Differential Binding of an SRF/NK-2/MEF2 Transcription Factor ...

THE JOURNAL OF BIOLOGICAL CHEMISTRY © 2001 by The American Society for Biochemistry and Molecular Biology, Inc.

Vol. 276, No. 37, Issue of September 14, pp. 34637–34650, 2001 Printed in U.S.A.

Differential Binding of an SRF/NK-2/MEF2 Transcription Factor Complex in Normal Versus Neoplastic Smooth Muscle Tissues* Received for publication, June 22, 2001 Published, JBC Papers in Press, July 16, 2001, DOI 10.1074/jbc.M105826200

Christopher J. Phiel‡, Vijayalakshmi Gabbeta‡, Linda M. Parsons§, David Rothblat‡, Richard P. Harvey¶储, and Kirk M. McHugh‡** From the ‡Department of Pathology, Anatomy, and Cell Biology, Thomas Jefferson University, Philadelphia, Pennsylvania 19107, the §Department of Anatomy and Cell Biology, University of Melbourne, Parkville 3054, Victoria, Australia, ¶The Victor Chang Cardiac Research Institute, St. Vincent’s Hospital, Darlinghurst 2010, Australia, and the 储University of New South Wales, Kensington 2052, Australia

The malignant potential of smooth muscle tumors correlates strongly with the disappearance of ␥-smooth muscle isoactin, a lineage-specific marker of smooth muscle development. In this paper, we identify a 36-base pair regulatory motif containing an AT-rich domain, CArG box, and a non-canonical NK-2 homeodomainbinding site that has the capacity to regulate smooth muscle-specific gene expression in cultured intestinal smooth muscle cells. Serum-response factor associates with an NK-2 transcription factor via protein-protein interactions and binds to the core CArG box element. Our studies suggest that the NK-2 transcription factor that associates with serum-response factor during smooth muscle differentiation is Nkx2-3. Myocyte-specific enhancer factor 2 binding to this regulatory complex was also observed but limited to uterine smooth muscle tissues. Smooth muscle neoplasms displayed altered transcription factor binding when compared with normal myometrium. Differential nuclear accessibility of serum-response factor protein during smooth muscle differentiation and neoplastic transformation was also observed. Thus, we have identified a unique regulatory complex whose differential binding properties and nuclear accessibility are associated with modulating ␥-smooth muscle isoactin-specific gene expression in both normal and neoplastic tissues.

Smooth muscle neoplasms are a common tumor occurring in 25% of women (1). The distribution of smooth muscle tumors roughly parallels the concentration of smooth muscle tissues in the body with the majority of all smooth muscle tumors arising in the female genital tract. Smooth muscle tumors represent a major health concern being associated with infertility, significant morbidity, and ⬃1/3 of all hysterectomies performed in the United States (1). Recent evidence also suggests a rising incidence of smooth muscle tumors in immunodeficiency states and/or virally infected individuals (2, 3). A better knowledge of * This work was supported by the Cooperative Human Tissue Network, which is funded by NCI, National Institutes of Health and by National Institutes of Health Grants HD27252 and DK55791. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EBI Data Bank with accession number(s) AF040977. ** To whom correspondence should be addressed: Dept. of Pathology, Anatomy, and Cell Biology, Thomas Jefferson University, 1020 Locust St., Philadelphia, PA 19107. Tel.: 215-503-7835; Fax: 215-923-3808; E-mail: [email protected]. This paper is available on line at http://www.jbc.org

how smooth muscle tumors arise and their relationship to normal smooth muscle development could lead to improved diagnosis, treatment, and the prevention of unnecessary surgical procedures. The factors responsible for controlling smooth muscle development and its resulting neoplastic transformation have yet to be identified. To begin to elucidate the molecular mechanisms responsible for these critical processes, we took advantage of a well defined in vitro model of smooth muscle differentiation (4). Primary cultures of rat intestinal smooth muscle cells (ISMCs)1 express markers of smooth muscle determination and differentiation in a manner similar to that seen in vivo. One marker of differentiation, ␥-smooth muscle isoactin, is expressed in differentiated smooth muscle cells from early embryonic development into adulthood (5, 6). Additionally, the absence of ␥-smooth muscle isoactin expression correlates strongly with the malignant potential of smooth muscle tumors (7–9). Therefore, ␥-smooth muscle isoactin represents an excellent model for understanding the events leading to smooth muscle-specific gene expression in both normal and neoplastic tissues. We have cloned and analyzed the promoter region of ␥-smooth muscle isoactin. Deletion analysis identified a corebinding site for a transcription factor complex that has the capacity to regulate smooth muscle-specific gene expression. This transcriptional complex differentially binds serum-response factor (SRF), myocyte enhancer factor 2 (MEF2), and an NK-2 homeobox protein in both a developmental and tissuespecific manner. In addition, uterine smooth muscle neoplasms showed altered transcription factor binding when compared with normal myometrium. Differential nuclear accessibility of SRF protein was also observed during smooth muscle development and following the neoplastic transformation of uterine smooth muscle cells. This study represents the first identification of an endogenous in vivo transcriptional complex necessary for the expression of a marker of smooth muscle differentiation in mammalian cells, and suggests that smooth muscle cell pathologies may be associated with an alteration of this complex. EXPERIMENTAL PROCEDURES

Cloning and Reporter Assays—A P1 bacteriophage rat genomic clone containing the 5⬘ promoter region of ␥-smooth muscle isoactin was 1 The abbreviations used are: ISMC, intestinal smooth muscle cells; SRF, serum-response factor; rSRF, recombinant SRF; MEF, myocytespecific enhancer factor 2; EMSA, electrophoretic mobility shift assay; ACH, AT-rich/CArG box 1/homeodomain-binding site; CArG, CArG box; HDBS, homeodomain-binding site; pLuc, plasmid luciferase reporter vector; MADS, MCM1, Agamous, Deficiens, SRF; TSC, tuberous sclerosis; PBS, phosphate-buffered saline; RT, room temperature; bp, base pair; NMC, neonatal smooth muscle myocytes; NE, nuclear extract.

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Regulation of Smooth Muscle Differentiation

FIG. 1. Cellular phenotypes observed during ISMC development. Time line showing approximate stage where each rat ISMC phenotype first becomes evident during development. The ability of ISMC to dedifferentiate is reflected by the reverse arrows. Abbreviations include embryonic day 9 (E9), 11 (E11), and 13 (E13), neonatal day 3 (NEO3), and adult (AD).

FIG. 2. CArG box 2 is essential for ␥-smooth muscle isoactin expression. A, reporter gene assays of ␥-smooth muscle isoactin promoter activity. Diagram shows deletion mutants of the rat ␥-smooth muscle isoactin promoter including the relative position of putative cis-acting elements and exon 1. Negative numbers represent the 5⬘ end of the various deletion constructs, and positive 61 represents the 3⬘ end of the construct. The complete nucleotide sequence includes 744-bp of proximal promoter, exon 1, and 34-bp of intron 1. GenBankTM accession number for this sequence is AF040977. Mutagenized CArG box 1 is shown as a hatched box. Luciferase reporter constructs based upon the ␥-smooth muscle isoactin promoter were cotransfected into cultured rat ISMCs with a constitutively expressed control construct for normalization. The horizontal bars show the average activity (n ⫽ 3) and standard deviation for each construct after normalization. B, alignment of the proximal promoter sequences surrounding CArG box 2 of the rat, mouse (80), human (81), and chicken (82) is shown. CArG box 2 is 100% conserved between all species. The HDBS domain is 100% conserved between rat, mouse, and human and 63% conserved in chickens. The AT-rich domain is 100% conserved between rat and mouse, 86% conserved between rat and humans, and 71% conserved between rat and chicken. obtained from Genome Systems. A 4.3-kilobase pair EcoRI fragment of the promoter was subcloned into pBluescript. The clone was completely sequenced and is deposited in GenBankTM (accession number AF040977). Polymerase chain reaction was employed to create the various promoter fragments cloned into pGL2-Basic (Promega). Sitedirected mutagenesis was performed using the QuikChange Site-directed Mutagenesis Kit (Stratagene). All clones were confirmed by DNA sequencing. Culture and Transfection of Rat ISMCs—The isolation and maintenance of primary cultures of rat neonatal ISMCs have been described in detail elsewhere (4). Cultured ISMCs were examined at low density (day 1 of culture, smooth muscle myoblasts) and at confluency (day 10 of culture, immature smooth muscle myocytes). In addition, mature smooth muscle myocytes were isolated from both neonatal (post-coital days 3–5) and adult rats. Primary cultures of cardiomyocytes were established from neonatal rat hearts and grown under identical culture conditions as ISMCs. NIH3T3 fibroblasts were obtained from ATCC and grown under identical culture conditions as ISMCs. ISMCs to be transfected were plated at a density of 5 ⫻ 105 cells/ 35-mm plate 24 h prior to transfection. A calcium phosphate-DNA precipitate for each condition was prepared using the Calcium Phosphate Transfection System as outlined by the supplier (Life Technologies, Inc.). Calcium phosphate-DNA precipitates were applied to cultured ISMCs for 4 h, followed by a 3-min glycerol shock. 30 ␮g of reporter DNA was used for all transfections. 0.1 ␮g of pRL-SV40 Renilla (Promega) was included as an internal control for transfection efficiency. After 48 h, cells lysates were collected, and dual-luciferase assays were performed in a TD-20/20 luminometer (Turner Designs) as described by the supplier (Promega). Electrophoretic Mobility Shift Assay (EMSA)—A total of 3 normal

human myometrium, 12 human leiomyomas, and 5 human leiomyosarcomas as determined by pathologic diagnosis were collected from surgical resections, immediately frozen in liquid nitrogen, and stored at ⫺70 °C. Nuclear and cytoplasmic proteins were isolated from each sample by standard techniques. Briefly, 5–10 plates of cultured cells were serially scraped using 1 ml of 1⫻ phosphate-buffered saline (PBS). Tubes were spun at 15,000 ⫻ g for 15 s at 4 °C, and the PBS was removed. For nuclear proteins, the pellet was resuspended in 400 ␮l of cold buffer containing 10 mM HEPES, pH 7.9, 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM dithiothreitol, and 0.5 mM phenylmethylsulfonyl fluoride (Buffer A). Cells were allowed to swell on ice for 15 min. 25 ␮l of IGEPAL (Sigma) was added, and the tube was mixed vigorously for 30 s. The homogenate was then centrifuged for 30 s, 4 °C, at 15,000 ⫻ g. The supernatant was removed, and the nuclear pellet was resuspended in 50 ␮l of ice-cold buffer containing 20 mM HEPES, pH 7.9, 500 mM KCl, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, and 0.5 mM phenylmethylsulfonyl fluoride. Tubes were rocked vigorously at 4 °C for 15 min. Extracts were centrifuged at 15,000 ⫻ g for 5 min at 4 °C, and the supernatant was removed and frozen in aliquots at ⫺70 °C. Tissue samples were handled in the same manner after an initial mincing and homogenization in 1⫻ PBS. For cytoplasmic proteins, the initial pellet was resuspended in M-Per Mammalian Protein Extraction Reagent (Pierce) with 100⫻ Protease Inhibitor Mixture (Sigma). Tube was placed on ice for 30 min and then centrifuged at 15,000 ⫻ g for 30 min at 4 °C. Supernatant was removed and frozen in aliquots at ⫺70 °C. All proteins were quantitated spectrophotometrically using the Bradford assay (Bio-Rad). Double-stranded oligonucleotide probes were commercially synthesized (Bio-Synthesis, Inc) and included the 36-bp AT-rich/CArG box 2/HBDS (ACH) element (5⬘-GGATCTTTATTAAAAAAAACCCACCT-


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FIG. 3. Distinct smooth muscle cell phenotypes show unique nuclear protein binding patterns. Representative EMSA analysis of the 36-bp ACH ␥-smooth muscle isoactin promoter fragment. Labeled ACH was incubated with 10 ␮g of rat nuclear proteins isolated from smooth muscle myoblasts (lanes 1– 8), immature smooth muscle myocytes (lanes 9 –16), and neonatal (lanes 17–24) and adult mature smooth muscle myocytes (lanes 25–32). Lanes 1, 9, 17, and 25, nuclear extract (NE) ⫹ labeled ACH; lanes 2, 10, 18, and 26, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled ACH; lanes 3, 11, 19, and 27, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled AT-rich domain; lanes 4, 12, 20, and 28, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled CArG box 2; lanes 5, 13, 21, and 29, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled HDBS; lanes 6, 14, 22, and 30, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled AT-rich/CArG box 2; lanes 7, 15, 23, and 31, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled CArG box 2/HDBS; and lanes 8, 16, 24, and 32 NE ⫹ labeled ACH 100-fold excess unlabeled nonspecific (NS) DNA. Primary bands are identified alphabetically.

FIG. 4. SRF and an NK-2 protein form a transcriptional complex centered on CArG box 2. Representative EMSA analysis of the 36-bp ACH ␥-smooth muscle isoactin promoter fragment using antibodies specific for SRF (Santa Cruz Biotechnology, Inc.), MEF2A (11), MEF2B (12), MEF2D (12), and the TN domain of NK-2 proteins. Labeled ACH was incubated with 10 ␮g of rat nuclear proteins isolated from smooth muscle myoblasts (lanes 1– 6), immature smooth muscle myocytes (lanes 7–12), and neonatal (lanes 13–18) and adult mature smooth muscle myocytes (lanes 19 –24). Lanes 1, 7, 13, and 19, NE ⫹ labeled ACH; lanes 2, 8, 14, and 20, NE ⫹ labeled ACH ⫹ SRF antibody; lanes 3, 9, 15, and 21, NE ⫹ labeled ACH ⫹ MEF2A antibody; lanes 4, 10, 16, and 22, NE ⫹ labeled ACH ⫹ MEF2B antibody; lanes 5, 11, 17, and 23, NE ⫹ labeled ACH ⫹ MEF2D antibody; and lanes 6, 12, 18, and 24, NE ⫹ labeled ACH ⫹ TN domain antibody. Primary bands are identified alphabetically in series with reference to previous figures (i.e. band A is the same in all figures). The position of the SRF supershifted band is marked S1, and the TN domain supershifted band S2. Although MEF2A, MEF2B, and MEF2D antibodies produced no apparent supershifted bands, inclusion of these antibodies with neonatal smooth muscle myocyte nuclear proteins appeared to intensify band B. No differences in banding patterns were observed whether individual antibodies were added prior to or after the addition of labeled probe.

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FIG. 5. Individual elements comprising the ACH display unique nuclear protein binding patterns. Representative EMSA analysis of CArG box 2 (lanes 1–5), AT-rich domain (lanes 6 –13), HDBS (lanes 14 –19), consensus oligonucleotide homeodomain-binding site 5⬘-TCGGGATCGCCCAGTCAAGTGC-3⬘ (Nkx2-1, lanes 20 and 21), and the consensus oligonucleotide homeodomain-binding site 5⬘-TCGGGATCCGAGTTAATTGCGC-3⬘ (A20, lanes 22 and 23) incubated with 10 ␮g of rat nuclear proteins isolated from smooth muscle myoblasts (lanes 1–13) and immature smooth muscle myocytes (lanes 14 –23). Lane 1, NE ⫹ labeled CArG box 2; lane 2, NE ⫹ labeled CArG box 2 ⫹ 100-fold excess unlabeled CArG box 2; lane 3, NE ⫹ labeled CArG box 2 ⫹ SRF antibody; lane 4, NE ⫹ labeled CArG box 2 ⫹ 100-fold excess unlabeled CArG box 2 mutant; lane 5, NE ⫹ labeled CArG box 2 ⫹ 100-fold excess unlabeled NS DNA; lane 6, NE ⫹ labeled AT-rich domain; lane 7, NE ⫹ labeled AT-rich domain ⫹ 100-fold excess unlabeled AT-rich domain; lane 8, NE ⫹ labeled AT-rich domain ⫹ MEF2A antibody; lane 9, NE ⫹ labeled AT-rich domain ⫹ MEF2B antibody; lane 10, NE ⫹ labeled AT-rich domain ⫹ MEF2D antibody; lane 11, NE ⫹ labeled AT-rich domain ⫹ 100-fold excess unlabeled ACH; lane 12, NE ⫹ labeled AT-rich domain ⫹ 100-fold excess unlabeled AT-rich domain mutant; lane 13, NE ⫹ labeled AT-rich domain ⫹ 100-fold excess unlabeled NS DNA; lane 14, NE ⫹ labeled HDBS; lane 15, NE ⫹ labeled HDBS ⫹ 100-fold excess unlabeled HDBS; lane 16 NE ⫹ unlabeled HDBS ⫹ TN domain antibody; lane 17, NE ⫹ labeled HDBS ⫹ 100-fold excess unlabeled ACH; lane 18, NE ⫹ labeled HDBS ⫹ 100-fold excess unlabeled HDBS mutant; lane 19, NE ⫹ labeled HDBS ⫹ 100-fold excess unlabeled NS DNA; lane 20, NE ⫹ unlabeled Nkx2–1 oligonucleotide; lane 21, NE ⫹ unlabeled Nkx2-1 oligonucleotide ⫹ TN domain antibody; lane 22, NE ⫹ unlabeled A20 oligonucleotide; lane 23, NE ⫹ unlabeled A20 oligonucleotide ⫹ TN domain antibody. Specific bands identified by Nkx2-1 oligonucleotide and A20 oligonucleotide are marked with asterisks. Note similarity of band using the A20 oligonucleotide to that observed in the HDBS lanes (arrowhead). A CArG box 2 oligonucleotide containing five additional sequence base pairs both 5⬘ and 3⬘ resulted in complete competition for CArG box 2 binding when used in 100-fold excess (data not shown). Utilization of this oligonucleotide was limited, however, because the addition of 5 bp 3⬘ resulted, in essence, in the recreation of the NK-2 homeodomain-binding site and hence corresponds to the CArG box 2/HDBS oligonucleotide.

TATATGGTAATATTGC-3⬘), the AT-rich domain (5⬘-GGATCTTTATTAAAAAAAA-3⬘), CArG box 2 (5⬘-GGATCCCTTATATGG-3⬘), HDBS (5⬘-GGATCGTAATATTG-3⬘), AT-rich/CArG box 2 (5⬘-GGATCTTTATTAAAAAAAAGGATCCCTTATATGG), CArG box 2/HDBS (5⬘-GGATCCCTTATATGGGGATCGTAATATTG-3⬘), nonspecific DNA (5⬘-GG ATCTTCTAAAAGATGCACACTTGCA-3⬘), the consensus c-fos-SRE (5⬘GGATCGGATGTCCATATTAGGACATCTG-3⬘), the ACH substituting the consensus c-fos-SRE for CArG box 2 (5⬘-GGATCTTTATTAAAAAAAACCCACCATATTAGGTAATATTGC-3⬘), and CArG box 2/HDBS substituting the consensus c-fos-SRE for CArG box 2 (5⬘-GGATCGGATGTCCATATTAGGTAATATTG-3⬘). Double-stranded oligonucleotide probes corresponding to the ␥-smooth muscle isoactin ACH or paired elements within the ACH were disrupted by changing the wild-type elements in combination or individually from AT-rich domain 5⬘-TTTATTAAAAAAAA-3⬘ to 5⬘-TTTACTGAACAAGA-3⬘ (AmutCH), CArG box 2 domain 5⬘CCTTATATGG-3⬘ to 5⬘- ACTTACATGT-3⬘ (ACmutH), and HDBS domain 5⬘-GTAATATTG-3⬘ to 5⬘-GTCAGACTG-3⬘ (ACHmut). Oligonucleotide probes corresponding to the consensus homeodomain-binding sequence 5⬘-TCGGGATCGCCCAGTCAAGTGC-3⬘ (Nkx2-1) and the consensus homeodomain-bind-

ing sequence 5⬘-TCGGGATCCGAAGTTAATTGCGC-3⬘ (A20) were also utilized (10). 5 pmol of double-strand DNA probe was end-labeled with [␣-32P]dCTP using 2 units of Klenow (Promega). Unincorporated nucleotides were removed using Sephadex G-25 spin columns (Amersham Pharmacia Biotech). All EMSAs were performed by incubating 50,000 – 100,000 cpm double-strand DNA probe, 10 ␮g of nuclear extract, 1 ␮g of poly(dI-dC), and 100⫻ cold competitor, if necessary, at RT for 30 min. Complexes were then separated on a 7.5% non-denaturing acrylamide gel. For supershift analysis, the same procedure was followed, except 3 ␮l of rabbit polyclonal SRF antibody (Santa Cruz Biotechnology, Inc.), monoclonal MEF2-A, -B, and -D (11, 12), or polyclonal TN domain antisera were added to nuclear extracts prior to the addition of labeled oligonucleotide probe. In initial experiments, each antibody was also added after the addition of labeled probe, and no differences in banding patterns were observed. Recombinant SRF Production—The pT7⌬ATG expression vector described previously (13), containing a full-length SRF open reading frame, was used to produce recombinant SRF protein (rSRF). rSRF was generated using the TnT Quick-coupled Transcription/Translation Sys-


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FIG. 6. Nuclear protein binding is differentially controlled by the individual elements comprising the ACH regulatory triad. Representative EMSA analysis of the 36-bp ACH ␥-smooth muscle isoactin promoter fragment containing mutations in the AT-rich domain (AmutCH), CArg box 2 (ACmutH), and the HDBS domain (ACHmut). Labeled ACH mutants were incubated with 10 ␮g of rat nuclear proteins isolated from smooth muscle myoblasts (MB), immature smooth muscle myocytes (IMC), and neonatal (NMC), adult mature smooth muscle myocytes (AMC), normal uterine myometrium (NM), uterine leiomyomas (UL), and uterine leiomyosarcomas (US). In addition, labeled ACH was incubated with 10 ␮g of rat nuclear proteins isolated from immature smooth muscle myocytes (IMC) as a control (lanes 8 and 16). The following lane pattern is repeated for each ACH mutant. Lanes 1, 9, and 17, smooth muscle myoblasts ⫹ labeled ACH mutant; lanes 2, 10 and 18, immature smooth muscle myocytes ⫹ labeled ACH mutant; lanes 3, 11 and 19, neonatal smooth muscle myocytes ⫹ labeled ACH mutant; lanes 4, 12 and 20, adult mature smooth muscle myocytes ⫹ labeled ACH mutant; lanes 5, 13, and 21, normal uterine myometrium ⫹ labeled ACH mutant; lanes 6, 14 and 22, uterine leiomyomas ⫹ labeled ACH mutant; and lanes 7, 15 and 23, uterine leiomyosarcomas ⫹ labeled ACH mutant. Primary bands are identified alphabetically in series with reference to previous figures. Bands in parentheses represent approximate position of previously identified bands C–E.

tem as described by the supplier (Promega). Translation products were analyzed using SDS-polyacrylamide gel electrophoresis. Verification of SRF production was performed by Western blot analysis using the SRF polyclonal antibody (Santa Cruz Biotechnology) as described below. Northern Blot Analysis—Northern blot analysis was performed as described previously (5). 20 ␮g of total cellular RNA were loaded per lane. Rat Nkx2-3 probe was derived from reverse transcriptase-polymerase chain reaction using the published degenerate primers Nkxupstream 5⬘-GC(ACGT)CA(AG)GT(ACGT)TA(TC)GA(AG)(CT)T(ACGT)GA(AG)(AC)G-3⬘ and Nkx-downstream 5⬘-(AG)TT(CT)TG(AG)AA CCA(AGT)AT(CT)TT(ACGT)AC(CT)TG(ACGT)GT-3⬘ (14). Nkx specificity was confirmed by DNA sequencing. [32P]dCTP-labeled probe was incubated with blot overnight at 55 °C. Blot was stripped and reprobed with glyceraldehyde phosphate dehydrogenase cDNA as a loading control. Western Blot Analysis—Nuclear and cytoplasmic proteins were isolated and quantitated as described above. Nuclear proteins from primary cultures of rat neonatal cardiomyocytes and NIH3T3 fibroblasts (ATCC) were included as non-muscle controls. 25 ␮g of total protein was combined with 2⫻ loading buffer to a final concentration of 50 mM Tris-HCl, pH 6.8, 2% SDS, and 1% 2-mercaptoethanol. Samples were fractionated by SDS-polyacrylamide gel electrophoresis and transblotted onto nitrocellulose membranes using standard techniques. Nitrocellulose membranes were blocked with 5% nonfat dry milk in Trisbuffered saline with 1% Tween 20 (TBS-T) for 1 h at RT and rinsed twice for 5 min each in TBS-T. Membranes were incubated overnight at 4 °C with a 1:1000 dilution of SRF polyclonal antibody (G-20, Santa Cruz Biotechnology). Membranes were washed in TBS-T and incubated for 1 h with a 1:5000 dilution of horseradish peroxidase secondary antibody (Pierce, Inc.). Following incubation with Pierce SuperSignal

chemiluminescence reagent (Pierce), immunoreactivity was visualized by exposure to film (Kodak Biomax MR). Immunohistochemical Analysis—Primary cultures of ISMC were grown on sterile glass coverslips placed in 6-well Costar Brand tissue culture plates (Fisher). Media were removed, and each well was washed with phosphate-buffered saline, pH 7.4, containing 0.1% Tween 20 (PBS-T), 3 times for 5 min each. Cells were fixed for 20 min at RT in PBS-T containing 1% paraformaldehyde. Cells were washed with PBS followed by incubation in PBS-T containing 1% Triton X-100 for 15 min at RT. Cells were washed in PBS-T and blocked for 1 h at RT in PBS-T containing 3% bovine serum albumin followed by a single wash in PBS-T. Cells were incubated at RT overnight with a 1:100 dilution of SRF polyclonal antibody (Santa Cruz Biotechnology). The following day, cells were washed 3 times for 10 min each in PBS-T. Cells were then incubated at RT for 2 h with rhodamine-conjugated secondary antibody (Jackson ImmunoResearch Laboratories). Cells were washed twice for 10 min each with PBS-T. Slides were mounted using Slow Fade Media (Molecular Probes). Isolated neonatal smooth muscle strips were fixed in 3.7% formaldehyde at 4 °C overnight. Muscle strips were serially immersed in 3 and 15% sucrose solutions for 1 h each at 4 °C, followed by immersion in a 30% sucrose solution at 4 °C overnight. Muscle strips were mounted in a cryostat block using tissue-freezing medium, and 0.6-␮m frozen sections serially cut and mounted to Superfrost Plus microscope slides (Fisher). Samples were incubated in PBS-T containing 1% bovine serum albumin for 45 min at RT followed by a single wash with PBS-T. Samples were then incubated for 1 h at RT with a 1:100 dilution of SRF polyclonal antibody (Santa Cruz Biotechnology). Slides were washed with PBS-T and then incubated at RT for 30 min with a 1:200 dilution of rhodamine-conjugated secondary antibody (Jackson ImmunoRe-

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search Laboratories). Slides were washed with PBS-T followed by PBS and mounted. All slides were stored at 4 °C. Immunoreactivity was detected using a Zeiss Axiovert 100 inverted microscope equipped with the Bio-Rad MRC600 Laser Scanning System. For negative controls, slides were incubated in the absence of primary antibody. RESULTS

␥-Smooth Muscle Isoactin Promoter Characterization—Distinct ISMC phenotypes were identified based upon criteria described previously (4). For the purpose of this study, mature smooth muscle myocytes were isolated from both neonatal and adult rats (Fig. 1). A genomic clone containing 1.1 kilobase pairs of the proximal promoter region of the rat ␥-smooth muscle isoactin was isolated and sequenced. Promoter deletion constructs were generated in order to establish the regions of the promoter that were necessary for gene expression (Fig. 2A). Transfection of pLuc744 into ISMCs resulted in a pattern of luciferase expression that was similar to endogenous ␥-smooth muscle isoactin expression over a 10-day culture period (4), whereas transfection of this same construct into NIH3T3 cells resulted in no detectable luciferase activity (data not shown). Transfection of 245 bases of promoter (pLuc-245) was sufficient for the highest levels of luciferase activity. Removal of 134 bases (pLuc-111) resulted in a 10-fold decrease in reporter activity, whereas deletion of an additional 13-bp including CArG box 2 (pLuc-98) completely abolished reporter activity. Two point mutations in CArG box 2 within the context of the full-length promoter (pLuc-744mut) resulted in minimal reporter activity. Taken together, these results demonstrate the requirement of an intact CArG box 2 element for normal ␥-smooth muscle isoactinmediated reporter activity. Distinct ISMC Phenotypes Display Unique Nuclear Protein Binding Patterns—EMSAs were performed to determine if endogenous SRF protein was binding to the ␥-smooth muscle isoactin promoter. A 36-bp probe containing CArG box 2 flanked by a 14-bp AT-rich region and a canonical homeodomain-binding site (HDBS) for Antennapedia class homeobox proteins was used for these studies (Fig. 2B). Inclusion of these flanking elements was based upon their potential involvement in the binding of transcriptional co-activators. Smooth muscle myoblasts showed five primary bands upon EMSA analysis (Fig. 3). The banding pattern for ISMC was distinct from that observed for cultured cardiomyocytes and NIH3T3 fibroblasts (data not shown). The intensity of bands A and B increased in immature smooth muscle myocytes and then decreased and disappeared in neonatal and adult smooth muscle myocytes, respectively. Whereas band C initially decreased in intensity in immature smooth muscle myocytes, it became the predominant band in neonatal and adult smooth muscle myocytes. Bands D and E were only observed in smooth muscle myoblasts. DNA binding specificity was demonstrated by competition with unlabeled ACH versus nonspecific DNA. Bands A and B were selectively competed by inclusion of CArG/HDBS. Band C was selectively competed by inclusion of cold AT/CArG in smooth muscle myoblasts and immature smooth muscle myocytes. However, in neonatal and adult smooth muscle myocytes, selective competition of band C occurred with CArG/ HDBS, whereas AT/CArG only minimally competed for the band. This suggests that in neonatal and adult smooth muscle myocytes the composition of band C is distinct from that observed in smooth muscle myoblasts and immature smooth muscle myocytes. Consequently, this band was designated C⬘. Bands D and E were selectively competed by the inclusion of cold AT/CArG. These observations indicate that distinct ISMC phenotypes display unique binding patterns to the 36-bp ACH ␥-smooth muscle isoactin promoter fragment.

FIG. 7. Binding to the ACH domain is CArG box 2 sequencespecific. Representative EMSA analysis comparing replacement of CArG box 2 with the consensus c-fos SRE. 10 ␮g of rat nuclear proteins isolated from immature smooth muscle myocytes were incubated with labeled ACH, c-fos-substituted ACH (AFH), CArG-HDBS, c-fos-substituted CArG-HDBS (fCArG-HDBS), CArG box 2 (CArG), and c-fos SRE (fCArG). Lane 1, NE ⫹ labeled ACH; lane 2, NE ⫹ labeled ACH ⫹ 100-fold excess ACH; lane 3, NE ⫹ labeled AFH; lane 4, NE ⫹ labeled AFH ⫹ 100-fold excess AFH; lane 5, NE ⫹ labeled CArG-HDBS; lane 6, NE ⫹ labeled CArG-HDBS ⫹ 100-fold excess CArG-HDBS; lane 7, NE ⫹ labeled fCArG-HDBS; lane 8, NE ⫹ labeled fCArG-HDBS ⫹ 100-fold excess fCArG-HDBS; lane 9, NE ⫹ labeled CArG box 2; lane 10, NE ⫹ labeled CArG box 2 ⫹ 100-fold excess CArG box 2; lane 11, NE ⫹ labeled c-fos SRE; lane 12, NE ⫹ labeled c-fos SRE ⫹ 100-fold excess c-fos SRE. Primary bands are identified alphabetically in series with reference to previous figures. Bands in parentheses represent approximate position of previously identified bands C-E. Each c-fos SRE lane shows the presence of additional bands marked with arrowheads.

SRF Binds to the ␥-Smooth Muscle Isoactin Promoter—The addition of SRF antibody caused bands A and B to supershift, confirming that SRF was indeed binding to this portion of the promoter (Fig. 4). The presence, intensity, and eventual disappearance of bands A and B mirrored the pattern of SRF supershift. SRF binding increased in immature smooth muscle myocytes relative to smooth muscle myoblasts and then decreased and eventually disappeared in neonatal and adult smooth muscle myocytes, respectively. Utilization of CArG box 2 as a probe produced bands A and B and both were supershifted upon the addition of SRF antibody (Fig. 5). Mutation of CArG box 2 within the context of the ACH (ACmutH) resulted in the disappearance of bands A, B, and C⬘, whereas bands C, D, and E remained unaffected (Fig. 6). Substitution of the consensus c-fos SRE for CArG box 2 altered the binding pattern from that observed with native ACH, CArG-HDBS, and CArG box 2 (Fig. 7). Although bands


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FIG. 8. SRF binding to the ACH domain involves additional cofactors. A, verification of recombinant SRF (rSRF) production synthesized from the plasmid pT7⌬ATG (13). Western blot analysis using SRF antibody of 150 ␮g of rSRF (lane 1), 15 ␮g of rSRF (lane 2), and 1.5 ␮g of rSRF (lane 3); 5 ␮l of 35S-labeled rSRF (lane 4). Lower molecular weight protein band marked by arrowhead most likely represents early termination product following in vitro translation. Extrapolated molecular weight of upper SRF band is 62 kDa. B, representative EMSA analysis of the 36-bp ACH ␥-smooth muscle isoactin promoter fragment comparing the binding pattern of 10 ␮g of nuclear protein isolated from immature smooth muscle myocytes with rSRF protein. Lane 1, NE ⫹ labeled ACH; lane 2, NE ⫹ labeled ACH ⫹ SRF antibody; lane 3, rSRF protein ⫹ labeled ACH; lane 4, rSRF protein ⫹ labeled ACH ⫹ SRF antibody; lane 5, rSRF protein ⫹ labeled ACH ⫹ TN domain antibody. Note faster migration of rSRF bands when compared with nuclear proteins and the inability of TN domain antibody to supershift rSRF band.

FIG. 9. Nkx2-3 expression increases as ISMCs differentiate in vitro. Northern blot analysis of Nkx2-3 expression identified a single band that was differentially expressed in neonatal mature smooth muscle myocytes (NMC), and primary cultures of rat ISMCs on days 1 (D1, smooth muscle myoblasts, MB), 3 (D3), 5 (D5), 7 (D7), and 10 (D10, immature smooth muscle myocytes, IMC).

A–C were present, the appearance of additional unique lower bands dominated the EMSA pattern. Inclusion of SRF antibody resulted in the supershift of bands A and B in a manner similar to that observed for native CArG box 2 (data not shown). These experiments show that unique patterns of SRF binding to the ␥-smooth muscle isoactin promoter are observed during smooth muscle differentiation. This binding is sequence-specific and appears to involve additional co-factors in association with SRF. An NK-2 Homeobox Transcription Factor Is Expressed in Cultured ISMCs and Binds to the ␥-Smooth Muscle Isoactin Promoter—Because a putative TAAT cis-binding element was found immediately adjacent to CArG box 2, we investigated homeodomain proteins that may act as mediators of smooth muscle-specific gene expression. One obvious candidate was the NK family of transcription factors (15–17). EMSA analysis using an antibody that recognizes a conserved amino-terminal domain (TN domain) found in a subset of NK-2 proteins resulted in a single supershifted band S2 (Fig. 4). Inclusion of the TN domain antibody completely supershifted all bands present in smooth muscle myoblasts and immature smooth muscle myocytes. In contrast, the TN domain antibody only partially supershifted band C⬘ in neonatal and adult smooth muscle

myocytes. NK-2 protein binding appeared to increase as ISMCs progressively differentiated. Interestingly, both the SRF and TN domain antibodies supershifted bands A and B, and the sequential administration of both antibodies resulted in a super-supershift of each individual band (data not shown). Competition analysis showed that the ACH and CArG/HDBS partially blocked the TN supershifted band, whereas the AT/CArG and HDBS had no effect (data not shown). Utilization of the HDBS domain as a probe resulted in the production of a single band (Fig. 5). Although unaffected by the inclusion of NS DNA, the specificity of this band could not be confirmed by competition with excess HDBS or ACH. However, utilization of oligonucleotide probes containing either a consensus 5⬘-CAAGTG-3⬘ (Nkx2-1) or 5⬘-TTAATT-3⬘ (A20)binding site resulted in the production of unique bands (Fig. 5), one of which appeared to overlap with the HDBS-specific band. Prior affinity studies have suggested that the consensus 5⬘CAAGTG-3⬘ and 5⬘-TTAATT-3⬘-binding sites are much higher affinity sites than the HDBS (5⬘-GTAATATTG-3⬘) site, a fact supported by our EMSA data. Mutation of the HDBS element within the context of the ACH (ACHmut) resulted in a diminution of binding intensity for all bands present, rather than ablation of individual bands (Fig. 6). To confirm that bands A and B represented a protein complex containing both SRF and an NK-2 factor, we performed EMSA analysis using recombinant SRF protein (Fig. 8, rSRF). rSRF produced a single unique band using both the ACH (A⬘) and CArG box 2 (data not shown) as probes. This band migrated slightly faster than band A and supershifted in the presence of the SRF antibody but not the TN domain antibody. The rSRF supershifted band (S1⬘) also migrated slightly faster than that observed for nuclear proteins. Taken together, these data indicate that bands A and B are composed of a protein complex minimally containing SRF and an NK-2 factor. In addition, binding of the NK-2 factor appears to occur principally through protein-protein interactions with SRF, rather

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FIG. 10. Human smooth muscle neoplasms display unique nuclear protein binding patterns to the ACH domain. Representative EMSA analysis using the 36-bp ACH promoter fragment following incubation with human nuclear proteins isolated from normal myometrium (lanes 1– 8, experimental n ⫽ 3), uterine leiomyomas (lanes 9 –16, experimental n ⫽ 12), uterine leiomyosarcomas (lanes 17–24, experimental n ⫽ 5), and cultured rat (R) and human (H) smooth muscle myoblasts (MB). Lanes 1, 9, and 17, NE ⫹ labeled ACH; lanes 2, 10, and 18, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled ACH; lanes 3, 11, and 19, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled AT-rich/CArG box 2; lanes 4, 12, and 20, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled CArG box 2/HDBS; lanes 5, 13, and 21, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled CArG box 2; lanes 6, 14, and 22, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled AT-rich; lanes 7, 15, and 23, NE ⫹ labeled ACH ⫹ 100-fold excess unlabeled HDBS; and lanes 8, 16, and 24, NE ⫹ labeled ACH 100-fold excess unlabeled NS DNA; lane 25, rat intestinal smooth muscle myoblast NE ⫹ labeled ACH; lane 26, human intestinal smooth muscle myoblast NE ⫹ labeled ACH. Primary bands are identified alphabetically in series with reference to previous figures. Primary bands were competed by inclusion of 100-fold excess ACH and CArG-HDBS, whereas 100-fold excess NS DNA is ineffective. Bands C⬙ and C⵮ have overlapping positions with original bands C/C⬘. However, this band is uniquely competed by inclusion of 100-fold excess AT-CArG in normal myometrium and hence is designated C⬙ and is uniquely composed of a clear doublet in both uterine leiomyomas and leiomyosarcomas and hence designated C⵮. Band F in normal myometrium is also competed by inclusion of 100-fold excess AT-CArG. Note similar banding patterns between rat and human intestinal smooth muscle myoblasts in lanes 25 and 26 suggesting that the unique uterine binding patterns observed for uterine smooth muscle cells are tissue rather than species-specific.

than direct protein binding to the HDBS element. In an effort to identify which NK-2 protein was binding to the ␥-smooth muscle isoactin promoter, we performed a degenerate PCR screen for NK homologues expressed in ISMCs (data not shown). Sequence analysis revealed the presence of only Nkx2-3 clones (data not shown). Northern blot analysis verified Nkx2-3 expression in rat ISMCs (Fig. 9). The moderate level of Nkx2-3 expression observed in neonatal smooth muscle myocytes (NMC) was maintained in smooth muscle myoblasts even after 24 h in culture (MB/D1). Nkx2-3 expression then steadily increased as ISMCs differentiated into immature smooth muscle myocytes (IMC/D10). This observation is consistent with our EMSA data showing NK-2 protein binding increased steadily as ISMCs differentiated. Although indirect evidence, these data suggest that the NK-2 factor found associated with SRF in ISMC is Nkx2-3. Myocyte Enhancer Factors Do Not Bind to the 36-bp Transcriptional Complex in Rat ISMCs—Although no canonical binding site for MEF2 is present in the ␥-smooth muscle isoactin promoter, the AT-rich region found within our 36-bp element possesses some similarities to a degenerate MEF2binding site (18). In addition, MEF2 proteins, like SRF, can participate in transcriptional activation and repression independent of DNA binding via protein-protein interactions mediated through a MADS domain (18 –21). By using isoform-specific antibodies, we assayed for the presence of MEF2A, -B, and -D in each distinct smooth muscle cell

phenotype. No supershifted bands were observed for MEF2 isoforms in any of the ISMC phenotypes (Fig. 4). When the AT-rich domain was used as a probe, a single unique band was observed upon EMSA analysis (Fig. 5). Interestingly, this band was only present in smooth muscle myoblasts and did not supershift with any of the MEF2 antibodies. Mutation of the AT-rich domain within the context of the ACH (AmutCH), resulted in the disappearance of bands C–E and a moderate to significant decrease in intensity for band C⬘ in neonatal and adult smooth muscle myocytes, respectively (Fig. 6). These observations suggest that while MEF2A, -B, and -D do not appear to be involved in the formation of the core transcriptional complex found associated with the 36-bp ACH fragment in smooth muscle myoblasts, an unidentified protein is binding to this domain, perhaps in association with SRF and the NK-2 factor. Analysis of Transcription Factor Binding in Normal and Diseased Human Uterine Smooth Muscle Tissue—The determination of the malignant potential of smooth muscle neoplasms is strongly correlated with an absence of ␥-smooth muscle isoactin expression. (7–9). With the knowledge that SRF and an NK-2 factor cooperatively bind to the ␥-smooth muscle isoactin promoter in a differentiation-specific manner, we decided to examine whether there were alterations in the binding of these proteins in nuclear extracts from normal uterine myometrium, uterine leiomyomas, and uterine leiomyosarcomas. The 36-bp promoter fragment used in this study is 92% conserved be-


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FIG. 11. SRF and MEF2A, -B, and -D binding are altered in neoplastic versus normal uterine smooth muscle cells. Representative EMSA supershift analysis using the 36-bp ACH promoter fragment following incubation with human nuclear proteins isolated from normal myometrium (lanes 1– 6), uterine leiomyomas (lanes 7–12), and uterine leiomyosarcomas (lanes 13–18). Lanes 1, 7, and 13, NE ⫹ labeled ACH; lanes 2, 8, and 14, NE ⫹ labeled ACH ⫹ SRF antibody; lanes 3, 9, and 15, NE ⫹ labeled ACH ⫹ MEF2A antibody; lanes 4, 10, and 16, NE ⫹ labeled ACH ⫹ MEF2B antibody; lanes 5, 11, and 17, NE ⫹ labeled ACH ⫹ MEF2D antibody; and lanes 6, 12, and 18, NE ⫹ labeled ACH ⫹ TN domain antibody. Primary bands are identified alphabetically in series with reference to previous figures. The position of the SRF supershifted band is marked S1; the TN domain supershifted band is S2, and the MEF2 supershifted band is S3. Note that inclusion of the MEF2 antibodies produced faster migrating bands in normal myometrium (band G) and uterine leiomyomas (band H). In addition, inclusion of the MEF2 antibodies resulted in intensification of band C⬙ in normal myometrium.

tween rat and human, suggesting that transcription factors of human origin should also be able to recognize and bind to this probe (Fig. 2B). Supporting this contention, nuclear proteins from cultured rat and human ISMCs display similar EMSA patterns (Fig. 10). Two primary bands were observed upon EMSA analysis of normal myometrium (Fig. 10), with band C⬙ showing an overlapping migration with the previously identified bands C/C⬘ in ISMCs. Leiomyomas and leiomyosarcomas possessed a unique, well defined doublet, C⵮, that ran in a position overlapping band C⬙. All leiomyomas examined also possessed varying intensities of bands A and B as originally identified in cultured ISMCs, whereas these same bands were faint and only found in about 60% of leiomyosarcomas. DNA binding specificity was demonstrated by competition with unlabeled ACH versus nonspecific DNA. Competition analysis revealed that the ACH, AT/CArG, and CArG/HDBS all partially blocked binding of the uterine nuclear proteins to the 36-bp promoter fragment. SRF antibody only produced a supershift when bands A and B were present, and therefore was only observed in uterine leiomyomas and 60% of uterine leiomyosarcomas (see Fig. 11, S1). The addition of TN domain antibody consistently produced a supershifted band in normal myometrium, uterine leiomyomas, and uterine leiomyosarcomas, with NK-2 factor binding intensity appearing almost identical in all three tissues. In normal myometrium, inclusion of the MEF2A, -B, and -D antibodies produced several effects. First, two unique bands appeared. One supershifted band S3 migrated to a position just beneath band B, and the second band G migrated slightly faster than band F. Second, the addition of the MEF2 antibodies appeared to enhance the intensity of band C⬙. In uterine leiomyomas, the addition of MEF2 antibodies also produced a faster migrating band H whose position was unique from that

observed in normal myometrium. Both bands G and H appeared distinct from bands D and E in ISMCs. The MEF2 antibodies had no apparent effect upon nuclear protein binding in uterine leiomyosarcomas. Whether the higher mobility MEF2 bands represent a unique supershift of previously uncomplexed proteins or the removal of MEF2 from the assay by antibody competition permitting an additional factor(s) to bind to the ACH remains to be determined. These experiments indicate that distinct alterations in MEF2 and SRF transcription factor binding were observed in normal versus neoplastic smooth muscle tissues. In addition, the observed similarities in EMSA banding patterns between rat and human ISMCs suggest that the unique banding patterns observed for uterine tissues probably represent tissue-specific utilization of the 36-bp promoter domain. Cytoplasmic Versus Nuclear Translocation of SRF—Recent evidence indicates that the physiologic control of smooth muscle-specific gene expression may be regulated through the nuclear translocation of SRF (22). To investigate whether a similar mechanism may play a role in ISMC differentiation, we compared SRF protein expression in nuclear versus cytoplasmic fractions of each smooth muscle cell phenotype and uterine smooth muscle tumor (Fig. 12A). The pattern of SRF protein expression in the nuclear fraction was similar to that observed by EMSA analysis. SRF protein was expressed at high levels in immature smooth muscle myocytes, low levels in smooth muscle myoblasts, neonatal mature smooth muscle myocytes, uterine leiomyomas, and undetectable levels in adult mature smooth muscle myocytes, normal myometrium, and leiomyosarcomas. In the cytoplasmic fraction, SRF protein was expressed at moderate levels in immature and neonatal smooth muscle myocytes, low levels in smooth muscle myoblasts and leiomyosarcomas, and undetectable levels in adult smooth

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FIG. 12. Nuclear translocation of SRF is modulated during ISMC differentiation and smooth muscle tumorigenesis. A, comparative Western blot analysis of SRF expression in the nuclear versus cytoplasmic pool of smooth muscle myoblasts (MB), immature smooth muscle myocytes (IMC), neonatal mature smooth muscle myocytes (NMC), adult mature smooth muscle myocytes (AMC), normal myometrium (NM), uterine leiomyoma (UL), and uterine leiomyosarcoma (US). Extrapolated molecular mass of band is 62 kDa. Note discordance in expression levels of cytoplasmic SRF versus nuclear SRF protein in NMC and UL. B, representative EMSA analysis of the 36-bp ACH promoter fragment incubated with 10 ␮g of cytoplasmic proteins isolated from smooth muscle myoblasts (lanes 1–3), immature smooth muscle myocytes (lanes 4 – 6), neonatal mature smooth muscle myocytes (lanes 7–9), adult mature smooth muscle myocytes (lanes 10 and 11), normal myometrium (lanes 12–14), uterine leiomyomas (lanes 15–17), and uterine leiomyosarcomas (lanes 18 –20). Lanes 1, 4, 7, 10, 12, 15, and 18, NE ⫹ labeled ACH; lanes 2, 5, 8, 11, 13, 16, and 19, NE ⫹ labeled ACH ⫹ SRF antibody; and lanes 3, 6, 9, 14, 17, and 20, NE ⫹ labeled ACH ⫹ TN domain antibody. Primary bands are identified alphabetically in series with reference to previous figures. The position of the SRF supershifted band is marked S1. The positions of the TN domain supershifted bands are marked S2 and S4. Compare intensity of A/B and S1 bands in neonatal mature smooth muscle myocytes to that previously observed for nuclear proteins in Fig. 4, lanes 13 and 14.

muscle myocytes, normal myometrium, and leiomyomas. EMSA analysis of cytoplasmic proteins revealed a similar pattern of SRF binding to the ACH (Fig. 12B). Four primary bands were observed, including the original bands A and B seen in nuclear extracts. Addition of SRF antibody produced a single supershifted band S1. Although inclusion of TN domain antibody produced a single supershifted band S2 in smooth muscle myoblasts and immature smooth muscle myocytes, a second faster migrating band S4 was also observed in neonatal mature smooth muscle myocytes and uterine leiomyosarcomas. The pattern of cytoplasmic SRF protein expression and ACH binding was distinct from that observed for nuclear protein. The difference was most apparent in neonatal mature smooth muscle myocytes where modest/high levels of SRF protein expression and ACH binding were observed in the cytoplasmic fraction compared with low/undetectable levels of SRF protein expression and ACH binding in the nuclear fraction. This observation suggests that during certain stages of smooth muscle development nuclear access of SRF may be limited. This hypothesis was supported by immunohistochemical analysis of SRF expression in smooth muscle myoblasts, immature smooth muscle myocytes, and neonatal mature smooth muscle myocytes (Fig. 13). SRF expression in smooth muscle myoblasts was predominantly nuclear in localization. Imma-

ture smooth muscle myocytes showed mixed expression with SRF being present in both the nuclear and cytoplasmic compartments. Finally, SRF expression in neonatal mature smooth muscle myocytes appeared predominantly cytoplasmic with only a small number of nuclei stained. These studies support a role for nuclear translocation of SRF in the regulation of smooth muscle-specific gene expression. DISCUSSION

The necessity of CArG box 2 and its ability to bind SRF during smooth muscle differentiation have been clearly demonstrated. SRF is a member of the MADS box transcription factor family and binds to the consensus CArG box element as a homo- or heterodimeric complex (23, 24). SRF is capable of displaying distinct biological functions based upon its specificity for co-regulatory protein partners and DNA-binding sites (15–17, 25, 26). Although SRF is ubiquitously expressed during development, it has been shown to preferentially accumulate in all three muscle lineages (15). SRF regulates muscle-specific gene expression (15, 16, 27–30), and binding of SRF to the chicken homologue of the ␥-smooth muscle isoactin promoter has been reported previously (31). Our results demonstrate that SRF activity also plays an important role in regulating visceral smooth muscle-specific


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FIG. 13. SRF protein shows differential expression in nuclear versus cytoplasmic compartments during smooth muscle development. Immunohistochemical localization of SRF protein in smooth muscle myoblasts (A), immature smooth muscle myocytes (B), and neonatal smooth muscle strips composed of mature smooth muscle myocytes (C). A and B, all of the cells/nuclei (n) were labeled. C, all of the cells but only some of the nuclei (n) were labeled.

gene expression in mammals. In visceral smooth muscle cells, SRF binding to CArG box 2 appears not only dependent upon expression of SRF protein but also its accessibility to the nuclear compartment. A similar block in the nuclear localization of SRF has been observed during adipocyte differentiation (32). In addition, recent evidence suggests that physiologic regulation of smooth muscle myosin heavy chain and SM22 transcription is mediated through nuclear to cytoplasmic redistribution of SRF (22). Our studies provide the first evidence that similar mechanisms may exist in the regulation of SRF expression during visceral smooth muscle development. Future analysis of smooth muscle-specific gene expression must therefore include an assessment of SRF expression in conjunction with its cellular distribution. SRF binding appears most prevalent when ISMCs are in a less differentiated, highly proliferative state. This may suggest that SRF expression is involved in establishing the modulatory potential frequently observed during smooth muscle cell dedifferentiation. As differentiation progresses, SRF binding decreases. The initial decrease in SRF binding occurs in neonatal smooth muscle myocytes via cytoplasmic translocation, which is then followed by a precipitous decrease in SRF protein expression in adult smooth muscle myocytes. The cytoplasmic accessibility of SRF protein in neonatal smooth muscle myocytes may explain, in part, why these cells show increased modulatory potential when compared with adult smooth muscle myocytes. Although SRF binding appears to play a critical role in regulating smooth musclespecific gene expression, its activity alone is insufficient for the proper developmental and tissue-specific expression of ␥-smooth muscle isoactin (Fig. 14). The ability of SRF to interact with accessory factors, and thereby differentially regulate tissue-specific gene expression, has been clearly demonstrated (15–17, 25, 26). Our data also support a role for an NK-2 homeobox factor participating in the regulation of a smooth muscle-specific promoter. The NK-2 class of homeobox genes constitutes a subfamily of sequencespecific transcription factors, several members of which play an essential role in mesoderm specification, cardiac development, and visceral smooth muscle lineage determination (14, 33–36). NK-2 homeodomain factors regulate muscle development by several potentially overlapping mechanisms, including cooperative association with MADS box factors (15–17). The mamma-

lian homologues of the Drosophila genes bagpipe (Nk-3) and tinman (Nk-2) represent two attractive NK-2 candidates that may be participating in smooth muscle development (14, 33– 36). Although both of these molecules are expressed in undifferentiated visceral mesoderm during murine embryogenesis (14, 36), Nkx2-3 appears to be the predominant NK-2 transcription factor expressed in rat ISMCs. Our data suggest that SRF is cooperatively associating with Nkx2-3 through protein-protein interactions, with direct DNA binding of this regulatory complex occurring principally through CArG box 2. The ␥-smooth muscle isoactin gene therefore represents the first direct target for endogenous NK-2 transcription factor binding. Previous studies suggest that the cooperative association of Nkx2-3 with SRF would result in an increased affinity of the regulatory complex for CArG box 2 (16, 37). Nkx2-3 binding remains relatively constant throughout smooth muscle development and lacks the nuclear to cytoplasmic translocation properties associated with SRF (Fig. 14). However, since SRF binding decreases during smooth muscle development, Nkx2-3 binding eventually predominates in the more mature smooth muscle cell phenotypes. Although recent in vitro studies indicate that SRF and Nkx3-1 interact to cooperatively activate avian ␥-smooth muscle isoactin promoter constructs, expression analysis did not detect this NK homologue in rat ISMCs. In addition, the developmental and tissuespecific expression pattern of Nkx3–1 is somewhat inconsistent with its participation in regulating visceral smooth muscle development in mammals (38 – 40). The presence of a potential degenerate MEF2-binding site in close approximation to CArG box 2 suggested that MEF2 may also participate in the transcriptional regulation of ␥-smooth muscle isoactin expression. Four genes encoding MEF2 (MEF2A–D) have been identified (41), and members of the MEF2 family have been shown to regulate the expression of genes involved in promoting and maintaining muscle differentiation (19, 20, 42– 48). MEF2A, -B, and -D are expressed in developing and mature smooth muscle tissues, whereas expression patterns and functional studies link MEF2C with cardiac and skeletal muscle development (41, 49 –52). Loss of function studies indicate that MEF2 is critical for the progression of myoblast differentiation in all three muscle lineages (43– 47). MEF2A, -B, and -D binding to the 36-bp ␥-smooth muscle isoactin promoter fragment was not observed in ISMCs. How-

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FIG. 14. Summation of transcription factor binding versus ␥-smooth muscle isoactin expression, SMC differentiation, and SMC proliferation. Nuclear versus cytoplasmic binding of SRF and an NK-2 factor to the 36-bp ACH promoter element is correlated with smooth muscle cell (SMC) phenotype, ␥-smooth muscle isoactin (␥-SMA) gene expression, cellular differentiation, cellular proliferation, MEF2 binding, and binding to the AT-rich element. Statistical analysis was performed by randomized, one-way analysis of variance followed by the post hoc Newman-Keuls multiple comparisons test. For all developmental time points there were significant differences in SRF (F(7,26) ⫽ 42.7; p ⬍ 0.0001) and Nkx (F(7,24) ⫽ 3.1; p ⫽ 0.0277) binding to the ACH. Significant differences in SRF (F(5,20) ⫽ 19.0; p ⬍ 0.0001) and Nkx (F(5,17) ⫽ 7.1; p ⫽ 0.0027) binding to the ACH were also observed for uterine smooth muscle tumors versus normal myometrium. Probability values († ⫽ p ⬍ 0.01; †† p ⬍ 0.05) were sequentially calculated for each ISMC phenotype and individually calculated as normal versus neoplasm for uterine smooth muscle tissues. Probability values along the x axis represent a statistical comparison of nuclear versus cytoplasmic SRF and Nkx binding to the ACH.

ever, binding to the AT-rich element alone was detected in smooth muscle myoblasts but was not observed in any other ISMC phenotype. The specificity of this band to smooth muscle myoblasts, where ␥-smooth muscle isoactin is minimally expressed, makes it a tempting inhibitory target for this regulatory complex. Studies are currently underway to identify and further characterize the factor that binds to this element. In addition to a role in smooth muscle development, ␥-smooth muscle isoactin represents a reliable marker for the malignant potential of smooth muscle tumors, with ␥-smooth muscle isoactin expression being significantly reduced in leiomyomas and completely absent in leiomyosarcomas (7–9). Over 95% of all smooth muscle tumors arise in the female genital tract, with the few remaining lesions variably arising in the gastrointestinal tract, vasculature, skin, and bladder (1, 53–54). Pathological evidence indicates that leiomyosarcomas do not arise from pre-existing leiomyomas (9). Examination of a variety of muscle-specific markers suggests that smooth muscle tumors are composed of less differentiated smooth muscle cell phenotypes (7, 8, 55–59). Clonality studies of leiomyomas suggest that chromosomal abnormalities occur secondary to tumor development (60). Therefore, it is likely that the etiology of smooth muscle oncogenesis involves a dysregulation in the differentiation of smooth muscle cells as a result of alterations in both genetic and epigenetic factors that converge on a common biologic pathway. Based upon these observations, we extended our studies to include an examination of the activity of the 36-bp ␥-smooth muscle isoactin promoter fragment in normal and neoplastic uterine smooth muscle tissues. The unique EMSA patterns and tissue-specific utilization of MEF2 isoforms by normal myometrium provide the first transcriptional evidence that uterine smooth muscle cells may dif-

ferentiate by a mechanism distinct from ISMCs. This molecular evidence is consistent with prior morphologic data that suggested urogenital smooth muscle development was distinct from that observed for gastrointestinal smooth muscle (6). Utilization of MEF2 isoforms, as well as an NK factor, by normal myometrium is consistent with the development of a fully differentiated tissue, as both transcription factors are presumed to promote smooth muscle cell differentiation. Although we are unsure of the significance of multiple MEF2 isoforms binding to the ␥-smooth muscle isoactin promoter, it may represent a novel feature of uterine smooth muscle cells, which are known to possess unique sensitivities to cycling steroid hormones (61– 66). At the very least, this observation would support a possible tissue-specific role for MEF2 regulation in smooth muscle differentiation. The reappearance of SRF binding in benign and malignant uterine smooth muscle neoplasms occurs as MEF2 binding concomitantly decreases and disappears. The discordant utilization of these two transcription factors is highly suggestive of a loss in differentiation potential, thereby creating a smooth muscle cell phenotype that is more proliferative and most similar to that observed during embryonic and early neonatal development. The observed transcriptional profile is therefore consistent with prior characterization studies that have suggested that smooth muscle tumors are composed of less differentiated smooth muscle cell phenotypes (8, 9, 53, 54). In uterine smooth muscle neoplasms, this transcriptional shift is marked by a decrease and eventual disappearance of ␥-smooth muscle isoactin expression in leiomyomas and leiomyosarcomas, respectively. Interestingly, SRF expression in leiomyosarcomas shows a modest shift to the cytoplasmic pool (4 cytoplasmic to

Regulation of Smooth Muscle Differentiation 1 nuclear) suggesting that a vestige of this regulatory paradigm may exist in these tumors. The molecular basis for the neoplastic transformation of smooth muscle cells is unknown. Gross chromosomal anomalies have been reported in 40 –50% of smooth muscle tumors, with the most common genetic alteration being a deletion of the long arm of chromosome 7 (67–73). Deletion mutations in two of the six collagen type IV subunits have been shown to result in Alport syndrome as well as diffuse leiomyomatosis (74). Collagen type IV, a critical component of basement membrane, has been postulated as important in the normal development of smooth muscle tissues (75, 76). Therefore, defects in its structure may result in a higher incidence of developmental miscues resulting in smooth muscle tumor formation. A rodent model that possesses a germ line mutation in the tumor susceptibility gene, tuberous sclerosis 2 (TSC2), has been shown to develop uterine leiomyomas in conjunction with renal cell carcinomas and angiosarcomas (77). Unfortunately, the functional role that TSC2 plays in the development of these neoplasms is not known. A novel mutation in a member of the ras gene family, TC21, has also been reported in a single leiomyosarcoma cell line (78). Our data suggest that two members of the MADS box transcription factor family, SRF and MEF2, display aberrant binding patterns in normal versus neoplastic smooth muscle tissues. Neoplastic transformation has been shown to block the differentiation-induced inhibition of SRF interactions with serum-response elements (32). However, whether MADS box transcription factors represent downstream markers or primary determinants of the neoplastic transformation of smooth muscle cells remains to be determined. Of the transcription factors examined, the NK-2 protein is the only factor that consistently binds to the core promoter probe in all smooth muscle cell phenotypes (Fig. 14). This finding is consistent with the requirement of an NK2 protein for specification and maintenance of the visceral smooth muscle cell phenotype. While we hypothesize that Nkx2-3 is the factor binding to our promoter domain, this gene has recently been knocked out in mice, and there is no overt disruption of ISMC specification or differentiation (79). This absence of phenotype may be explained by the compensation for a lack of Nkx2-3 by another NK-2 family member. Unfortunately, even the cooperative binding of Nkx2-3 and SRF to the ␥-smooth muscle isoactin promoter seems insufficient for the proper regulation of this gene during development (Fig. 14). In fact, a significant development conundrum exists in that a variety of studies, including our own, have demonstrated that smooth muscle-specific gene expression is positively regulated CArG box elements. However, SRF binding appears least prevalent in mature smooth muscle cell phenotypes, exactly where one would expect to see the highest levels of smooth muscle-specific gene expression. Even if the cooperative association of Nkx2-3 with SRF increases binding affinity, there is a lack of differential binding of Nkx2-3 in distinct ISMC phenotypes. These observations clearly suggest that additional regulatory factors and/or elements are involved in the developmental and tissue-specific regulation of smooth muscle genes. In summary, smooth muscle differentiation appears to be controlled by a complex interplay of multiple transcription factors including SRF, MEF2, and an NK-2 protein. The precise balance and utilization of these factors may provide a basis for the highly plastic nature associated with smooth muscle development and pathogenesis. A continuing goal for future study will be to clarify the complex molecular pathways controlling normal and abnormal smooth muscle differentiation.

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