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Originally published In Press as doi:10.1074/jbc.M501044200 on May 9, 2005

J. Biol. Chem., Vol. 280, Issue 27, 25854-25863, July 8, 2005
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Regulation of Smooth Muscle-specific Gene Expression by Homeodomain Proteins, Hoxa10 and Hoxb8*

Omar El-Mounayri, Jason W. Triplett, Charles W. Yates, and B. Paul Herring{ddagger}

From the Department of Cellular and Integrative Physiology, Indiana University School of Medicine, Indianapolis, Indiana 46202

Received for publication, January 27, 2005 , and in revised form, April 28, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Smooth muscle cells arise from different populations of precursor cells during embryonic development. The mechanisms that specify the smooth muscle cell phenotype in each of these populations of cells are largely unknown. In many tissues and organs, homeodomain transcription factors play a key role in directing cell specification. However, little is known about how these proteins regulate smooth muscle differentiation. Using degenerate reverse transcription-PCR coupled to cDNA library screening we identified two homeodomain proteins, Hoxa10 and Hoxb8, which are expressed in adult mouse smooth muscle tissues. All three of the previously described transcripts of the Hoxa10 gene, Hoxa10-1, Hoxa10-2, and Hoxa10-3, were identified. Hoxa10-1 directly activated the smooth muscle-specific telokin promoter but did not activate the SM22{alpha}, smooth muscle {alpha}-actin, or smooth muscle myosin heavy chain promoters. Small interfering RNA-mediated knock-down of Hoxa10-1 demonstrated that Hoxa10-1 is required for high levels of telokin expression in smooth muscle cells from uterus and colon. On the other hand, Hoxb8 inhibited the activity of the telokin, SM22{alpha}, and smooth muscle {alpha}-actin promoters. Cotransfection of Hoxa10-1 together with Hoxa10-2 or Hoxb8 suggested that Hoxa10-2 and Hoxb8 act as competitive inhibitors of Hoxa10-1. Results from gel mobility shift assays demonstrated that Hoxa10-1, Hoxa10-2, and Hoxb8 bind directly to multiple sites in the telokin promoter. Mutational analysis of telokin promoter reporter genes demonstrated that the three homeodomain protein binding sites located between -80 and -75, +2 and +6, and +14 and +17 were required for maximal promoter activation by Hoxa10-1 and maximal inhibition by Hoxb8. Together these data demonstrate that the genes encoding smooth muscle-restricted proteins are direct transcriptional targets of clustered homeodomain proteins and that different homeodomain proteins have distinct effects on the promoters of these genes.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Smooth muscle cells (SMC)1 are derived from diverse populations of precursor cells during embryonic development. For example, coronary artery SMC are derived from proepicardial cells (1, 2), whereas SMC of the aortic arch and thoracic aorta are derived partly from the neural crest (3–5). In the peripheral vasculature, SMC are recruited from the surrounding mesenchyme by endothelial cells; similarly, in the gut a primitive epithelial tube recruits SMC from the surrounding mesenchyme (6). With these diverse origins, it is likely that distinct nuclear factors are involved in regulating smooth muscle development and differentiation in different smooth muscle tissues.

Differentiated smooth muscles are characterized by the presence of a set of unique isoforms of contractile proteins, ion channels, and signaling molecules that are necessary for the contractile properties of the tissue. Several groups have investigated the mechanisms regulating the expression of unique smooth muscle-specific isoforms of contractile proteins, and experiments using transgenic mice have identified minimal promoter regions that are necessary to mediate smooth muscle-specific expression of several of these genes. For example, studies have demonstrated that a 16-kb fragment of the smooth muscle myosin heavy chain extending from -4.2 to +11.6 kb (7), a -2,600 to +2,784 bp fragment of the smooth muscle {alpha}-actin gene (8), a 13.7-kb (-2.7 to +11 bp) fragment of the smooth muscle {gamma}-actin gene (9), and a 370-bp (-190 to +180 bp) fragment of the telokin gene (10, 11) are required to mimic expression of the corresponding endogenous genes. In addition, transgenes driven by 435 bp of the proximal SM22{alpha} promoter, although not mimicking endogenous SM22{alpha} expression, were restricted to arterial SMC in adult mice (12–14). Analysis of the pattern of expression of these transgenes and other truncated constructs has suggested that there are distinct regulatory modules that control expression of genes in different smooth muscle tissues (11). Progress has also been made in identifying some of the major transcription factors that regulate the expression of smooth muscle-specific genes, although their tissue-specific roles have, thus far, been poorly defined. Transcription factors identified to regulate smooth muscle-specific genes include serum response factor (SRF), myocyte enhancer factor 2B (MEF2B), MEF2C, myocardin, myocardin-related factor-A (MRTFA, MLK1, MAL), GATA family members, GATA 4/5/6, Krupple-like zinc finger proteins such as Sp1/3, BTEB3, single-stranded DNA-binding proteins Pur{alpha} and Pur{beta}, and homeodomain proteins such as Gax, Hex, Nkx3.1, Nkx3.2, Barx1b, Barx2b, and Hoxb7 (for review, see Refs. 15 and 16).

Homeodomain (Hox)-containing transcription factors play a crucial role in organogenesis and pattern formation during embryogenesis and regulate proliferation, differentiation, and migration in multiple cell types (17). In addition, expression of these proteins often persists in adult tissues where they play a role in cell type specification. Homeobox genes were first identified in Drosophila as genes whose mutations caused body segment transformation, or homeotic transformation (18). Homeobox genes encode evolutionary conserved transcription factors with a common 60-amino acid DNA-binding motif that folds into three {alpha}-helices and is referred to as the homeodomain (19). In addition to the clustered Hox genes, there are many additional proteins in mammals which contain a homeodomain and play important roles in development. Several of these homeodomain proteins, including Mhox(Prx1), Nkx3.1, Barx1b, and Barx2, have been proposed to play a role in regulating expression of smooth muscle genes through their ability to interact with SRF and promote its DNA binding activity (20–23). The expression of homeodomain-containing proteins has been characterized in vascular SMC (for review, see Ref. 24). For example, the homeodomain protein Gax is expressed at high levels in differentiated vascular smooth muscle, and its expression is down-regulated during phenotypic modulation and dedifferentiation. It has also been shown that Gax is important to induce cell cycle arrest in vascular SMC through its activation of p21CIP-1 (25). Several other Hox genes, including HOXA5, HOXA11, HOXB1, HOXB7, and HOXC9 were found to be expressed in fetal human vascular SMC, of these only Hoxb7 and Hoxa11 were detected in adult mouse smooth muscle tissues (in intestine and uterus, respectively) (26). Hoxa2, Hoxa4, Hoxa5, and Hoxb7 have been cloned from an adult rat vascular smooth muscle cDNA library, although the functions of these proteins were not determined in this study (27). Subsequently, Hoxb7 has been shown to increase expression of SM22{alpha} in 10T1/2 cells (28). Clustered Hox genes have been shown to be important for vascular development and have been implicated in several pathological processes such as arterial restenosis after balloon dilatation, and abnormal expression of homeobox genes has also been found to contribute to infertility and sterility in humans (29).

Although Hox proteins have been implicated in the regulation of smooth muscle differentiation, the downstream targets of these proteins are poorly defined. To elucidate further the role of Hox genes in adult SMC, we used degenerate RT-PCR and cDNA library screening to identify Hox proteins expressed in the smooth muscle of adult mouse bladder and then examine the role of these proteins in regulating smooth muscle-specific genes. Clones encoding Hoxa10 and Hoxb8 were isolated from this screen. Reporter gene cotransfection studies demonstrated that Hoxa10-1 activated the smooth muscle-specific telokin promoter in fibroblasts. siRNA-mediated knock-down of Hoxa10-1 demonstrated that Hoxa10-1 plays an important physiological role in activating telokin expression in SMC. In contrast, Hoxb8 repressed the telokin promoter and several other smooth muscle-specific promoters in vascular SMC. Gel mobility shift assays demonstrated binding of Hoxa10-1, Hoxa10-2, and Hoxb8 to four AT-rich regions in the telokin promoter. One of these regions includes the CArG box. which binds SRF, and Hox binding to this region is abolished by SRF binding. Mutational analysis of the telokin promoter revealed that each of the other three AT-rich regions in the telokin promoter is required for Hoxa10-1 and Hoxb8 to regulate telokin promoter activity. Taken together, these data demonstrated a critical role of clustered homeodomain proteins in regulating smooth muscle-specific genes.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
RT-PCR and cDNA Library Screening—Total RNA was isolated from mouse bladder using a single step guanidinium isothiocyanate procedure. mRNA was isolated from total RNA using magnetic oligo(dT) beads (Invitrogen). Prior to use poly(A)+ mRNA was treated with DNase to remove genomic DNA contamination. cDNA was generated from poly(A)+ mRNA using random primers and murine leukemia virus-reverse transcriptase. Hox cDNA was then amplified by PCR using degenerate sense and antisense oligonucleotides (Hox sense, CTRGARCTRGARAARGARTTYCAYTT; and Hox antisense, RTTYTGRAACCARATYTTWACYTG), derived from helix 1 and 3 of the Hox domain. PCR was carried out for 35 cycles, using an annealing temperature of 51 °C. Gel-purified PCR products were then used as probes to screen a mouse bladder cDNA library in {lambda}gt11. Library filters were hybridized overnight at 65 °C with a 32P-labeled probe. Filters were washed in 2 x SSPE (0.36 M NaCl, 0.02 M NaH2PO4, and 0.002 M EDTA, pH 7.7) + 1.0% SDS at room temperature for 15 min, 2 x SSPE + 1.0% SDS at 65 °C for 15 min followed by 0.2 x SSPE + 0.1% SDS at 65 °C for 10 min. {lambda}-DNA was isolated using Lambdasorb (Promega, Madison, WI), digested with EcoRI, and the resulting fragments were subcloned into pGEM 7Z (Promega) and sequenced by automated sequencing.

Construction of Hoxa10, Hoxb8, and Hoxb7 Mammalian Expression Vectors—The Hoxa10-1 cDNA isolated from the {lambda} library extended from base 206 through 2450 and was thus missing 198 bp of 5'-coding sequence. To generate a full-length clone, a 603-bp fragment at the 5'-end of the Hoxa10 gene was amplified from genomic DNA using PCR. Oligonucleotides were created from the published Hoxa10 sequence. The amplified 5'-end was joined to the 3'-portion of Hoxa10-1 cDNA at a common SacII restriction site to generate full-length cDNA. The full-length Hoxa10-1 was ligated into the pcDNA3 HisC mammalian expression vector (Invitrogen). The integrity of cDNA was confirmed by sequencing. The coding region of Hoxa10-2 was amplified by PCR from the cDNAs isolated from the bladder cDNA library and cloned in-frame into a modified pShuttle vector (Clontech) that included an amino-terminal hemagglutinin epitope tag. Because the Hoxb8 clones isolated from our library did not contain the entire coding sequence, a 740-bp fragment of mouse Hoxb8, including the entire coding region, was isolated by RT coupled to PCR from mouse bladder mRNA. RT was performed using random primers and SuperScript reverse transcriptase (Invitrogen). PCR was then performed using Deep Vent DNA polymerase and the following Hoxb8-specific primers: sense, actcagaatgagctcttatttcgtcaactc; antisense, aggatcctacttcttgtcacccttctgcgcatc. After sequence confirmation the Hoxb8 coding region was cloned in-frame into pcDNAHisC (Invitrogen) or pShuttle (Clontech) for expression experiments. A Hoxb7 cDNA was amplified from Image clone 4413080 (Invitrogen) and ligated into the pShuttle expression vector (Clontech).

Reporter Gene Constructs—Promoter-luciferase constructs were generated in the pGL2BorpGL3B luciferase vectors (Promega) as described previously (10). Promoter fragments used were -256 to +147 of the rabbit telokin promoter (T400 (10)), -4,200 to +11,600 of the smooth muscle myosin heavy chain gene (7), -2,555 to +2,813 of the smooth muscle {alpha}-actin gene (8), -435 to +60 of the SM22{alpha} gene (11), and -113 to +20 of thymidine kinase gene (TK). For mutational analysis of the telokin promoter, deletion of the AT-rich regions was generated using a -82 to +82 fragment of the mouse telokin promoter as a template. All mutant reporter gene constructs were initially generated in pCRBlunt vector (Invitrogen) by a QuikChange mutagenesis kit (Stratagene) and then transferred to the pGL2B luciferase reporter vector. The resultant plasmids were sequenced to verify the integrity of the insert.

Cell Transfection and Reporter Gene Assays—Plasmids were transfected into A10 vascular SMC and 10T1/2 embryonic fibroblast cells using FuGENE 6 (Roche Applied Science). 10T1/2 cells were grown in high glucose Dulbecco's modified Eagle's medium (Roche Applied Science) containing 5 units/ml penicillin, 50 µg/ml streptomycin, and 10% fetal bovine serum. A10 cells were grown in the same medium, but fetal bovine serum was increased to 20%. A10 smooth muscle and 10T1/2 fibroblast cells were seeded at a density of 2.5 x 104/well in a 24-well plate. 16–18 h postseeding cells were washed once with phosphate-buffered saline, pH 7.4, and replaced with 0.5 ml of complete medium. Cells were then incubated with a total of 1 µg of plasmid DNA (0.25 µg of promoter-pGL2B, 0.5 µg of various expression plasmids or empty vectors, and 0.25 µg of pRLTK-Renilla luciferase as an internal control) and 2 µl of FuGENE 6 in 50 µl of Dulbecco's modified Eagle's medium. 24 h after transfection, extracts (100 µl/well) were prepared for measurement of luciferase activity using a dual luciferase assay as described by the manufacturer (Promega). The level of firefly luciferase activity was normalized to the control Renilla luciferase activity. Measurements were made from a minimum of six independent transfections, and all assays were repeated at least twice. Results were reported as the mean ± S.E., and all variables were analyzed by t test, and significance was set at p ≤ 0.01.

Gel Mobility Shift Assays—Gel mobility shift assays were conducted in a 15-µl final volume. 0.2 ng (15 x 103 counts/min) of 32P-labeled double-stranded DNA probe was added to the binding mix. The binding mix also contained 200 ng of poly(dI-dC), 4.5 µg of bovine serum albumin, and different amounts of recombinant protein diluted in binding buffer containing 12 mM HEPES, pH 7.9, 60 mM KCl, 4 mM MgCl2, 10% glycerol, and 1 mM dithiothreitol, as needed. The mix was incubated for 15 min at room temperature during the binding reaction. For antibody supershift assays, incubations were extended for 1 additional h on ice, after the addition of the appropriate antibody. Antibodies used were anti-MEF2{alpha}, SRF, Omni, and SP1, obtained from Santa Cruz (Santa Cruz, CA), and anti-hemagglutinin, obtained from Covance (Richmond, CA). In addition, unlabeled double-stranded DNA probes were used as competitors in some experiment as indicated in the figure legends. The sequences of the sense strand of each probe used in gel mobility shift experiments were as follows: -90 to -53 core wild type probe that encompasses the -80 to -67 5'-AT-rich region and -65 to -56 CArG box, 5'-GCTTTATATAAACTATCCCTTTTATGGGAGCT-3'; -90 to -67 probe that encompass the -80 to -67 5'-AT-rich region, 5'-CGATCTGCAGTTGCTTTATATAAACTAT-3'; -69 to -55 CArG probe, 5'-CGATATCCCTTTTATGGG-3'; -2 to +23 region that encompasses the +2 to +6 3'-AT-rich region and the +14 to +17 region, 5'-ACTGTCACATTAACTCGCACATCAGTTCCA-3'; -2 to +23 region with +14 to +17 deleted, 5'-CATGCACATTAACTCGCACGTTCCA-3'; -2 to +23 region with +2 to +6 deleted, 5'-CATGCACCTCGCACATCAGTTCCA-3'; -2 to +35 region with +2 to +6 and +14 to +17 mutations, 5'-ACTGTCACGCGCGCTCGCACGTCGGTTCCAGAACCCATTCCA-3'; AP2, 5'-CCGATCGAACTGACCGCCCGCGGCCCGT-3'; core wild type CArG mut (-90 to -53), 5'-GCTTTATATAAACTATCCCTTTTCTACGAGCT-3'; -68 to -48 probe with CArG mut, 5'-CGATATCCCTTTTCTACGAGCTGAA-3'. Annealed oligonucleotides were labeled with [{alpha}-32P]-dCTP by Klenow DNA polymerase. Free [{alpha}-32P]dCTP was removed by agarose gel electrophoresis. The DNA-protein complexes were resolved by electrophoresis through 4% polyacrylamide gels containing 6.5 mM Tris, pH 7.9, 3.3 mM sodium acetate, pH 7.9, 1 mM EDTA, and 2.5% glycerol. The gel was then dried and exposed to photographic film with intensifying screens at -80 °C.



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  		FIG. 1.
Expression of Hoxa10 and Hoxb8 in mouse tissues and cells. RNase protection analyses were performed using probes specific for Hoxa10-1 (A), Hoxa10-2 (B), and Hoxb8 (C), respectively, as indicated. Differences in the sequence of Hoxa10-2 in mouse and rat resulted in a partial protection of the A10-2 probe by rat Hoxa10-2 mRNA. In D, RT-PCR was conducted using RNA isolated from mouse bladder, colon, liver, and brain, from a colon smooth muscle cell line (colon SMC), rat embryo fibroblasts (REF), 10T1/2 fibroblasts, A10 vascular SMC, and primary uterine SMC (UT-SMC). A pair of unique primers for Hoxb8 was designed as sense 5'-CTGGTGCAGTACGCAGACTGCAAGC-3' and anti-sense 5'-CCACTTCATTCTCCGATTCTGGAACC-3', yielding a 288-bp product. 400 ng of RNA was utilized as a template for RT-PCR with the Hoxb8-specific primers using the SuperScript One-step RT-PCR system (Invitrogen).

 
Protein Expression and Western Blotting—For preparation of extracts for gel mobility shift assays COS cells were plated at a density of 1.8 x 106/100-mm dish. 16–20 h postplating, cells were transfected with appropriate expression plasmids using FuGENE 6. 16 µg of plasmid and 32 µl of FuGENE 6 were used for each dish of cells. 24 h post-transfection cells were washed three times on ice with rinsing buffer containing 40 mM Tris, pH 7.5, 1 mM EDTA, and 0.15 M NaCl. COS cells were then scraped in 1 ml of rinsing buffer and transferred to microcentrifuge tubes. Cells were collected by centrifugation and resuspended in 200 µl of resuspension buffer mix containing 40 mM HEPES, 0.4 M KCl, 1 mM dithiothreitol, 0.1 mg/ml phenylmethylsulfonyl fluoride, 0.1 mg/ml aprotinin, and 0.02 mg/ml leupeptin. The cells were then lysed by freeze thawing three times, the lysates clarified by centrifugation for 5 min, and the supernatant was stored at -80 °C in aliquots. Protein concentrations were determined using the Bradford protein determination method. The expression of recombinant proteins was verified by Western blotting as described previously (21).

Reverse Transcription Coupled to PCR—Total RNA was isolated as described previously (30). 400 ng of RNA template was used for RT and PCR with specific primers using the SuperScript One-step RT-PCR system (Invitrogen). Unique primers were designed for Hoxa10-1, telokin, and glyceraldehyde-3-phosphate dehydrogenase as follows: Hoxa10-1 sense, 5'-AGCGAGTCCTAGACTCCACGCCACC-3', and antisense, 5'-TCACTTGTCTGTCCGTGAGGTGGACG-3', yielding a 339-bp product; telokin sense, 5'-GACACCGCCTGAGTCCAACCTCCG-3', and antisense, 5'-GGCTTTTCCTCAGCAACAGCCTCC-3', yielding a 214-bp product; glyceraldehyde-3-phosphate dehydrogenase sense, 5'-GCAGTGGCAAAGTGGAGATTGTTGCC-3', and antisense, 5'-GGAGATGATGACCCTTTTGGCTCCAC-3', yielding a 294-bp product.



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  		FIG. 2.
Effects of Hoxa10, Hoxb8, and Hoxb7 on the activity of smooth muscle-specific promoters. 0.5 µg of Hoxa10-1 (A) or Hoxb8 (B) expression plasmid was cotransfected together with 0.25 µg of promoter-luciferase reporter genes containing either the -256 to +147 fragment of the rabbit telokin promoter (T400), -2,555 to +2,813 fragment of the smooth muscle {alpha}-actin promoter ({alpha}-actin), -435 to +60 fragment of the SM22{alpha} promoter (SM22), -4,200 to 11,600 fragment of the smooth muscle myosin heavy chain promoter (MHC), or -113 to +20 fragment of the thymidine kinase promoter (TK) and 0.25 µg of the TK promoter-driven Renilla luciferase internal control plasmid into 10T1/2 fibroblast cells (A) or A10 SMC (B), respectively. 0.5 µg of Hoxa10-1, Hoxa10-2, Hoxb8, Hoxb7, or empty (control) expression vector plasmid was cotransfected together with 0.25 µg of T400 and 0.25 µg of the TK promoter-driven Renilla luciferase internal control plasmid into 10T1/2 fibroblast cells (C) or A10 SMC (D). The level of promoter activity was determined by measuring the level of firefly luciferase activity relative to the control Renilla luciferase. -Fold activation or repression of the promoter activity relative to vector control transfections is shown as the mean ± S.E. of a minimum of six samples. * indicates values that are statistically different from controls, p < 0.01.

 
Adenovirus-mediated Expression of siRNA Directed against Hoxa10 —A plasmid-based system for the production of siRNA was generated by inserting oligonucleotides specific to Hoxa10-1 CAATGTCATGCTCGGAGAG downstream of an H1 promoter in the adenoviral shuttle vector pRNAT-H1.1/Shuttle (GenScript, Piscataway, NJ). This shuttle vector is compatible with the Adeno-X adenoviral system Clontech and was used to generate adenovirus expressing the siRNA as described previously (30). For adenoviral infection, primary mouse uterine and colonic SMC were prepared as described previously (31), seeded in 6-well plates, and allow to grow to a density of 2–2.5 x 105 cells/well. Primary cells were washed with phosphate-buffered saline and infected with Hoxa10-1 siRNA or control siRNA adenovirus in phosphate-buffered saline for 4 h at 37 °C. 72 h after infection protein extracts were prepared using radioimmune precipitation assay buffer (1% Nonidet P-40, 1% sodium deoxycholate, 0.1% SDS, 0.15 M NaCl, 2 mM EDTA, 0.01 M sodium phosphate, pH 7.2) containing protease inhibitors. Protein concentrations of the extracts were determined using the BCA protein assay. For RT-PCR analysis total RNA was extracted using TRIzol reagent as described previously (30).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Hoxa10 and Hoxb8 Are Expressed in Adult Mouse Bladder— RT-PCR, using degenerate oligonucleotides prepared to the first and third {alpha}-helixes of the conserved homeodomain of mouse Hox genes, amplified a 110-bp fragment from mRNA isolated from mouse bladder. The cDNAs present in this fragment were gel purified and used as probes to screen a {lambda}gt11 cDNA library generated from mouse bladder. Five different positive clones were obtained from the library screen, three represented Hoxa10, and two represented Hoxb8. Each of the Hoxa10 clones represented a distinct isoform. The Hoxa10-1 clone obtained was 2,244 bp and extended from 206–2450 of the published mouse Hoxa10-1 sequence. This clone was missing 198 bp of the 5'-coding sequence, hence, PCR was used to obtain the missing 5'-end and generate a full-length Hoxa10-1 expression construct as described under "Experimental Procedures." A 1,600-bp clone encoding the entire coding sequence of Hoxa10-2 and a 752-bp fragment of Hoxa10-3 was obtained. Hoxb8 clones were 317 bp and ~1,067 bp, extending from 1,539 to 1,856 and from 1,440 to 2,507 bp, respectively, of the published Hoxb8 cDNA. A full-length Hoxb8 coding region (1,059–1,787) was isolated by RT-PCR using mouse bladder mRNA as a template as described under "Experimental Procedures."



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  		FIG. 3.
Effect of siRNA-mediate knock-down of Hoxa10-1 on endogenous telokin gene expression in primary SMC. Primary uterine SMC (A and B) and primary colonic SMC (C) were transduced with control siRNA or Hoxa10-1 siRNA adenovirus for 4 h. 72 h after infection protein extracts were prepared and analyzed by Western blotting for endogenous telokin and SRF protein expression (B), or RNA was harvested from cells using TRIzol reagent, and RT-PCR was performed to detect endogenous Hoxa10-1 (A and C) and telokin (C). The glyceraldehyde-3-phosphate dehydrogenase (GAPDH) housekeeping gene was used as an internal control showing equal RNA input and the efficiency of the RT-PCR.

 



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  		FIG. 4.
Mapping the Hox binding sites on the telokin promoter. A, schematic diagram showing the CArG box (-65 to -56), 5'-AT-rich region (-80 to -71), and the 3'-AT-rich region (+2 to +6) and the potential Pbx binding site (+14 to +17) on the telokin promoter. B and C, electrophoretic mobility shift assays conducted using a probe that encompasses a -90 to -53 region of the telokin promoter including both the -80 to -71 AT-rich region and the -65 to -56 CArG box. A single DNA-protein complex was seen when reactions included 0.5 µg of Hoxa10-1 extract, and two complexes were seen when 1.7 µg of Hoxa10-1 nuclear extract was added (B). Both complexes were supershifted with the {alpha}-Omni antibody directed against the epitope on Hoxa10-1 but not with nonspecific antibody. In C reactions included a 25-fold excess of unlabeled competitor oligonucleotides encompassing the entire -90 to -53 region (Core WT), this same region with three point mutations in the CArG box (CoreCArG-mut), the -90 to -67 region (E-TATA), the CArG box (-70 to -55), a mutant CArG box (CArG mut), or a consensus AP2 site, as indicated. Of these competitors only the AP2 consensus site was unable to compete for binding to Hoxa10-1. D, electrophoretic mobility shift assay conducted with a probe encompassing the CArG box (-70 to -55). 1.7 µg of Hoxa10-1 nuclear extract was added together with increasing amounts of SRF (0, 50, 150, and 200 ng of SRF). The identity of the protein in each mobility-shifted complex was confirmed using specific antibodies to supershift the complexes, as indicated. E and F, electrophoretic mobility shift assays were conducted using probes that included either the -90 to -67 region (E) or the -2 to +23 region (F).

 
Hoxa10 and Hoxb8 mRNA Are Expressed at a High Level in Adult Smooth Muscle Tissues and Cells—RNase protection analysis of total RNA isolated from adult mouse tissues revealed expression of Hoxa10-1 at high levels in uterus and bladder and at lower levels in colon, kidney, skeletal muscle, and aorta (Fig. 1A). Hoxa10-1 was also detected in intestinal SMC (colon SMC) (Fig. 1A). RNase protection analysis of Hoxa10-2 expression showed that Hoxa10-2 is expressed in a pattern similar to that of Hoxa10-1 with the exception that Hoxa10-2 but not Hoxa10-1 was detected in A10 vascular SMC (Fig. 1, A and B). RNase protection analysis revealed high levels of Hoxb8 mRNA in uterus, colon, and kidney and lower levels in bladder and aorta (Fig. 1C). In addition, RT-PCR data indicated that Hoxb8 is also expressed in primary uterine SMC and a colon smooth muscle cell line (Fig. 1D).

Hoxa10-1 Activates and Hoxb8 Represses Smooth Muscle-specific Promoters—To determine whether Hoxa10 and Hoxb8 directly regulate the expression of smooth muscle-restricted proteins, expression constructs were cotransfected together with smooth muscle-specific promoter reporter genes into A10 vascular smooth muscle and 10T1/2 fibroblast cells, and the effects on luciferase activity were determined. Results from these experiments demonstrated that in 10T1/2 cells Hoxa10-1 increased the activity of the telokin promoter 3-fold without affecting the activity of the other promoters analyzed (Fig. 2A). Similar results were also observed in A10 SMC (data not shown). In contrast, Hoxb8 significantly repressed the activity of the telokin, smooth muscle {alpha}-actin, and SM22{alpha} promoters by 70, 50, and 70%, respectively, without significantly altering the activity of the smooth muscle myosin or TK kinase promoters in A10 cells (Fig. 2B). Hoxb8 had little effect on the activity of these promoters in 10T1/2 cells (data not shown). In contrast to Hoxa10-1, Hoxa10-2 had no significant effect on telokin promoter activity in 10T1/2 cells (Fig. 2C) and resulted in a 30% reduction of promoter activity in A10 cells (Fig. 2D). Sequence analysis indicated that Hoxa10-3 does not encode for a homeodomain protein, and the protein product of this mRNA has not been defined, hence Hoxa10-3 was not pursued further in this study. Because Hoxb7 has also been suggested to play a role in regulating smooth muscle-specific genes we also examined the effects of Hoxb7 on telokin promoter activity. This analysis revealed that Hoxb7 had little effect on telokin promoter activity in A10 cells (Fig. 2D) and no effect in 10T1/2 fibroblasts (Fig. 2C).



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  		FIG. 5.
Mapping the Hox binding sites within the -2 to +23 region of the telokin promoter. Electrophoretic mobility shift assays were performed using either a full-length probe encompassing the -2 to +23 region or truncated probes in which either the +2 to +6 AT-rich region was deleted (-2 to +23 (ATTA-D)) or the +14 to +17 region was deleted (-2 to +23 (ATCA-D)), or both regions mutated in the context of a probe that extended from -2 to +35 (-2 to +35 (All-M)). Probes were incubated with in vitro transcribed and translated Hoxa10-1 (A) or Hoxb8 (B), and specific complexes were supershifted with antibodies to the Omni epitope tag (*) but not by nonspecific antibodies (SP1).

 
siRNA-mediated Knock-down of Hoxa10-1 Decreases Endogenous Telokin Gene Expression in Primary Visceral SMC—To examine the physiological role of Hoxa10-1 in regulating endogenous telokin gene expression, primary cultures of visceral SMC, which express both Hoxa10-1 and telokin, were infected with Hoxa10-1 siRNA to reduce endogenous levels of Hoxa10-1. RT-PCR analysis demonstrated that Hoxa10-1 siRNA decreased expression of both endogenous Hoxa10-1 and telokin without affecting expression of glyceraldehyde-3-phosphate dehydrogenase (Fig. 3, A and C). Similarly, Western blot analysis revealed a significant down-regulation of telokin protein expression in primary uterine cells infected with Hoxa10-1 siRNA but not with control siRNA (Fig. 3B). These data indicate a critical physiological role of Hoxa10-1 in regulating endogenous telokin gene expression in SMC.

Hoxa10-1 Binds to Multiple Sites in the Core of the Telokin Promoter—To determine whether the physiological effects of Hoxa10-1 on telokin promoter activity result from direct binding of Hoxa10-1 to the telokin promoter, gel mobility shift assays were performed. Because homeodomain-containing proteins are known to bind to AT-rich sequences and previous studies have shown that an AT-rich region and adjacent CArG box are important for telokin promoter activity (32), gel mobility shift assays were first used to determine whether Hoxa10-1 bound to the -90 to -53 AT-rich/CArG-containing region of the telokin promoter (Fig. 4). When high amounts of Hoxa10-1 extract were utilized, two mobility-shifted complexes were seen on this probe (compare 0.5 µg and 1.7 µg of extract in Fig. 4B). Both mobility-shifted complexes were supershifted by antibody to the Omni epitope tag on the Hoxa10-1 protein but not by irrelevant antibodies (Fig. 4B). The presence of two Hoxa10-1-containing complexes suggests that there are at least two binding sites for Hoxa10-1 within this region of the telokin promoter. Consistent with this proposal, competition experiments with unlabeled oligonucleotides corresponding to either the 5'-half of the probe (-84 to -66 AT-rich region) or the 3'-half of the probe (CArG, -69 to -55) demonstrated that both of these oligonucleotides can compete for Hoxa10-1 binding to the AT-rich/CArG probe (Fig. 4C). In addition, gel mobility shift assays demonstrated a Hoxa10-1 mobility-shifted complex on probes derived from either the CArG or -84 to -66 AT-rich regions (data not shown). Previously we have shown that SRF binds to the CArG element of the telokin promoter (32). Experiments were therefore performed to determine whether both SRF and Hoxa10-1 could simultaneously bind to this element. These experiments demonstrated that SRF competes with Hoxa10-1 for binding to the CArG probe, suggesting that in the presence of SRF, Hoxa10-1 does not bind to the CArG element (Fig. 4D).

Similar to Hoxa10-1 both Hoxa10-2 and Hoxb8 were shown to bind to the -84 to -66 AT-rich region (Fig. 4E). Sequence analysis of the telokin promoter fragment, utilized in reporter assays described in Fig. 2, revealed the presence of two other potential homeodomain protein binding sites between residues -2 and +23. Gel mobility shift assays using a probe that encompasses this -2 to +23 region demonstrated that Hoxa10-1, Hoxa10-2, and Hoxb8 can also bind to this region of the promoter (Fig. 4F). To determine whether both of the potential binding sites within this probe can bind to Hoxa10-1 or Hoxb8, mobility shift assays were performed using probes with deletions of either the +2 to +6 or the +14 to +17 Hox sites. These assays demonstrated that Hoxb8 can bind to both homeodomain protein binding sites in this region, but Hoxa10-1 can only bind to the +2 to +6 site (Fig. 5).

Hoxa10-2 and Hoxb8 Competitively Inhibit the Activity of Hoxa10-1—Because Hoxa10-1 and Hoxa10-2 share an identical DNA binding domain it would be anticipated that they would compete for binding to similar DNA binding sites. Consistent with this model, gel mobility shift assays demonstrated direct competition between Hoxa10-1 and Hoxa10-2 for binding to the -82 to -69 Hox site within the telokin promoter (Fig. 6A). Because Hoxa10-2 lacks the amino-terminal transcription activation domain found in Hoxa10-1, these data would also predict that Hoxa10-2 would competitively inhibit the transcription activation caused by Hoxa10-1. To test this possibility directly, reporter gene assays were performed in which the telokin promoter was activated with a constant amount of Hoxa10-1, and the effects of increasing concentrations of Hoxa10-2 were determined. These assays revealed that the activation of the telokin promoter by Hoxa10-1 could be reversed by overexpression of Hoxa10-2 (Fig. 6B). Although Hoxb8 binds to the same sites in the telokin promoter as Hoxa10-1, it inhibits promoter activity. This would suggest that Hoxb8 may also act, in part, by competing with Hoxa10-1. This proposal is supported by the observation that Hoxb8 can also block the activation of the telokin promoter induced by Hoxa10-1 (Fig. 6C). In contrast to Hoxb8, Hoxb7 did not significantly block the ability of Hoxa10-1 to activate the telokin promoter (Fig. 6D).



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  		FIG. 6.
Hoxa10-2 and Hoxb8 but not Hoxb7 block the telokin promoter activation induced by Hoxa10-1. A, electrophoretic mobility shift assay conducted using the -90 to -67 probe that encompasses the -81 to -71 AT-rich region of the telokin promoter. The probe was incubated with a constant amount of Hoxa10-1 (1 µg) together with an increasing amount of Hoxa10-2 nuclear extract (0, 0.5, 1, 1.5, 2, 2.5, and 3 µg). Control COS cell extract was added to maintain equivalent total protein input into each reaction. The Hoxa10-1 complex was reduced significantly as the levels of the Hoxa10-2 complex increased. B, C, and D, telokin promoter luciferase reporter gene assays were performed in 10T1/2 cells. Reporter genes were transfected into cells together with a constant amount of Hoxa10-1 expression plasmid and increasing amounts of Hoxa10-2 (B), Hoxb8 (C), or Hoxb7 (D) expression plasmids. Total plasmid concentration was kept constant through the addition of empty expression vector. The level of promoter activity was determined by measuring the level of firefly luciferase activity relative to the control Renilla luciferase. -Fold activation or repression of the promoter activity relative to vector control transfections is shown as the mean ± S.E. of six samples. * indicates values significantly different from Hoxa10-1 alone, p < 0.01.

 
Hoxa10-1 and SRF Synergistically Activate the Telokin Promoter—Because gel mobility shift assays demonstrated that SRF and Hoxa10-1 compete for binding to the CArG box in the telokin promoter we examined the effects of coexpression of these proteins on telokin promoter activity. Cotransfection of SRF and Hoxa10-1 into 10T1/2 cells was found to result in synergistic activation of the telokin promoter (Fig. 7A). In contrast, cotransfection of SRF with Hoxa10-2 or Hoxb8 resulted in no significant change in the activation of the promoter compared with the activation induced by SRF alone (Fig. 7, B and C).

Multiple Binding Sites Are Required for Hoxa10-1 and Hoxb8 Regulation of the Telokin Promoter—Gel mobility shift assays demonstrated that Hoxa10-1, Hoxa10-2, and Hoxb8 bind to multiple sites in the telokin promoter (Fig. 4). To determine which of these binding sites are important for the Hox proteins to regulate telokin promoter activity, telokin promoter reporter genes with deletions in each of the Hox sites were generated and analyzed. Cotransfection experiments showed that deletion of any one of the Hox sites resulted in a significant inhibition of the ability of Hoxa10-1 to activate the promoter (Fig. 8A). Similarly, all three Hox sites were required for Hoxb8 to repress telokin promoter activity maximally (Fig. 8B).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In the current study we identified two homeodomain proteins, Hoxa10 and Hoxb8, which are expressed in adult smooth muscle tissues, and we demonstrated that the promoters of smooth muscle-restricted genes are direct transcriptional targets of these proteins. Hoxa10 specifically activates the telokin promoter and is required for telokin expression in uterine and colonic SMC. In contrast, Hoxb8 represses the activity of several smooth muscle-restricted promoters. These data suggest that clustered homeodomain proteins play an important role in regulating expression of genes in smooth muscle tissues and cells and that different homeodomain proteins have very distinct effects on the promoters of smooth muscle-specific genes.



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  		FIG. 7.
Hoxa10-1 and SRF synergistically activate the telokin promoter, whereas Hoxa10-2 and Hoxb8 do not affect the SRF-dependent activation of the promoter. Telokin promoter reporter genes were transfected into 10T1/2 cells together with Hoxa10-1, SRF, Hoxa10-2, or Hoxb8 expression plasmids as indicated. In A, cells were transfected with either 0.25 µg of Hoxa10-1, 0.25 µg of SRF, or 0.25 µg of each plasmid together with empty expression vector where appropriate, to maintain equal total plasmid concentrations. In B and C, cells were transfected with 0.2 µg of SRF expression plasmid together with increasing amounts of Hoxa10-2 (B) or Hoxb8 (C) expression plasmid and empty vector to maintain equal total plasmid concentrations. The level of promoter activity was determined by measuring the level of firefly luciferase activity relative to the control Renilla luciferase. -Fold activation or repression of the promoter activity relative to vector control transfections is shown as the mean ± S.E. of a minimum of six samples. * indicates values that are statistically different from controls, p < 0.01.

 
RNase protection analysis indicated that Hoxa10-1 transcripts are expressed at high levels in smooth muscle tissues of the uterus and bladder and at lower levels in kidney. In addition, Hoxa10-1 is also expressed in primary cultures of SMC and in a colonic smooth muscle cell line. Previous studies indicated the presence of all three Hoxa10 transcripts during embryonic development with increased expression from day 9 to day 12 followed by a decrease in expression by day 15 (33). In situ hybridization demonstrated expression of Hoxa10-1 and Hoxa10-2 in the posterior somites, neural tube, and the intermediate mesoderm of the herniated gut and the urogenital system, in day 12.5 mouse embryos. This pattern of embryonic expression suggests a possible role of Hoxa10 in regulating the development of the hind gut and urogenital tract and coincides with the induction of telokin expression in these tissues (34). This observation together with our findings that Hoxa10-1 activates the telokin promoter and that down-regulation of Hoxa10-1 by siRNA in uterine and colonic SMC results in decreased telokin expression (Fig. 3) suggests that Hoxa10-1 may be involved in the induction of telokin during smooth muscle development in these tissues. Hoxa10 has also been identified previously in endometrial epithelial cells and myometrial SMC (35). Several studies have indicated that Hoxa10 is important for the process of uterine epithelium cell differentiation and the development of subcellular structures required for implantation (36). In addition, the expression of Hoxa10 in the myometrium is regulated during the menstrual cycle, with high levels of estrogen increasing Hoxa10 expression (37). Previously we have shown that telokin expression is also regulated during the menstrual cycle with high levels of expression being observed during the estrogen-dominant follicular phase and low levels of expression during the progesterone-dominant luteal phase. Telokin expression is also elevated in the uterus of pregnant women (38). Reporter cotransfection experiments demonstrated that both the estrogen receptor and Hoxa10-1 can activate the telokin promoter (Fig. 2A and Ref. 38), suggesting that these two proteins may synergize with each other to mediate hormone-dependent changes in telokin expression in the uterus. Because telokin has been proposed to play a role in regulating smooth muscle relaxation (39) the elevation of telokin expression during pregnancy may contribute to the maintenance of uterine quiescence during pregnancy.

Although Hoxa10-1 may play an important role in regulating telokin expression in uterus, colon, and bladder, telokin is also expressed in a number of smooth muscle tissues that have undetectable expression of Hoxa10-1 such as jejunum, stomach, and lung (Fig. 1). In these other tissues it is probable that other homeodomain proteins that are expressed in more anterior regions of the body substitute for the function of Hoxa10-1. For example, the paralogous group 4 Hox genes have been found to be expressed in more anterior portions of the intestinal tract, including the stomach and small intestine (40). Additional studies will be needed to determine whether members of this group of Hox genes regulate expression of telokin in more anterior regions of the gut.

The three Hoxa10 transcripts isolated from mouse bladder in the current study are identical to those reported previously (33). The Hoxa10-1 transcript uses a distinct 5'-exon compared with the Hoxa10-2/3 transcripts, and Hoxa10-2 and Hoxa10-3 transcripts are distinguished by alternative splice donor sites in exon 1 (33). Because the first exons of Hoxa10-1 and Hoxa10-2/3 are distinct, this would suggest that these transcripts result from the activity of independent promoters within the gene. Only two transcripts, Hoxa10-1 and Hoxa10-2, are capable of producing homeodomain-containing proteins. A potential open reading frame in Hoxa10-3 is not in the correct reading frame to encode the homeobox domain, and the protein product of this mRNA has not been identified. The Hoxa10-1 transcript encodes a homeodomain protein of 55 kDa (399 amino acids) with a 325-amino acid amino-terminal region that contains a proline/glutamine-rich domain similar to those shown to function as transcriptional activators. Hoxa10-2 encodes a homeodomain protein of 16 kDa (96 amino acids) with a small amino-terminal domain of 20 amino acids that lacks the proline/glutamine-rich transcriptional activation domain. The coexpression of Hoxa10-1 and Hoxa10-2 in tissues, and in at least one cell line derived from colonic SMC, has important implications for the physiological effects of these homeodomain proteins. Because Hoxa10-2 competes with Hoxa10-1 for the same binding sites (Fig. 6) and Hoxa10-2 blocks the ability of Hoxa10-1 to activate a promoter, this would suggest that the relative ratio of Hoxa10-1 and Hoxa10-2 in cells will likely determine the level of gene activation induced by Hoxa10-1. The coexpression of an activator and repressor in the same cell type is a phenomenon seen with many proteins, for example alternative splicing generates SRF isoforms that can act as competitive inhibitors of full-length SRF (41). Similarly, alternative splicing of the focal adhesion kinase gene leads to the production of focal adhesion kinase-related non-kinase, which competitively inhibits the activity of focal adhesion kinase (42). The ability of a SMC to express either Hoxa10-1, an activator of telokin gene expression, or Hoxa10-2, a repressor of telokin gene expression, likely allows the cells to fine tune the expression of telokin by independently regulating the expression of the two Hox proteins.



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  		FIG. 8.
Mapping the Hox sites in the telokin promoter required for Hoxa10-1 and Hoxb8 activity. A mouse telokin promoter reporter gene extending from -82 to +82 (0.25 µg) or the indicated deletion mutants of this fragment, and TK-Renilla control plasmid (0.25 µg) were transfected into 10T1/2 cells together with a Hoxa10-1 expression vector (0.5 µg) (A) or into A10 cells together with a Hoxb8 expression vector (B). The level of promoter activity was determined by measuring the level of firefly luciferase activity relative to the control Renilla luciferase. -Fold activation or repression of the promoter activity relative to vector control transfections is shown as the mean ± S.E. of six samples. Relative luciferase activities that were significantly different from the activity of Hoxa10-1 (A) or Hoxb8 (B) on the -82 to +82 wild-type reporter gene (bottom bars) are indicated by the asterisks, p < 0.01.

 
The -83 to -73 region of the telokin promoter (TGCTTTATATAAACTA) resembles the previously defined consensus binding site for Hoxa10 (AA(A/T)TTTATTAC), with the TTTAT core recognition sequence being entirely conserved. There are no consensus binding sites for the PBC family of homeodomain proteins adjacent to the Hoxa10 binding site, suggesting that Hoxa10 likely binds to this site in the telokin promoter alone rather than as a complex with PBC proteins. However, there is a consensus binding site for Pbx, a member of the PBC family of proteins, between +12 and +16 adjacent to the Hoxa10-1 binding site between +2 and +6 (ATTAA). Because deletion of this consensus Pbx binding site between +12 and +16 prevented Hoxa10-1 from activating the telokin promoter (Fig. 8), yet Hoxa10-1 itself does not bind to this site (Fig. 5), this would suggest that Hoxa10-1 bound to the +2 to +6 Hox site may require heterodimerization with Pbx for Hoxa10-1 to activate transcription. In support of this proposal, previous studies have shown Pbx interactions with Hox proteins B1 through B9 and HoxA10, and these studies also showed that this interaction is critical for restricting the DNA binding specificity of Hox proteins (43).

Hoxb8 strongly repressed the activity of several smooth muscle-specific promoters, including the telokin promoter. Within the telokin promoter, Hoxb8 inhibited promoter activity, at least in part through its ability to block promoter activation induced by Hoxa10-1. However, because we did not detect any endogenous Hoxa10-1 expression in A10 cells and Hoxb8 expression in these cells repressed telokin promoter activity, this would suggest that Hoxb8 represses telokin promoter activity through an independent mechanism. One such mechanism could be through competition with other homeodomain protein activators that may be expressed in A10 cells. Alternatively, interaction of Hoxb8 with Pbx may recruit corepressor molecules to inhibit promoter activity. Similar to its role in inhibiting smooth muscle-specific genes Hoxb8 has been shown to prevent myeloid differentiation through a mechanism that requires interaction with Pbx on target gene promoters. The expression of both Hoxa10-1 and Hoxb8 in the same SMC (colon SMC, Fig. 1) would again suggest that either the relative ratio of Hoxa10-1 and Hoxb8 will determine the level of telokin gene activation or that the activity of these proteins is modified post-transcriptionally. In support of the latter model, tyrosine phosphorylation of Hoxa10 has been shown to decrease in vitro binding of Hoxa10 to a Pbx-Hoxa10 binding site (44). Alternatively the interaction of the Hox proteins with other factors, such as Pbx or Mies proteins, may alter their ability to regulate the promoters of smooth muscle-specific genes.

Our findings that Hoxb8 repressed the activity of several smooth muscle promoters is perhaps surprising in light of a previous study that demonstrated that Hoxb7 increased expression of SM22{alpha} in 10T1/2 cells (28). However, we also demonstrated that Hoxb7 has little effect on telokin promoter activity (Fig. 2) or on the activity of the -435 to +60 bp SM22{alpha} promoter (data not shown). This may suggest that Hoxb7 binds to a region of the SM22{alpha} promoter not included in our reporter gene or that the ability of Hoxb7 to induce SM22{alpha} in stably transfected 10T1/2 cells may have resulted from indirect activation of the promoter by Hoxb7. For example, Hoxb7-mediated activation of basic fibroblast growth factor (45) may have stimulated the proliferation of a subpopulation of 10T1/2 cells that had elevated expression of SM22{alpha}. Our findings, together with the observed expression of Hoxb7 in atherosclerotic plaques and the ability of Hoxb7 to increase smooth muscle cell proliferation (28) may suggest that Hoxb7 is more important for regulating the proliferation of SMC rather than regulating the expression of smooth muscle contractile proteins directly.

In summary, our data demonstrate that the promoters of smooth muscle-restricted genes are direct transcriptional targets of clustered homeodomain proteins and that these proteins have distinct effects on different smooth muscle-restricted promoters. Hoxa10-1 specifically regulates telokin gene expression and together with direct estrogen activation through the estrogen receptor, may play a particularly important role in maintaining high levels of telokin expression in uterine smooth muscle, thereby helping to keep the uterus quiescent during pregnancy.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants HL58571, DK61130, and DK65644 (to B. P. H.). 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. Back

{ddagger} To whom correspondence should be addressed: Dept. of Cellular and Integrative Physiology, Indiana University School of Medicine, 635 Barnhill Dr., Indianapolis, IN 46202. Tel.: 317-278-1785; Fax: 317-274-3318; E-mail: pherring{at}iupui.edu.

1 The abbreviations used are: SMC, smooth muscle cells; Hox, homeodomain; RT, reverse transcription; siRNA, small interfering RNA; SRF, serum response factor; TK, thymidine kinase. Back


   ACKNOWLEDGMENTS
 
We are grateful to April Hoggatt and Ketrija Touw for expert technical assistance and to members of the Herring laboratory for helpful discussions.



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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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