent in the Xenopus SRF. A higher degree of sequence divergence between mammalian and amphibian SRF species was observed in the N-terminal domain as compared with the carboxyl-terminal domain, which is involved in transcriptional activation.
Characterization of the Murine SRF PromoterThe 5-cap site
was identified by primer extension analysis. A 23-nucleotide-long
antisense olignucleotide from the 5
-end of the murine cDNA, which
corresponded to nucleotides +82 to +104, was end-labeled, hybridized to
total mouse muscle RNA, and extended with reverse transcriptase.
Results of primer extension analysis indicated that the majority of the
SRF transcripts were initiated from the guanosine located 347 bp 5
of
the methionine start codon (Fig. 3A), which
we previously reported (37). The major cap site for mSRF transcripts
was 4 nucleotides downstream of that for human SRF (35). Several minor
transcriptional start sites were also detected downstream of the major
initiation site.
Inspection of the genomic sequences immediately upstream of the cap
site revealed a TATA box at positions 16 to
22 (Fig. 3B). The TATA box sequence also resembled the consensus
binding element for YY1 and a tissue-restricted transcription factor, MEF-2. There are two consensus SREs located at
42 and
62. In addition, there are two divergent SREs located at
142 and
222. Potential binding sites for other immediate-early gene products (AP-1,
Egr-1, and Ets-1) are identified. Overlapping the Egr-1 sites are
Sp1-binding sites. Potential binding sites for NF-Y, GATA factors, and
TEF-1 are also present within the SRF promoter (Fig.
3B).
We asked, how does SRF fulfill its role in
regulating striated -actin genes, and how might SRF contribute to
tissue-restricted expression? To investigate the expression pattern of
SRF, total RNA from various tissues was analyzed by Northern blot
analysis. As shown in Fig. 4A, a coding
region probe downstream of the MADS box detected SRF transcripts of 2.5 and 4.5 kilobases, which were abundantly expressed in mesoderm-derived
tissues such as skeletal and cardiac muscle and, to a lesser extent, in
neuroectoderm-derived brain tissue. Liver, spleen, lung, and kidney
tissues, which are derived from the endoderm, barely expressed SRF
mRNA (Fig. 4). Close examination of the mSRF genomic sequence
revealed the presence of two polyadenylation signal sequences, each
separated from the other by 1.5 kbp. Differential utilization of these
two polyadenylation signals for mRNA 3
-end formation could have
contributed to the size differences observed for the two SRF RNA
species. To examine this possibility, we used a probe that overlapped
the second polyadenylation region, which detected only the 4.5-kilobase
species, as shown in Fig. 4B. Thus, the two mSRF RNAs arose
from post-transcriptional processing of the two polyadenylation sites.
As observed for blots probed with the SRF coding region, SRF mRNA
was detected primarily in skeletal and cardiac muscle and brain, but
not in endoderm-derived tissues, reinforcing our conclusion that the
expression of SRF is tissue-restricted (33).
SRF Transcripts Appear Enriched in Embryonic Skeletal, Cardiac, and Vascular Smooth Muscle
During vertebrate embryogenesis,
expression of striated -actin transcripts serves as an early marker
for differentiation of cardiac, skeletal, and vascular smooth muscle
cell lineages. We asked if SRF mRNA expression patterns were also
locally restricted to early embryonic cardiac, skeletal, and smooth
muscle cell types. In sectioned day 11.5 mouse embryos, SRF transcripts
were seen in the neuroectoderm of the brain and the neural tube, but
were absent in the underlying notochord (Fig.
5A). Transverse sections revealed high levels
of SRF expression in the bulbus cordis and the right atrial portions of
the myocardium, as shown in Fig. 5B. SRF was also detected
at high levels in the myotomal portion of somites and in the emerging
smooth muscle cells surrounding the second branchial arch artery (Fig.
5A). Lower levels of SRF was also detected in the
sympathetic trunk (Fig. 5B) and in the cardinal vein. SRF
was barely detected in the lung bud and liver (Fig. 5B).
Sense probes in all cases showed background levels of hybridization in
all tissues (data not shown). These in situ hybridization
experiments demonstrated that SRF gene expression was developmentally
regulated and largely restricted to the cardiac and skeletal muscle
cell lineages, consistent with the early specific expression of the
-actin genes in the embryo.
Tissue-regulated Expression of SRF
To investigate the
molecular basis for the muscle tissue enriched expression and to map
the essential cis-acting elements, we transiently
transfected chicken primary myotubes and the human liver cancer cell
line HepG with SRF promoter constructs. The p456 plasmid was 80-fold
more active in myotubes than in HepG2 cells (Fig. 6). A
further deletion to
310 uncovered the presence of a negative-acting
element(s). This deletion of 146 bp resulted in a nearly 2.2-fold
increase in the promoter activity in myotubes. The p
310 construct was
96-fold more active in myotubes than in HepG2 cells. A deletion of 18 bp to
292 decreased the promoter activity only slightly in myotubes
and by 56% in HepG2 cells. A further deletion to
227 decreased the
promoter activity by ~60% in myotubes, but not in HepG2 cells.
Additional muscle-specific positive-acting element(s) were revealed by
deletion to
136, which resulted in a 4-fold decrease in the promoter
activity in myotubes and a slight decrease in HepG2 cells. Two
non-consensus SREs are present within this deleted 91-bp region. A
further deletion of 79 bp to
57, which removed one of the two
consensus SREs (SRE2) and overlapping Egr-1- and Sp1-binding sites,
reduced the promoter activity by 12-fold in myotubes. Plasmid p
57,
which contains a single consensus SRE (SRE1) and the TATA box region,
was 25-fold more active than the promoterless control plasmid pGL2 in
myotubes. The p
57 construct was not active in HepG2 cells, which
express only low levels of endogenous SRF. A deletion of 20 bp to
37, which eliminated SRE1, reduced the promoter activity to background levels in myotubes, suggesting that the SREs were required for SRF
promoter activity.
Multiple SRF-binding Sites in the SRF Promoter
Results of
promoter deletion analysis suggested that the SREs present within the
SRF promoter were required for transcriptional activity. DNase I
protection assays performed with bacterially expressed purified SRF
were used to ascertain if the two consensus SREs present in the SRF
promoter bind SRF. Both SRE1 and SRE2 were well protected at the lowest
levels of GST-SRF. In addition, the TATA box region was also protected,
but at considerably higher inputs of GST-SRF (Fig. 7).
No protection was observed over the two potential, but non-consensus
SREs located at positions 142 and
222.
SREs may also serve as binding targets for YY1, NFIL-6, SRE-ZBP,
SRE-BP, and several other uncharacterized factors. Many of these
factors are also expressed in skeletal muscle tissue (Refs. 7 and 38;
reviewed in Ref. 11). Having demonstrated specific binding of
bacterially expressed SRF, we investigated the interaction of proteins
from myotube nuclear extracts with SREs from the SRF promoter.
Double-stranded oligonucleotide probes corresponding to SRE1 and SRE2
were incubated with the nuclear extract prepared from chicken embryo
myotubes. A doublet of slowly migrating complexes (complexes I and II)
and a fast migrating complex (complex III) were observed with the SRE2
probe (Fig. 8A). Nuclear extracts from the
fibroblast cell line NIH3T3 also gave rise to a similar doublet of SRF
complexes when a longer probe was used for gel shift assays (30).
Complexes I and II were competed by a 50-fold molar excess of cardiac
SRE1 and wild-type SRE1 and SRE2 oligonucleotides from the SRF
promoter, but not by nonspecific Sp1/Egr-1 and mutant SRE1 and SRE2
oligonucleotides (Fig. 8A) (data not shown). Furthermore, complexes I and II were either supershifted or abolished by SRF antiserum, but not by YY1 antiserum, indicating that these two complexes contain SRF (Fig. 8A) (data not shown). Complex
III was identified as a YY1-containing complex based on several
criteria. First, this complex was abundant in extracts from replicating myoblasts and decreased in myotube extracts as myogenesis progressed (data not shown). Second, this complex was competed by SRE2 and the
YY1-binding site-containing skeletal actin SRE1 oligonucleotide, but
not by SRE1 and cardiac actin SRE1 oligonucleotides, which do not bind
YY1 (Fig. 8A) (data not shown). Third, complex III was
abolished by YY1 antiserum and unaffected by SRF antiserum. SRE1 was
similar to SRE2 with respect to SRF binding, except that it did not
bind YY1 (Fig. 8A). We have also examined the binding of SRF
to noncanonical SREs (SRE3 and SRE4) by competition gel shift assays in
which SRE1 was the probe. Both SRE3 and SRE4 oligonucleotides competed
for SRF binding to the SRE1 probe, albeit poorly (data not shown).
Mutations at Both (but Not Individual) SREs Block Promoter Activity
To investigate the role of SRE1 and SRE2 in the muscle
enriched expression of SRF, site-directed mutational analysis of these SREs was performed in the context of the p456 promoter. The
BglII restriction site, which prevented the binding of SRF,
was substituted for these SREs (Fig. 8A). Mutation of SRE1
did not affect the activity of the promoter, suggesting that the SREs
may have partially redundant functions as SRE2 could functionally
substitute for mutated SRE1 (Fig. 8C). In contrast to the
SRE1 mutation, the SRE2 mutation increased the activity of the promoter
by 2.2-fold (Fig. 8C). However, mutation of both SRE1 and
SRE2 resulted in ~75% reduction in the promoter activity, suggesting
that either of the two SREs is required for the complete promoter
activity in myotubes.
In
the SRF promoter, there were two potential Sp1 sites located at 86 bp
(proximal Sp1) and
251 bp (distal Sp1). Overlapping these Sp1 sites
were potential binding sites for the zinc finger-containing transcription factor Egr-1. Sp1 sites from several other promoters contain similarly overlapping Egr-1 sites and bind both Sp1 and Egr-1.
We tested if the Sp1 sites from the SRF promoter could bind both Sp1
and Egr-1 present in myotube nuclear extracts. Three closely migrating
complexes were seen with the distal Sp1 site probe (Fig.
8B). These complexes were specifically competed by a 50-fold
molar excess of unlabeled consensus Sp1 site and proximal and distal
wild-type Sp1 sites, but not mutated Sp1 sites. The identity of these
complexes was confirmed by an antibody supershifting experiment. The
complexes were supershifted by the addition of Sp1 antibody, but not by
Egr-1 antibody, indicating that the complexes contain Sp1 and not
Egr-1. Even though the proximal Sp1 site contains an overlapping
consensus Egr-1 site, binding of only Sp1 was evident under our
electrophoretic mobility shift assay conditions (data not shown).
Spencer and Misra (30) showed by mutational analysis that the two Sp1 sites present in the murine SRF promoter were essential for serum induction of the promoter in 3T3 fibroblasts, but mutated Sp1 sites did not affect the basal promoter activity. The role of these Sp1 sites in the muscle enriched expression of the SRF promoter was also examined by mutagenesis. Sp1 site-directed mutations abolished the binding of Sp1 from myotube nuclear extracts (Fig. 8B). Mutagenesis of the proximal Sp1 site actually increased the promoter activity by 2-fold (Fig. 8D), whereas a mutation over the distal Sp1 site reduced the promoter activity by 60%.
Myogenic Up-regulation of SRF Promoter ActivityRecently,
avian SRF RNA and protein and SRF DNA binding activity were shown to
increase when cultured primary myoblasts were allowed to withdraw from
the cell cycle and fuse to form multinucleated myotubes (7, 27). We
wanted to determine if the up-regulation of SRF gene activity was under
transcriptional control by examining the activity of transfected SRF
promoter-reporter constructs in primary chicken embryo myogenic
cultures. The SRF promoter fragment from 456 to +23 linked to a
luciferase reporter gene displayed low activity in replicating
pre-fusion myoblasts, but was up-regulated ~6-fold in late-stage
myotubes (Fig. 9A). Thus,
cis-acting sequences required for the up-regulation of SRF
gene activity are contained within the 456-bp promoter region.
The up-regulation of SRF during myogenesis, the presence of multiple
SREs in the SRF promoter, and the high affinity binding of SRF to these
SREs suggested the possibility that SRF autoregulates itself. Thus, we
compared luciferase reporter activities of the p456 SRF promoter
construct cotransfected with or without SRF expression vectors in
non-myogenic CV1 fibroblasts. Coexpression of an exogenous SRF resulted
in up to a 5-fold increase in SRF promoter activity (Fig.
9C). We then asked if dominant-negative SRF mutants would
block SRF promoter function in transfected chicken embryo myotubes. The
SRFpm1 mutant (39) dimerizes with other SRF monomers and interferes
with wild-type SRF by forming DNA binding-defective heterodimers. In
addition, another dominant-negative SRF mutant, SRF
C, in which the
C-terminal transcription activation domain (amino acids 266-504) was
deleted, acts as a de facto repressor by occupying SREs
through specific DNA binding, but is incapable of activating
SRE-dependent transcription. Cotransfection of either SRFpm1 or SRF
C with p
456 resulted in 40 and 95% decreases in SRF
promoter activity, respectively, thus suggesting that the myogenic
up-regulation of SRF promoter activity was mediated by SRF (Fig.
9B). Inhibition by SRFpm1 and SRF
C was specific to the
SRF promoter because SV40 promoter activity was not significantly affected by these dominant-negative SRF mutants. These results indicate
that SRF autoregulates its own promoter and that this autoregulation is
primarily mediated through SRE1 and SRE2.
The hypothesis that introns and RNA splicing facilitated the evolution of ancient genes in the progenote organism was recently reviewed (40). The function of introns in the evolution of genes can be explained by the proposal that either introns appeared late in evolution and could not participate in the construction of primordial genes or that RNA splicing and introns existed in the earliest organisms, but were lost during the evolution of the modern prokaryotes. Blake (41) suggested that evidence for intron-facilitated evolution of a gene might be found by comparing the borders of functional protein domains with the placement of introns. The recent elucidation of the x-ray crystal structure of the SRF MADS box demonstrated a novel DNA-binding motif, a coiled-coil, and a stratified structural subdomain involved in dimerization (2). We showed here (Fig. 1 and Table I) that the murine SRF gene consists of seven exons interrupted by six introns. Exon/intron borders are well conserved between the Xenopus SRF (42) and murine SRF genes. The first intron was found to sever the N-terminal extension, which makes specific base contacts within the minor grove of an SRE half-site, from the dimerization subdomain encoded in exon II. In comparison with the genomic organization of MEF-2B (43), which was conserved in Drosophila d-mef-2, other MEF-2 relatives, and the plant AGL3 gene (44), the first intron also bisected the unstructured N-terminal extension, whereas the second intron was close to the C-terminal border of MADS boxes found in animal and plant SRFs and the MADS/MEF-2 boxes in all MEF-2 genes. Thus, in all MADS box-containing genes yet examined, introns closely circumscribed the dimerization subdomain. Based on conservation of primary sequence of the MADS box region and gene organization analysis, introns might have participated in the construction of the earliest MADS box-containing genes, prior to the diversion of plants and animals that occurred at least one billion years ago.
Despite these similarities, MADS box proteins also have evolved to perform diverse functions such as specification of mating type in yeast, homeotic activities in plants, pulmonary system development in Drosophila, and elaboration of mesodermal structures in vertebrates. Interestingly, the overall structural divergence of SRF proteins among evolutionarily distant species of animals appears to be related to differences in the spatial expression pattern. For example, of the variety of animal species examined, pruned, the SRF homolog of Drosophila SRF (45), was the most divergent, in which sequence conservation was limited only to the MADS box domain. The localization of Drosophila SRF expression was different from that of Xenopus, avian, and murine SRFs, which are more unified in structure and tissue expression. Drosophila SRF was localized to the insect tracheal system (46), whereas vertebrate SRFs, like several of their MEF-2 counterparts (reviewed in Ref. 5), were localized to avian and murine skeletal and cardiac muscle and neuroectoderm-derived tissues (Figs. 4 and 5) (27).
How does SRF play a central role in regulating muscle-specific genes
that are expressed under cell differentiation-promoting conditions? We
have shown that mSRF has a distinct striated muscle tissue enriched
expression pattern and further identified the SRE as a mediator of SRF
promoter regulation. Although it is generally assumed that SRF serves a
role as a constitutive factor during its association with accessory
factors, we have shown that SRF binding activity actually increased
dramatically following the ending of cell replication primarily due to
change in the cellular content of SRF in primary myoblasts (7). Surveys
of early avian (27) and murine (Fig. 5) embryos also indicated
tissue-restricted expression of SRF transcripts, which substantially
increased the cellular mass of SRF in the myotomal portion of somites,
cardiac myocytes, and vascular smooth muscle cells. The expression of SRF continues in these tissues at high levels through adulthood (Fig.
4). During primary myogenesis in culture, SRF promoter activity, RNA,
and protein mass increase significantly preceding fusion of myoblasts
and the appearance of muscle-specific structural genes (33). We showed
that mutations of both SRE1 and SRE2 were required to down-regulate SRF
promoter activity, suggesting that SRE1 and SRE2 were functionally
redundant (Fig. 8C). Overexpression of SRF from a plasmid
vector substantially increased SRF promoter activity in transfected
fibroblasts (Fig. 9C). In comparison, the dominant-negative
mutants of SRF, SRFpm1 and SRFC, inhibited the muscle enriched
expression of SRF and other muscle-specific genes.2 Thus, SRF has a primary role in
directing muscle differentiation.
Tissue-restricted expression of SRF was also evident from comparison of its promoter activity, in which the SRF promoter activity was at least 2 orders of magnitude greater in primary cultured myotubes than in liver HepG2 cells. Low SRF promoter activity cannot be attributed to the presence of strong liver tissue-specific silencer elements in the SRF promoter because serial deletions in the promoter did not activate the promoter. Although less likely, it is possible that a silencer element located within the 37-bp cap upstream region strongly repressed the SRF activity in liver cells. Another possibility for the lack of SRF promoter activity in liver cells could be the absence of SRF promoter-specific trans-acting factors from this cell type. Endoderm-derived liver tissue did not express endogenous SRF (Fig. 4). Furthermore, a minimal SRF promoter containing a single SRE, which was otherwise active in myotubes, was not active in HepG2 cells, indicating that the SRF promoter activity was SRF-dependent. The SREs were required for the higher basal level of expression in myotubes (Fig. 8C).
The basal activity of the SRF promoter in NIH3T3 cells was shown to be
dependent on the more ubiquitous CAAT box-binding factor, whereas the
SREs and Sp1 sites were dispensable (30). In contrast, SRE1 and Sp1
sites were required for both serum-induced promoter activity (30) and
muscle tissue-restricted activity. SRF promoter deletion analysis
demonstrated that SRF was necessary but not totally sufficient for
driving the SRF promoter. Furthermore, autoregulation of the SRF
promoter by SRF alone cannot account for the muscle tissue enriched
expression of the SRF gene. The tissue enriched expression of SRF might
be accomplished by the concerted action of SREs with other
cis-acting elements. Accordingly, deletion of sequences 5
to SRE1 and SRE2 decreased the promoter activity, suggesting that the
interaction of these deleted sequences with SRE1 and SRE2 may be
required for the complete activity of the promoter. Of the two
Sp1-binding sites present in the SRF promoter, only the distal Sp1 site
appeared to be important for SRF expression. Sartorelli et
al. (47) have shown that a functional interaction among SRF, the
ubiquitous transcription factor Sp1, and the cell type-restricted
myogenic factor MyoD is required for human cardiac
-actin gene
expression.
One mechanism by which SRF might promote muscle-specific gene
expression would be by interfering with the activity of negative regulators of muscle differentiation. YY1, a ubiquitously expressed C2H2 zinc-finger protein (48, 49) that binds
the consensus sequence AANATGGNG, has been shown to bind several SREs
(7, 50). We observed that in proliferating myoblasts, the skeletal -actin gene was repressed by mutually exclusive binding of YY1 over
SRE1 (7). Interestingly, gel mobility shift assays with SRE2 from the
SRF promoter also uncovered an overlapping YY1 site (Fig.
8A). Furthermore, comparison of nuclear extracts prepared from proliferating myoblasts with myotubes revealed mutually exclusive binding of SRF and YY1 over SRE2 (data not shown). Consistent with the
repressor role of YY1, mutation of SRE2 resulted in a 2.2-fold increase
in SRF promoter activity (Fig. 8C), indicating that the SRF
promoter activity might also be modulated by YY1. Displacement of YY1
by increased SRF binding activity during myogenesis may facilitate the
SRF autoregulatory loop.
The combinatorial interaction of SRF with tissue-specific accessory
factors might be another way to confer tissue specificity to SRF
(reviewed in Refs. 51 and 52). A cardiac-specific homeodomain protein,
Nkx-2.5, was shown to be expressed with a spatio-temporal pattern
similar to that observed for avian SRF. Physical interaction of SRF
with Nkx-2.5 resulted in enhanced expression of both endogenous and
transfected chicken cardiac -actin genes (52). Building up of
SRF·Nkx-2.5 complexes might compete off negative-acting factors such
as YY1 and allow for saturation of the multiple SREs with
positive-acting SRF complexes. Conversely, SRF·Nkx-2.5 complexes in
cardiac myocytes might serve to repress the c-fos promoter through forming nonproductive complexes via its SRE. Therefore, SRF
might be able to mediate accessory factor interactions with certain
homeodomain factors that either activate or repress transcription.
Likewise, interactions of SRF with skeletal muscle-restricted basic
helix-loop-helix proteins of the MyoD family may also confer skeletal
muscle specificity to SRF (53). Although consensus E box-binding sites
for basic helix-loop-helix proteins are absent in the SRF promoter, it
is still conceivable that the SRF promoter could be activated by
myogenic regulatory factors in an E box-independent manner, as has been
demonstrated for the chicken myoD promoter (54). The
mechanism of imparting muscle tissue specificity to SRF is not limited
to physical interaction between SRF and a tissue-restricted transfactor. Functional interaction of SRF and MyoD or myogenin bound
to different sites on the promoter can confer muscle tissue specificity
to SRF (47). A similar interaction of SRF on the interleukin-2 receptor
-chain gene promoter with the Rel homology protein NF-
B confers
T-cell specificity to SRF (55, 56). Thus, additional interactions of
SRF with different cell type-restricted coactivators may also determine
the response of different tissues to SRF.
The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EMBL Data Bank with accession number(s) U49759.
We thank Lisa Goering and Dr. Ruxandra Draghia for assistance in preparing the in situ hybridization figure.