Sp1 is required for transcriptional activation of the fibroblast growth factor receptor 1 gene in neonatal cardiomyocytes

Gene 400 (2007) 150 – 157 www.elsevier.com/locate/gene

Sp1 is required for transcriptional activation of the fibroblast growth factor receptor 1 gene in neonatal cardiomyocytes Mahdie Seyed, Joseph X. DiMario ⁎ Rosalind Franklin University of Medicine and Science, The Chicago Medical School, Department of Cell Biology and Anatomy, 3333 Green Bay Road, North Chicago, Illinois USA 60064 Received 26 April 2007; received in revised form 4 June 2007; accepted 5 June 2007 Available online 19 June 2007 Received by J.A. Engler

Abstract Fibroblast growth factor receptor 1 (FGFR1) is the predominant FGFR in cardiac tissue and regulates proliferation, differentiation, and maintenance of normal myocardium. During development of cardiac tissue, FGFR1 gene expression regulates cardiomyocyte proliferation. The focus of this study was to determine the molecular mechanism of transcriptional activation of the FGFR1 gene in proliferating neonatal cardiomyocytes. Analysis of DNA sequence of the FGFR1 gene identified three potential Sp factor binding sites located at 49 bp, 68 bp, and 100 bp upstream from the 3′ end of the promoter segment. Mutation of each of these sites resulted in a significant decline in FGFR1 promoter activity compared to wild type promoter activity, and combinatorial mutation of all three sites completely abrogated promoter activity to background levels. In addition, overexpression of Sp1 in neonatal cardiomyocytes resulted in a dose-dependent increase in wild type FGFR1 promoter activity. However, Sp1-mediated up-regulation of promoter activity was abrogated when all three Sp interacting sites were mutated. Chromatin immunoprecipitation (ChIP) assays were used to demonstrate direct interactions of Sp1 with the proximal promoter region of the FGFR1 gene in neonatal cardiomyocytes. ChIP assays using Drosophila Schneider Line 2 (SL2) cells transiently transfected with wild type or mutant FGFR1 promoter constructs verified the direct interaction between Sp1 and the three Sp1 interacting sites of the promoter. Western blot analyses indicated that Sp1 was present in cytoplasmic and nuclear extracts of neonatal myocardium. These results indicate that Sp1 is a necessary positive regulator of FGFR1 gene transcription in neonatal cardiomyocytes. © 2007 Elsevier B.V. All rights reserved. Keywords: FGFR1; Promoter; Proliferation; Muscle

1. Introduction Fibroblast growth factors (FGF) and their tyrosine kinase receptors play a significant role in proliferation and differentiation of numerous cell types. Since their first discovery more

Abbreviations: FGFR1, fibroblast growth factor receptor 1; bp, base pair; ChIP, chromatin immunoprecipitation; SL2, Schneider Line 2; FGF, fibroblast growth factor; HSPG, heparan sulfate proteoglycans; Ig, immunoglobulin; PLC, phospholipase C; PKC, protein kinase C; AV, atrioventricular; ES, embryonic stem; CMV, cytomegalovirus; PCR, polymerase chain reaction; SDS, sodium dodecyl sulfate; BS, blocking solution; PBS, phosphate buffered saline; HRP, horseradish peroxidase; SV40, simian virus 40; kb, kilobase; luc, luciferase; SEM, standard error of the mean; Ab, antibody. ⁎ Corresponding author. Tel.: +1 847 578 8633; fax: +1 847 578 3253. E-mail address: [email protected] (J.X. DiMario). 0378-1119/$ - see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.gene.2007.06.010

than 30 years ago as mitogenic factors in pituitary extracts (Gospodarowicz, 1974), over twenty different isoforms of FGF have been identified to date (Claus et al., 2004). These isoforms bind a family of four FGF receptors (FGFR1-4) that dimerize upon ligand binding and are activated by tyrosine autophosphorylation (Dell'Era et al., 2003). It has been proposed that heparan sulfate proteoglycans (HSPG) act as low-affinity coreceptors for FGFs and that formation of a ternary complex of FGF-FGFR-HSPG is essential for dimerization and activation of FGF receptors (Yayon et al., 1991; Rapraeger et al., 1991). Each FGFR monomer has two cytoplasmic tyrosine kinase domains, a single-pass transmembrane domain, and an extracellular immunoglobulin (Ig)-like domain composed of two or three Ig-like loops depending on the splicing of the specific FGFR isoform (Oslen et al., 2004). Autophosphorylated FGFR signaling is mediated via phospholipase C (PLC), protein kinase C (PKC),

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and other target proteins resulting in activation of signaling cascades involved in regulation of cell growth and differentiation in a variety of tissues (Dell'Era et al., 2003; Basilico and Moscatelli, 1992). The important role of FGFRs in differentiation, proliferation, and migration of various cell populations has been investigated previously. In chicken embryos FGFR1 plays a critical role in normal neurulation as well as limb bud formation (Deng et al., 1997). With respect to skeletal muscle development, overexpression of FGFR1 during embryonic stages promoted proliferation of myoblasts, maintained myoblast migration from somites to limb buds, and inhibited terminal differentiation of myogenic precursor cells (Itoh et al., 1996). In vivo, expression of a truncated, dominant negative FGFR1 resulted in a 30% loss of skeletal muscle mass, a 50% reduction in myofiber density, and a reduction in the number of nuclei in myotubes (Flanagan-Steet et al., 2000). In vivo inhibition of FGFR4 signaling also resulted in loss of muscle mass in the affected tissue. In contrast to FGFR1, FGFR4 appears to promote myogenic differentiation (Marics et al., 2002). FGFR2 and FGFR3 are both required for normal formation of cartilage for endochondral osteogenesis (Eswarakumar et al., 2004), and their mutations have been associated with long-bone and skull defects related to craniosynostosis (Wilkie, 1997). The presence of different FGFRs in a variety of cell types and tissues, as well as their role in up-regulation of cell-cycling pathways, has made FGFRs a frequent focus of numerous cancer-related studies (Eswarakumar et al., 2005). In developing heart, expression of FGFRs correlates with the proliferative capacity of cardiomyocytes (Sheikh et al., 1999). FGFR1 is the predominant high-affinity FGFR in avian and murine myocardium. Expression of FGFR2 and FGFR3 is restricted to atrioventricular (AV) cushion mesenchyme and AV endocardial cells, respectively (Sugi et al., 2003). In Drosophila, mutation of the heartless gene, which is homologous to vertebrate FGFR1, resulted in lack of development of the contractile dorsal vessel (Beiman et al., 1996). Also, while fgfr1−/− mice die during gastrulation, fgfr1−/− embryonic stem (ES) cells maintain their potential to differentiate into cardiomyocytes in vitro. However, this ability was lost upon treatment with the FGFR1 tyrosine kinase inhibitor, SU5402. Furthermore, lentivirus-mediated delivery of FGFR1 to fgfr1−/− ES cells rescued cardiomyocyte induction indicating that FGFR1 plays a non-redundant role in cardiogenesis (Dell'Era et al., 2003). As development proceeds from embryonic to postnatal to adult stages, FGFR1 RNA levels in myocardium decline, corresponding to a decrease in the proliferation rate of cardiomyocytes and a transition from a hyperplastic to a hypertrophic phenotype (Sheikh et al., 1999; Engelmann et al., 1993; Liu et al., 1995). The functional significance of FGFR1 activity on cardiomyocyte growth has been demonstrated in several studies. Overexpression of FGFR1 in neonatal rat cardiomyocytes and the H9c2 cardiogenic cell line resulted in increased DNA synthesis and an increased rate of cardiomyocyte proliferation (Sheikh et al., 1997, 1999). Conversely, antiFGFR1 antibodies retarded proliferation and multilayering of cardiogenic cells in in vitro models of cardiac morphogenesis

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(Sugi et al., 1995). These studies confirmed the critical role of FGFR1 in cardiogenesis and cardiomyocyte proliferation. Many growth factor receptor gene promoters share structural similarities. These transcriptional regulatory regions usually lack consensus TATA and CCAAT transcription initiation sites. Also, the GC-rich characteristic of growth factor receptor gene regulatory regions accounts for the numerous Sp transcription factor binding sites present in these regions (DiMario, 2002). Sp transcription factors comprise a family of four C2H2 zinc-finger DNA binding proteins (Sp1-4) that regulate basal and inducible transcription of many genes. They bind to GC boxes (GGGCGG), GT motifs (GGGTGTGGC), or CT elements (CCTCCTCCTCCTCGGCCTCCTCCCC) (Parakati and DiMario, 2004). Sp1, Sp2, and Sp4 often function as activators of promoter activity, whereas Sp3 can function as either a transcriptional activator or repressor (Hagen et al., 1994; Li et al., 2004). Here, we report the positive regulatory role that Sp1 plays in controlling the level of FGFR1 promoter activity in neonatal cardiomyocytes by binding to three identified Sp binding site elements in the proximal promoter. 2. Materials and methods 2.1. Cell culture Neonatal ventricular cardiomyocytes were isolated from day 1 postnatal Sprague-Dawley rats using Worthington's Cardiomyocyte Isolation System. Cells were plated in collagen coated 35 mm dishes at a density of 1 × 106 cells/dish or in 100 mm dishes at a density of 6 × 106 cells/dish. Growth medium contained 10% horse serum (Hyclone), 10% fetal bovine serum (Hyclone), 1.32 mM CaCl2, 1 × glutamine (Gibco), and 1 × penicillin/ streptomycin/Fungizone (Gibco) in F10 base medium (Gibco). Cells were cultured at 37 °C in a 5% CO2 humidified incubator. Drosophila SL2 cells (ATCC) were cultured in 75% Drosophila SFM/25% Schneider's Drosophila medium (Gibco) supplemented with 2 mM L-glutamine and 1 × antibiotic-antimycotic (Gibco) at 25 °C without CO2. After 3–4 days growth in culture medium, cells were plated at a density of 2.5 × 106 cells/60 mm dish for transfection the next day. 2.2. Transfections Cardiomyocytes were transfected with 6 μg of wild type or mutated promoter constructs linked to the luciferase gene using Lipofectamine Plus reagent (Invitrogen). Transfection of the promoterless pGL3-Basic (Promega) vector DNA served as a negative control. Transfection of 300 ng of Renilla luciferase construct (pRL-SV40; Promega) controlled for transfection efficiencies. After 24 h, firefly and Renilla luciferase assays were performed using the Dual-Glo Luciferase Assay System (Promega). The expression constructs, pCMVSp1 and pCMVSp3, were used to drive expression of Sp1 and Sp3, respectively, based on constitutive activity of the cytomegalovirus (CMV) promoter (Shen et al., 1995). Transfection of pCMVSp1 and pCMVSp3 into cardiomyocytes was performed as described above with two modifications. First, increasing amounts of

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pCMVSp1 were added to the transfection mixture, and pKS BlueScript (Stratagene) was used to bring the total DNA up to 8 μg per plate. For these experiments, luciferase assays were performed 48 h after transfection. Transfection of Drosophila SL2 cells occurred in Drosophila SFM medium using the Cellfectin reagent (Invitrogen). Expression of Sp1 in SL2 cells was transcriptionally regulated by the Drosophila actin promoter in the expression construct, pPacSp1 (Courey and Tjian, 1988). Each plate received 300 ng of pRL-SV40, increasing amounts of pPacSp1, 3 μg of promoter-luciferase reporter plasmid as well as pKS BlueScript (Stratagene) DNA to bring the total DNA up to 8 μg/plate. Luciferase assays were performed after 48 h. For all transfections, a minimum of 3 independent experiments were performed. 2.3. Mutagenesis The wild type FGFR1 promoter-luciferase construct (wtFGFR1-Luc) was used as the template DNA and the following mutagenic forward primers were used with their complementary anti-sense oligonucleotides: m49FGFR1: 5′G C G A G G C T C T C C A G A G TATA G A C C G C A G C - 3 ′ m64FGFR1: 5′-CACCCGCCGCGCCACTATAAGCTCTCCAGAGGCGG-3′ m68FGFR1: 5′-CACAGACACCCGCCGCGTAATGGCGAGCTCTCCA-3′ m72FGFR1: 5′-CACAGACACCCGCCGTATAACGGCGAGCTCTCCA-3′ m100FGFR1: 5′-GCCCGGAGACGAGCGTATAGAGCGCAAGA-3′. The bold, italicized letters represent nucleotide substitutions. The templates were PCR amplified using the mutagenic oligonucleotides as primers, and template DNAwas digested with DpnI restriction enzyme. Mutations were confirmed by DNA sequencing. 2.4. Western blot analysis For subcellular fractionation, hearts were minced on ice and homogenized in 0.5 ml of buffer A (10 mM HEPES, pH 7.9, 50 mM NaCl, 0.5 M sucrose, 1 mM EDTA, 0.5 mM spermidine, 0.15 mM spermine, 0.5% Triton X-100, protease inhibitors (Complete Mini, Roche)). Samples were centrifuged at 1000 ×g for 10 min at 4 °C, and the supernatant was collected as cytoplasmic extract. Nuclear pellets were washed with buffer A and resuspended in 50 μl of buffer B (10 mM HEPES, pH 7.9, 25% glycerol, 0.1 mM EDTA, 0.5 mM spermidine, 0.15 mM spermine, protease inhibitors). NaCl was added to a final concentration of 400 mM. Samples were incubated on ice for 30 min with frequent agitation, and then centrifuged at 2000 ×g for 10 min at 4 °C. Supernatant was collected as nuclear extract. Protein content was determined by BCA protein assay (Pierce).

For western blots, proteins were resolved in 7.5% SDSpolyacrylamide gels and transferred onto nitrocellulose membranes. The membranes were then incubated overnight at 4 °C in blocking solution (BS) containing 5% nonfat dry milk in phosphate buffered saline (PBS) with 0.05% Tween-20. Subsequently the membranes were incubated with rabbit polyclonal antibodies directed against individual Sp transcription factors (Santa Cruz Biotech), diluted 1:1000 in BS, antiE47 rabbit polyclonal antibody (Santa Cruz Biotech), diluted 1:1000 in BS, or monoclonal mouse anti-α-actin antibody (Sigma), diluted 1:5000 in BS, for 1 h at room temperature. After three washes with PBS containing 0.05% Tween-20, the membranes were incubated with horseradish peroxidase-(HRP) conjugated anti-rabbit IgG or HRP-conjugated goat anti-mouse IgM secondary antibodies (1:5000 dilution) for 1 h at room temperature. The membranes were washed 5–6 times as above, and the immunocomplexes were detected using the Super Signal ® Chemiluminescent Substrate (Pierce). 2.5. Chromatin immunoprecipitation Assays were performed as previously described (Parakati and DiMario, 2002). The following forward and reverse primers flanking the Sp binding sites within the FGFR1 promoter were used for PCR amplification of immunoprecipitated DNA from neonatal cardiomyocytes: 5′-CCGGGGCACCAGCTTCGGCTCCATTGTTC-3′ and 5′-GCTGACTCTCACTTGGCGCTGCGGTC-3′. For transfected SL2 cells, primers were designed to amplify transfected DNA and not genomic DNA. The following forward and reverse primers were used: 5′-GATCGAGGTACCTCCGGGGCACCAGCTTCGG-3′ and 5′GCTGCGGTCCCGCCTCTGGAGAGCTCGC-3′. PCR products were electrophoresed in 1% agarose gels. 2.6. Statistics Statistical comparisons were performed using unpaired t-tests with the level of significance set at 0.01. 3. Results 3.1. Sequence analysis on the proximal 240 base pairs of FGFR1 promoter The DNA sequence of the mouse FGFR1 promoter is shown in Fig. 1. The sequence extends 240 bp upstream from the initiating codon. Seventy-two percent of the sequence is comprised of GC base pairs, making it a GC-rich region like

Fig. 1. Sequence analysis of the proximal FGFR1 promoter. Potential Sp transcription factor binding sites were identified in the proximal 240 bp of the FGFR1 promoter. These sites, located at − 49 bp, − 68 bp, and −100 bp, are underlined.

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other growth factor receptor promoters (DiMario, 2002). GCrich transcriptional control regions often contain binding sites for the Sp family of transcription factors. Sequence analysis of the proximal FGFR1 promoter via TFSEARCH software and the TRANSFAC database identified three potential Sp binding sites on the promoter. The sites are most similar to consensus GC boxes (GGGCGG) that interact with Sp transcription factors. The sites are located at − 49 bp, − 68 bp, and − 100 bp upstream from the initiation codon. 3.2. Functional assessment of the potential transcription factor binding sites To assess the transcriptional activity of the FGFR1 promoter, the wild type DNA sequence shown in Fig. 1 was inserted upstream of the firefly luciferase reporter gene. To determine whether the potential Sp transcription factor binding sites at − 49 bp, − 68 bp, and − 100 bp were functional, a series of functional mutagenesis experiments was performed. FGFR1 promoter constructs were generated harboring mutations that replaced 4 bp GC-rich segments with A and T nucleotides (Fig. 2A). Mutant constructs were named using the number of the first 3′ base of the corresponding DNA site. Wild type and mutated FGFR1 promoter-reporter constructs were transfected into neonatal cardiomyocytes. Mutant promoter activities were compared to wild type promoter activity (Fig. 2B). Mutation of the potential sites at − 49 bp, −68 bp, and − 100 bp reduced FGFR1 promoter activity to 70%, 39%, and 64% of wild type promoter activity, respectively. To control for any non-specific effects of mutagenesis on FGFR1 promoter activity, two additional mutations, m64FGFR1 and m72FGFR1, flanking the − 68 bp site, were generated. These mutations yielded promoter activities of 96% and 103% which were not significantly different from the level of wild type FGFR1 promoter activity. Combinatorial mutations of the functional sites were also generated. The combined mutations of the − 49 bp and − 100 bp sites (m49/m100FGFR1) and of all three functional sites (m49/m68/m100FGFR1) further reduced FGFR1 promoter activity to 24% and 21% of wild type promoter activity, respectively. These promoter activities were not significantly different from activity generated from the promoterless pGL3Basic plasmid vector. These results indicate that the three sites at − 49 bp, − 68 bp, and − 100 bp interact with positive transcriptional regulators of the FGFR1 promoter.

Fig. 2. Effect of site-specific mutations on FGFR1 promoter activity. A. Mutant promoter constructs (e.g. m49FGFR1) fused to the luciferase (Luc) reporter gene were generated by substitution of GC-rich potential regulatory sites in the wild type FGFR1 (wtFGFR1) promoter with AT-rich sequences. In each case, 4 bp substitutions were introduced. B. Neonatal cardiomyocytes were transfected with 3 μg of either wtFGFR1-Luc or mutant reporter plasmids along with 300 ng of pRL-SV40 to control for transfection efficiency. Dual luciferase assays were performed 24 h after transfection to determine the level of promoter activity. The pGL3Basic vector, lacking the FGFR1 promoter, was used to control for background levels of luciferase activity. Asterisks indicate mutations that resulted in a significant change in the level of promoter activity compared to wtFGFR1 promoter activity ( p b 0.0001). Promoter activities of m49FGFR1, m68FGFR1, m100FGFR1, m49/m100FGFR1, and m49/m68/ m100FGFR1 were reduced compared to wtFGFR1 promoter activity. Activities of the m64FGFR1 and m72FGFR1 promoters were not significantly different from wtFGFR1 promoter activity. Activities of the double and triple mutant promoters (m49/m100FGFR1 andm49/m68/m100FGFR1) were not significantly different than activity from the promoterless pGL3Basic vector. Bars represent mean promoter activities ± S.E.M. (n = 9).

nuclear versus cytoplasmic protein extractions and provided protein loading controls between protein samples.

3.3. Sp1 and Sp3 are present in the nuclei of neonatal cardiomyocytes

3.4. Sp1 activates the FGFR1 promoter

To determine which Sp transcription factors were present in neonatal cardiomyocytes, nuclear and cytoplasmic protein extracts were obtained from neonatal rat cardiomyocytes and western blotted. Sp1-4 were detected with Sp1-4-specific antibodies (Fig. 3). Sp1 and Sp3 were detected in nuclear and cytoplasmic extracts of neonatal cardiomyocytes. Sp4 was detected only in the cytoplasmic fraction, and Sp2 was not detected in either nuclear or cytoplasmic fractions. Western blots in which E47 and α-actin were immunodetected with their respective antibodies verified

To investigate the ability of Sp1 and Sp3 transcription factors to regulate FGFR1 promoter activity in cardiomyocytes, cultures of neonatal cardiomyocytes were transfected with the wild type FGFR1 promoter reporter construct, wtFGFR1-Luc, as well as increasing amounts of the Sp1 or Sp3 expression vectors, pCMVSp1 or pCMVSp3. Expression of Sp1 increased FGFR1 promoter activity in a dose dependent manner (Fig. 4A). Transfection of 4 μg of pCMVSp1 increased FGFR1 promoter activity by approximately 2.5 fold. To verify that the increase of

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promoter, neonatal cardiomyocytes were co-transfected with promoter constructs harboring mutations in one or more of these elements along with the pCMVSp1 expression construct. Promoter activities were determined 24 h after co-transfection (Fig. 5). Co-transfection of 4 μg of pCMVSp1 increased wtFGFR1 promoter activity as before (Fig. 4). Each mutation of the FGFR1 promoter significantly reduced Sp1-mediated transcriptional activation. The m49FGFR1, m68FGFR1, and m100FGFR1 promoters demonstrated reductions in Sp1mediated transcriptional activation of 44%, 79%, and 47%, respectively. Combinatorial mutations of m49/m100FGFR1 and

Fig. 3. Immunodetection of Sp1 and Sp3. Western blot analysis was performed to determine which Sp transcription factors were expressed in neonatal cardiomyocytes. Equal amounts (60 μg) of nuclear and cytoplasmic extracts were resolved in a 7.5% SDS polyacrylamide gel. Proteins were transferred to nitrocellulose membranes, and Sp1-4 were detected with Sp transcription factor specific rabbit polyclonal antibodies followed by a horseradish peroxidase conjugated secondary antibody and chemiluminescence. Detection of the nuclear transcription factor E47 and the cytoplasmic α-actin in nuclear and cytoplasmic fractions, respectively, served as protein loading controls.

FGFR1 promoter activity was the result of expression of the Sp1 transcription factor and independent of the pCMV promoter, cultures of neonatal cardiomyocytes were transfected with 4 μg of an empty pCMV vector. FGFR1 promoter activity did not change in the presence of the empty pCMV vector (Fig. 4B). To determine whether the effect of Sp1 on transcriptional activation of the FGFR1 promoter was specific to Sp1, the Sp3 expression construct, pCMVSp3, was transfected into neonatal cardiomyocytes. Expression of Sp3 did not activate the FGFR1 promoter (data not shown). Drosophila SL2 cells lack endogenous Sp transcription factors and therefore can function as a neutral cell type with respect to Spmediated transcriptional regulation. The function of individual Sp factors can be assessed in a controlled manner without potential interfering or complicating effects by endogenous Sp factors. This cell line has proven useful for studying the role of the Sp family of proteins in the transcriptional regulation of multiple genes (Parakati and DiMario, 2002; Courey and Tjian, 1988). Co-transfection of SL2 cells with wtFGFR1-Luc and the Sp1 transcription factor expression construct, pPacSp1, yielded similar results to those observed with cardiomyocytes (Fig. 4C). Transfection of 4 μg of the pPacSp1 expression vector increased wild type FGFR1 promoter activity by approximately 4 fold. The greater effect of Sp1 expression on the level of FGFR1 promoter activity in SL2 cells compared to cardiomyocytes is likely due to lack of endogenous Sp1 in SL2 cells. In total, these results indicate that Sp1 can activate FGFR1 transcriptional activity in cardiomyocytes. 3.5. Mutation of the positive regulatory sites negates FGFR1 promoter activation by Sp1 To investigate the functional significance of each of the three positive transcriptional regulatory elements of the FGFR1

Fig. 4. Effect of Sp1 expression on FGFR1 promoter activity. A. Cardiomyocytes were transfected with 3 μg of wtFGFR1-Luc reporter plasmid along with variable amounts of pCMVSp1 (0, 1, 2, 3, or 4 μg). To keep the total amount of DNA constant, pKS Bluescript plasmid was used to bring the total DNA up to 8 μg / plate. Transfection efficiencies were normalized by co-transfection of 300 ng of pRL-SV40. Relative promoter activities were determined 24 h after transfection and compared to FGFR1 promoter activity in the absence of pCMVSp1. Cotransfection of increasing amounts of pCMVSp1 increased FGFR1 promoter activity in a dose-dependent manner (p b 0.001; n = 9). B. To control for any nonspecific effects due to transfection of the CMV promoter DNA, neonatal cardiomyocytes were transfected with 3 μg of wtFGFR1-Luc along with 0 μg or 4 μg of empty pCMV vector DNA. pKS-Bluescript was used to bring the total DNA content up to 8 μg /plate. Transfection of the empty pCMV vector DNA did not significantly affect the level of FGFR1 promoter activity (n = 3). C. Drosophila SL2 cells were co-transfected with 3 μg of wtFGFR1-Luc and increasing amounts (0–4 μg) of pPacSp1. pKS Bluescript DNAwas used to bring the total amount of DNA up to 8 μg /plate, and pRL-SV40 (300 ng) was used as an internal control for transfection efficiency. Transfection of increasing amounts of pPacSp1 increased FGFR1 promoter activity in SL2 cells in a dose-dependent manner with approximately a 4 fold increase in promoter activity in the presence of 4 μg of the Sp1 expression vector ( p b 0.01;n = 3).

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Fig. 5. Effect of mutations of the FGFR1 promoter on Sp1-mediated transcriptional activation. Neonatal cardiomyocytes were co-transfected with 3 μg of wtFGFR1-Luc or mutated FGFR1 promoter-luciferase reporter constructs along with 4 μg pCMVSp1. Promoter activities were normalized by co-transfection of pRL-SV40. Each mutation significantly (asterisk) reduced Sp1-mediated activation of the FGFR1 promoter ( p b 0.001; n = 3). m49FGFR1, m68FGFR1, and m100FGFR1 retained 56%, 21%, and 53% of the Sp1mediated transactivation potential, respectively. The m49/m100FGFR1 retained only 16% of Sp1-mediated transactivation potential, and the m49/m68/ m100FGFR1 promoter was not significantly activated by expression of Sp1.

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To further demonstrate that the interaction between Sp1 and the FGFR1 promoter is specific to the three identified Sp1 binding sites on the promoter, transient chromatin immunoprecipitation assays were performed using wild type and mutant FGFR1 promoter constructs in Drosophila SL2 cells. Cultures of Drosophila SL2 cells were transfected with either wtFGFR1Luc or mutant promoter plasmids as well as the pPacSp1 expression vector. ChIP analysis was performed 48 hours after transfection. The wild type FGFR1 construct was immunoprecipitated with the Sp1 antibody and PCR amplified. The same controls were used for both neonatal cardiomyocytes and SL2 cells. FGFR1 promoter DNAs containing individual Sp site mutations (m49FGFR1, m68FGFR1, and m100FGFR1) were also immunoprecipitated with the Sp1 antibody. Also, the double site mutation m49/m100FGFR1 was immunoprecipitated. However, mutation of all three Sp1 binding sites on the FGFR1 promoter eliminated immunoprecipitation of DNA fragments detectable by PCR amplification when using the Sp1 antibody. These results indicate that Sp1 directly interacts with the FGFR1 promoter in neonatal cardiomyocytes.

m49/m68/m100FGFR1 further negated Sp1-mediated transactivation by 84% and 91%, respectively. Activity of the triple m49/m68/m100FGFR1 mutant promoter co-transfected with 4 μg pCMVSp1 was not significantly different than activity in cardiomyocytes transfected with the mutant promoter and no pCMVSp1. These results indicate that the three identified transcriptional regulatory sites potentially interact with Sp1 and thereby activate the FGFR1 promoter in cardiomyocytes. Furthermore, since mutation of all three sites completely abrogated Sp1-mediated transcriptional activation, it is likely that no other unidentified, potential Sp binding site significantly mediates Sp1 activation of the promoter. 3.6. Sp1 interacts with the FGFR1 promoter in vivo To determine whether the Sp1 transcription factor interacts with the FGFR1 promoter in vivo, chromatin immunoprecipitation (ChIP) analysis was employed using an anti-Sp1 antibody (Fig. 6). The Sp1 specific antibody immunoprecipitated chromatin from neonatal cardiomyocytes. DNA was PCR amplified using primers that flanked all three Sp binding sites of the FGFR1 promoter, yielding the expected product size of 216 bp. Omission of antibody and inclusion of a non-specific antibody (anti-Gαq) did not result in immunoprecipitation of DNA fragments detectable by PCR amplification. PCR amplification of wtFGFR1-Luc plasmid and input chromatin before immunoprecipitation served as positive controls. The results indicate that Sp1 forms a protein-DNA transcriptional regulatory complex by binding to the FGFR1 promoter in neonatal cardiomyocytes.

Fig. 6. Endogenous and transient chromatin immunoprecipitation analyses of Sp1 binding to the FGFR1 promoter in vivo. A. Chromatin from neonatal cardiomyocytes was immunoprecipitated with the Sp1 antibody (Sp1 Ab) and endogenous FGFR1 promoter DNA was PCR amplified using primers that flanked all three Sp binding sites. Amplification of input chromatin (Input) prior to immunoprecipitation and wtFGFR1 promoter DNA (wtFGFR1) served as positive controls for chromatin extraction and PCR amplification. Chromatin immunoprecipitation using a non-specific antibody (Gαq Ab) or no antibody (No Ab) served as negative controls. B. Transient chromatin immunoprecipitation assays of the wild type and mutant FGFR1 promoter constructs in SL2 cells were also performed. SL2 cells were transiently transfected with wtFGFR1-Luc or one of the mutant FGFR1 promoter constructs along with pPacSp1. Chromatin immunoprecipitation assays, including controls, were performed as above. DNA was amplified using a 5′ flanking primer that hybridized to FGFR1 promoter sequence upstream from the three Sp1 binding sites and a 3′ flanking primer that hybridized to the luciferase gene DNA. The Sp1 antibody immunoprecipitated DNA fragments detectable by PCR amplification from SL2 cells transfected with wtFGFR1 and pPacSp1, indicating that Sp1 binds the FGFR1 promoter construct in transiently transfected SL2 cells. No immunoprecipitated PCR product was detected when all three Sp1 binding sites of the FGFR1 promoter were mutated.

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4. Discussion FGFR1 is involved in the development, growth, and survival of cardiac tissue. During embryogenesis, FGFR1 plays a nonredundant role in the early differentiation of cardiac cells and regulates neonatal cardiomyocyte proliferation (Dell'Era et al., 2003; Sheikh et al., 1999). Adult cardiac tissue, which is generally considered non-proliferative, has a low level of FGFR1 gene expression. Importantly, this low level of expression is responsible for maintaining homeostasis and promoting cardiomyocyte survival after hypoxic injury (Palmen et al., 2004). The focus of this study was the mechanism of transcriptional regulation of FGFR1 gene expression in neonatal cardiomyocytes. Western blot analysis revealed that Sp1 and Sp3 were present in nuclear and cytoplasmic fractions of neonatal cardiomyocytes. Therefore, Sp1 and Sp3 were candidates as transcriptional regulators of the FGFR1 promoter. To determine whether Sp1 or Sp3 activated the FGFR1 promoter, Sp1 and Sp3 were overexpressed in neonatal cardiomyocytes transfected with the wtFGFR1-Luc construct. Expression of the Sp1 transcription factor resulted in a dose dependent increase in FGFR1 promoter activity in neonatal cardiomyocytes. A similar Sp1-mediated dose dependent increase was detected in SL2 cells. The positive regulatory effect of Sp1 on FGFR1 promoter activity was reduced when each of the three Sp1 binding sites was mutated, and it was completely abolished when all three sites were mutated. Furthermore, activation of the FGFR1 promoter was specific to Sp1. These results indicate that Sp1 exerts its positive effect on the level of FGFR1 gene expression by binding to all three identified cis-elements. Interaction of Sp1 with the FGFR1 promoter was critically assessed by chromatin immunoprecipitation (ChIP) assays in neonatal cardiomyocytes, as well as by transient ChIP assays in Drosophila SL2 cells. Endogenous Sp1 was bound to the proximal region of the FGFR1 promoter in neonatal cardiomyocytes. To determine the precise binding sites within the promoter, ChIP assays were performed using FGFR1 promoters with single and multiple Sp binding site mutations. Expressed Sp1 interacted with the FGFR1 promoter in SL2 cells, and mutation of all three Sp1 binding sites was required to abrogate direct interaction between Sp1 and the FGFR1 promoter. These results indicate that Sp1 interacts with all three Sp binding sites within the proximal FGFR1 promoter and that no other direct Sp1 binding site exists within this region of the promoter. The transcriptional control regions of growth factor receptor genes share several structural similarities. The proximal promoter regions are typically GC-rich. For example, the promoters for the rat epidermal growth factor receptor gene, rat transforming growth factor-β receptor type III gene, and human FGFR3 gene are 64%, 69%, and 82% GC-rich, respectively. These promoters also typically lack canonical TATA and CCAAT motifs (DiMario, 2002). Rather, many growth factor receptor transcriptional regulatory regions including the mammalian and avian FGFR1 genes contain several functional Sp binding sites (Parakati and DiMario, 2002, 2004). Both promoters are transcriptionally activated by Sp1, and both

promoters contain three functional Sp1 binding sites within the proximal promoter region. The mouse FGFR1 promoter contains its three Sp1 binding sites within the proximal 100 bp, and the chicken FGFR1 promoter contains its three sites within the proximal 59 bp. In addition, each site confers partial activation to the promoter, although the second upstream site of the mouse FGFR1 promoter at – 68 bp showed the greatest activation potential in contrast to the chicken FGFR1 promoter which was primarily activated by the first (most proximal) site at − 29 bp. The FGFR1 proximal promoter contains several Sp binding sites located 49 bp, 68 bp, and 100 bp within the promoter region. The function of each of these Sp binding sites was assessed by mutagenesis of each site and in combination, followed by promoter activity assays in neonatal cardiomyocytes and Drosophila SL2 cells. Each identified Sp binding site contributed to FGFR1 promoter activity to varying degrees. Mutation of the Sp binding site at − 68 bp reduced promoter activity to a greater extent (39% of wtFGFR1) relative to the sites at − 49 bp and − 100 bp. This suggests that the −68 bp site functions as a strong transcriptional activation element. Interestingly, the mutant promoter m49/m100FGFR1, in which the − 49 bp and − 100 bp sites were mutated and the − 68 bp site was intact, did not display any significant activity. The inability of the − 68 bp site to function as a strong activation element in the context of the − 49 bp and − 100 bp site mutations may be due to cooperativity in transcriptional activation among the three sites. Such synergistic activation may be a common mechanism in Sp1-regulated genes since the transcriptional regulatory regions of these genes often have multiple Sp1 binding sites. The cooperative transcriptional activation of promoters by Sp1 appears to be a mechanism by which not only growth factor receptor genes are regulated but also, more generally, Sp1-regulated genes that have multiple Sp binding sites. For example, the simian virus 40 (SV40) promoter is positively regulated by Sp1 and also contains a cluster of Sp binding sites (Gidoni et al., 1984). Similarly, the human translation termination factor 1 gene has several Sp1 binding sites clustered in the proximal transcriptional control region (Dubourg et al., 2003). In addition to structural and functional similarities with regard to Sp1-mediated regulation of FGFR1 promoter activity, an interesting mechanistic difference also exists. In the mammalian FGFR1 promoter, the three proximal Sp1 binding sites were sufficient to activate the promoter. This is in contrast to the avian FGFR1 promoter in which the three proximal Sp1 binding sites were necessary, but not sufficient, for transcriptional activation. Two additional Sp1 binding sites located more than 2 kb upstream from the proximal sites were required for promoter activity (Patel and DiMario, 2001). Also, these distal sites did not confer incremental activation since mutation of either distal site completely abrogated promoter activity. This configuration of Sp sites into proximal and distal clusters has suggested a model of transcription in which distant DNA regions interact with proximal promoter regions by DNA looping, mediated by Sp1 (Su et al., 1991). Although this mechanism may be operative in the avian FGFR1 promoter, it

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does not appear to be required for activation of the mammalian FGFR1 promoter. It must be acknowledged that additional Sp1 binding sites may be located in the proximal FGFR1 promoter and/or more distal upstream regions, and they may provide further incremental activation. However, if they exist, they do not appear to be necessary for significant promoter activation as observed for the avian FGFR1 promoter. It is possible that the different modes of transcriptional regulation are dependent on cell-specific regulation. The studies focusing on avian FGFR1 promoter regulation used skeletal muscle cell proliferation and differentiation as a model system. In these cells FGFR1 gene expression is down-regulated to undetectable levels after differentiation into muscle fibers. In adult cardiomyocytes, FGFR1 gene expression persists, albeit at reduced levels. Therefore, the different expression profiles of FGFR1 in skeletal versus cardiac muscle may be due to the different modes of FGFR1 promoter activation. Further analysis of the transcriptional repression of the FGFR1 gene as neonatal cardiomyocytes lose proliferative capacity and differentiate into adult cardiomyocytes is needed to clarify the different expression patterns in cardiac versus skeletal muscle. Acknowledgements The Sp1 expression constructs were generous gifts from Dr. G. Suske (University of Marsburg, Germany). References Basilico, C., Moscatelli, D., 1992. The FGF family of growth factors and oncogenes. Adv. Cancer Res. 59, 115–165. Beiman, M., Shilo, B.Z., Volk, T., 1996. Heartless, a Drosophila FGF receptor homolog, is essential for cell migration and establishment of several mesodermal lineages. Genes Dev. 10, 2993–3002. Claus, P., Werner, S., Timmer, M., Grothe, C., 2004. Expression of the fibroblast growth factor-2 isoforms and the FGF receptor 1–4 transcripts in the rat model system of Parkinson's disease. Neurosci. Lett. 360, 117–120. Courey, A.J., Tjian, R., 1988. Analysis of Sp1 in vivo reveals multiple transcriptional domains, including a novel glutamine-rich activation domain. Cell 55, 887–898. Dell'Era, P., Ronca, R., Loco, L., Nicoli, S., Metra, M., Presta, M., 2003. Fibroblast growth factor receptor-1 is essential for in vitro cardiomyocyte development. Circ. Res. 93, 414–420. Deng, C., et al., 1997. Fibroblast growth factor receptor-1 (FGFR-1) is essential for normal neural tube and limb development. Dev. Biol. 185, 42–54. DiMario, J.X., 2002. Activation and repression of growth factor receptor gene transcription. Int. J. Mol. Med. 10, 65–71. Dubourg, C., Toutain, B., LeGall, J.Y., LeTreut, A., Guenet, L., 2003. Promoter analysis of the human translation termination factor 1 gene. Gene 316, 91–101. Engelmann, G.L., Dionne, C.A., Jaye, M.C., 1993. Acidic fibroblast growth factor and heart development: role in myocyte proliferation and capillary angiogenesis. Circ. Res. 72, 7–19. Eswarakumar, V.P., Horowitz, M.C., Locklin, R., Morriss-Kay, G.M., Lonai, P., 2004. A gain-of-function mutation of Fgfr2c demonstrates the roles of this receptor variant in osteogenesis. Proc. Natl. Acad. Sci. 101, 12555–12560. Eswarakumar, V.P., Lax, I., Schlessinger, J., 2005. Cellular signaling by fibroblast growth factor receptors. Cytokine Growth Factor Rev. 16, 139–149.

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