GATA-4 and MEF2C transcription factors control the tissue-specific exp
http://www.100md.com
《核酸研究医学期刊》
Unit of Molecular and Cellular Oncology and 1 Molecular Cell Biology Unit, Department for Molecular Biomedical Research, VIB-Ghent University, B-9052 Ghent-Zwijnaarde, Belgium
* To whom correspondence should be addressed. Tel: +32 9 3313740/3313600; Fax: +32 9 3313609; Email: geert.berx@dmbr.Ugent.be
Present address: Griet Vanpoucke, MRC—Human Reproductive Sciences Unit, 49, Little France Crescent, Edinburgh EH16 4SB, UK
DDBJ/EMBL/GenBank accession no. AF361938
ABSTRACT
T-catenin is a recently identified member of the -catenin family of cell–cell adhesion molecules. Its expression is restricted mainly to cardiomyocytes, although it is also expressed in skeletal muscle, testis and brain. Like other -catenins, T-catenin provides an indispensable link between a cadherin-based adhesion complex and the actin cytoskeleton, resulting in strong cell–cell adhesion. We show here that the tissue-specificity of T-catenin expression is controlled by its promoter region. By in silico analysis, we found that the T-catenin promoter contains several binding sites for cardiac and muscle-specific transcription factors. By co-transfection studies in P19 embryonal carcinoma cells, we demonstrated that MEF2C and GATA-4 each have an activating effect on the T-catenin promoter. Transfections with wild-type and mutant promoter constructs in cardiac HL-1 cells indicated that one GATA box is absolutely required for high T-catenin promoter activity in these cells. Furthermore, we showed that the GATA-4 transcription factor specifically binds and activates the T-catenin promoter in vivo in cardiac HL-1 cells. In vivo promoter analysis in transgenic mice revealed that the isolated T-catenin promoter region could direct the tissue-specific expression of a LacZ reporter gene in concordance with endogenous T-catenin expression.
INTRODUCTION
-Catenins are key molecules of the E-cadherin-mediated cell–cell adhesion complex, because they make the indispensable link to the actin cytoskeleton. The importance of this link to confer strong and functional cell–cell adhesion is illustrated by tumor cells that have lost a functional E-catenin protein, a change that is associated with the loss of cell–cell aggregation and the gain of invasive capacity (1–6). Re-introduction of exogenous functional -catenin results in the restoration of cell–cell adhesion and the inhibition of the invasive capacity in vitro and in vivo (3,6–9). Besides its role as an invasion-suppressor molecule, -catenin has a tumor-growth suppressive capacity. This was recently demonstrated by a conditional knock-out of E-catenin in the epidermis (10). Ablation of E-catenin expression in the skin results in a sustained activation of the Ras-MAPK pathway leading to hyperproliferation of the epidermal cells (10).
Recently, a new member of the -catenin family was identified and termed T-catenin. This novel -catenin shows very high sequence homology to both E- and N-catenins (11). T-catenin expression is restricted to certain tissues. It was first discovered in testis, but also found in cardiac and skeletal muscle and in the brain (11). The distinct expression pattern of T-catenin contrasts with the ubiquitous expression of the closely related E-catenin. In some cell types E- and T-catenin are co-expressed, as is found at the intercalated discs of cardiac muscle cells, whereas in other tissues, for instance in human testis, they are differentially localized (11).
Not much is known so far about the function of this recently identified T-catenin. It has been shown that T-catenin can bind strongly to ?-catenin in heart and testis tissues, and in vitro data show that T-catenin can function as a genuine -catenin by providing a link between a cadherin-mediated cell–cell adhesion complex and the actin cytoskeleton (11). N-cadherin-mediated adhesion is critical for proper myofibril organization in cardiomyocytes (12), and altered cadherin expression in the myocardium leads to dilated cardiomyopathy (DCM) (13). DCM is a ‘cytoskeletalopathy’, as most of the DCM genes identified so far have an influence on actin cytoskeleton organization (14). Interestingly, the gene encoding human T-catenin, CTNNA3, was mapped to chromosome 10q21 (15), a region that has been linked to certain cases of dominantly inherited autosomally DCM (16). In view of its high expression in the heart and its presumed role in anchoring the actin cytoskeleton to the cadherin-mediated cell–cell adhesion complex, T-catenin is considered as a good candidate gene for inherited or sporadic DCM. However, so far no final evidence for such causative link has been found (15). Besides their structural role in cell–cell adhesion, -catenins have also been implicated in signaling events. Similar to other -catenins, T-catenin is also able to inhibit Wnt signaling (17). Based on this finding and on the chromosomal position of T-catenin, a link between T-catenin gene variation and late-onset of Alzheimer disease has been proposed (17,18).
The close relation of -catenin genes and their partly overlapping expression patterns makes understanding the transcriptional regulation of these genes of considerable interest. Moreover, due to the particular expression pattern of T-catenin being restricted to a very limited number of tissues, its promoter must be very tightly regulated and is thus an interesting object to study.
Extensive study of the cardiac-specific gene regulatory network has shown that MEF2C, Nkx2.5 and GATA proteins are key transcriptional regulators in the heart (19). GATA factors are Cys4 zinc finger transcription factors with GATA-4, -5 and -6 playing roles in the development and proper function of the heart . Members of the MEF2 family of MADS-box transcription factors are expressed at high levels in all muscle cells, and MEF2C is essential for proper heart development . Overexpressed MEF2C can initiate cardiomyogenesis in embryonic carcinoma cells (22). In cardiomyocytes, MEF2C acts together with GATA factors to induce gene transcription (23). Apart from their role in heart-specific gene regulation, certain GATA factors and MEF2C are also expressed in other tissues exhibiting T-catenin expression. MEF2C is found in skeletal muscle cells, where it strongly potentiates the transcriptional activity of MyoD, and in post-mitotic neurons in the brain, whereas GATA-4 is involved in gene regulation in the testis (24).
In the present study, we analyze the human T-catenin promoter region both in vitro and in vivo and we determine the role of MEF2C and GATA-factors in the tissue-restricted expression of the T-catenin gene.
MATERIALS AND METHODS
Cloning and sequencing of the human T-catenin promoter
A human bacterial artificial chromosome (BAC) library (Genome Systems Inc.) was screened by a PCR specific for the first exon of the CTNNA3 gene (5'-TGTCATCTGCCTCTCAATTTG-3', 5'-ATGCTGCCTTTCTGTTTCTTC-3'). One positive BAC clone was obtained. Fragments containing exon 1 were identified by Southern hybridization with an exon-1-specific primer and subcloned into the pGEM11 vector to generate pGEM11-hTctnprom. Clones of interest were identified by colony hybridization and sequenced. A sequence of 3412 bp of the human T-catenin promoter was deposited in the GenBank database (GenBank accession no. AF361938 ).
Luciferase reporter plasmids and site-directed mutagenesis
A 3266 bp SacI–SpeI fragment from the pGEM11-hTctnprom construct was blunt-ended and cloned into the blunt-ended HindIII site of the pGL3-Basic Luciferase reporter vector (Promega, Madison, WI) to generate plasmid Tctnprom-luc1.
Site-directed mutations of two potential GATA-binding sites and one putative MEF2C-binding site were introduced into the Tctnprom-luc1 plasmid using the QuickChange site-directed mutagenesis kit (Stratagene). The following primers were used to generate the mutated constructs (mutations are in boldface and are underlined): 5'-TAACCTCCCCTTTCTTTCTTAGGCTGGGTGAACAACGCT-3' and 5'-GAGCGTTGTTCACCCAGCCTAAGAAAGAAAGGGGAGGTTA-3' for GATA box1. 5'-ACTAGCGGTTCAGGATTACCTACCACCCACCCTGGCTTG-3' and 5'-CAAGCCACCGTGGGTGGTAGGTAATCCTGAACCGCTAGT-3' for GATA box2. 5'-GTCTGGCCTTTTTCAACGAAGCACGCTCAGTAACAAGTTGTCAG-3' and 5'-CTGACAACTTGTTACTGAGCGTGCTTCGTTGAAAAAGGCCAGAC-3' for the MEF2C-binding site.
Eukaryotic expression vectors
A GATA-4 expression vector (pcDNA1/GATA-4) and an Nkx2.5 expression vector (pGK-Nkx2.5) were obtained from Dr J.Molkentin (25) and Dr I.Skerjanc (22), respectively. The MEF2C coding sequence was PCR amplified from human heart cDNA (BD Biosciences Clontech, Palo Alto, CA) using primers 5'-GGGACTATGGGGAGAAAAAAGA-3' and 5'-TGTTGCCCATCCTTCAGAAAGT-3', and cloned into the pEF6Ahismyc expression vector (Invitrogen, San Diego, CA).
Cell culture
P19 embryonal carcinoma cells were cultured in -minimal essential medium (Invitrogen) containing 5% newborn calf serum, 5% fetal calf serum, 2 mM L-glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin. HL-1 cardiac cells were cultured as described previously (26).
Transfections and reporter gene assays
Co-transfections in P19 cells were performed using FuGENE 6 Reagent (Invitrogen). These co-transfections included 0.5 μg of the wild-type or mutant T-catenin promoter luciferase reporter constructs combined with 0.5 μg of the pcDNA1/GATA-4, pGK-Nkx2.5 or pEF6Ahismyc-MEF2C expression vectors, in various combinations. Total DNA for each transfection was corrected by the addition of empty expression vector. In each transfection, 0.5 μg of a CMV driven LacZ expression vector (pUT651; Eurogentec) was added as a control for the transfection efficiency. HL-1 cells were transfected using calcium phosphate precipitation in the presence of serum. The calcium phosphate precipitate contained 2 μg of the luciferase reporter plasmid and 1 μg of the pUT651 LacZ expression vector as a control for transfection efficiency. Fresh medium was added to the cells 12 h after transfection.
Forty-eight hours after transfection, the cells were lysed and luciferase and ?-galactosidase activities were measured by adding a luciferase substrate buffer and a ?-galactosidase substrate (Galactostar kit, Tropix Inc., Bedford, MA), respectively.
Nuclear extract preparation and EMSA analysis
Nuclear extracts of HL-1 cells were prepared by controlled NP-40 lysis. Briefly, confluent cell monolayers in 75 mm2 dishes were washed twice with ice-cold phosphate-buffered saline (PBS), harvested by scraping in PBS, and centrifuged at 1500 g for 5 min. The pellets were washed once with PBS and pelleted again as before. The cell pellets were suspended in buffer 1 and the cells were allowed to swell on ice for 15 min. The cells were then lysed by adding 0.6% NP-40 in buffer 1 and vortex-mixed briefly. Lysates were centrifuged at full speed for 15 min and the cytoplasmic fraction was discarded. Nuclei pellets were resuspended in buffer 2 (20 mM HEPES, pH 7.5, 1 mM MgCl2, 400 mM NaCl, 10 mM KCl, 20% glycerol, 0.5 mM EDTA, 0.1 mM EGTA, 2 mM Pefabloc, 0.5 mM DTT and 0.15 IU/ml aprotinin) containing 1% NP-40, vortex-mixed well and centrifuged for 15 min at full speed. The supernatant (nuclear extract) was recovered and dialyzed overnight against 1.6x EMSA binding buffer (20 mM HEPES, pH 7.9, 100 mM KCl, 2 mM DTT, 1 mM EDTA and 20% glycerol) with one buffer change.
Oligonucleotide probes used for EMSA analysis are listed in Table 1. Each reaction mixture (20 μl) contained 12 mM HEPES, pH 7.9, 60 mM KCl, 1.2 mM DTT, 0.6 mM EDTA, 12% glycerol, 0.1% NP-40, 0.05% BSA, 1 μg poly(dI–dC), 2–10 μg nuclear extract and 50 fmol 32P end-labeled probe (10 000 c.p.m.). The mixture was incubated at 30°C for 30 min and loaded directly onto a 6% non-denaturing polyacrylamide gel in 0.5x Tris–borate-EDTA buffer. For competition experiments, cold competitors were added to the reaction mixture simultaneously with the labeled probe. For supershift assays, a GATA-4 antibody (Santa Cruz Biotechnology Inc., Santa Cruz, CA) was added to the DNA/nuclear extract mix, and the reactions were allowed to proceed for 20 min before loading the samples.
Table 1. Wild-type and mutant oligonucleotides for EMSA analysis and DNA precipitations
Biotinylated oligonucleotide precipitation assay
DNA precipitations were carried out as described previously (27). The biotinylated double-stranded wild-type and mutant oligonucleotides were the same as those used for the EMSA assays (Table 1). DNA bound proteins were collected with streptavidin–agarose beads (Pierce Biotechnology, Rockford, IL) and analyzed by western blotting using antibodies specific for GATA-4 and MEF2C (Santa Cruz Biotechnology).
Chromatin immunoprecipitation (ChIP)
We used the ChIP assay kit from Active Motif (Carlsbad, CA). Briefly, sample cell lysates were sonicated with a Sonics Vibra Cell (Newtown, CT), equipped with four 3 mm stepped microtips at 25% power, in this way simultaneously shearing four samples of 350 μl in 1.5 ml Eppendorf tubes. Samples were sonicated 15 times for 15 s with intermittent cooling on ice water for 30 s. The chromatin–antibody protein complex was eluted from the protein-G–Sepharose beads with freshly prepared elution buffer (0.2 M NaHCO3 and 1% SDS). NaCl was added to each ChIP eluate to a final concentration of 200 mM before heating the mixture at 65°C for 4 h to reverse the cross link. The samples were digested with proteinase K for 2 h at 42°C. The DNA from the samples was purified using DNA minicolumns and eluted in water. The GATA-3, GATA-4 and GATA-6 antibodies used were from Santa Cruz Biotechnology.
Quantitative real-time PCR
Primers for PCR analysis for ChIP of the mouse T-catenin promoter were designed using primer Express software (Perkin-Elmer Applied Biosystems). Sequences of primers for the murine proximal T-catenin promoter sequence (–146 to –15) were 5'-TTATTTAGAGCTTCCGAAGA-3' (forward), 5'-GGTGGGTGGTATCTAATCC-3' (reverse); primers for a distal region of the T-catenin promoter (–3373 to –3324), which we designated the distal promoter, were 5'-AATCCTGGGCCATACATGTT-3' (forward) and 5'-TGTGCTGGCATCCATAAGAC-3' (reverse). To detect the amplified promoter regions the SyBr green I reagent mix (Eurogentec, Belgium) was used. The average threshold cycle (Ct) values of triplicate measurements were used for all subsequent calculations on the basis of the delta Ct method.
RT–PCR analysis
RNA from cell lines and tissues was prepared using the RNeasy Mini Kit (Qiagen) according to the manufacturer's instructions. RNA was reverse transcribed to cDNA using pSuperscript Reverse Transcriptase (Invitrogen) and the residual RNA template was removed by RNAse H treatment. Primer sets were developed to specifically amplify T-catenin (human T-catenin: 5'-CTGCCTCTCAATTTGGTACT-3' and 5'-AGGGGTCATCTGTAAATCTC-3'; murine T-catenin: 5'-CCCCTTTCTCTCTTATCCTGAG-3' and 5'-GCTGCCAGCTCTTCCTTTAAA-3'), the different mouse GATA factors (GATA-1: 5'-GCCCAAGAAGCGAATGATTGT-3' and 5'-GCCAGATGCCTTGCGGTTCCT-3'; GATA-2: 5'-TGAAGACATGGAGGCGTTTG-3' and 5'-TCTAGGGCTGCGTCAGGG-3'; GATA-3: 5'-GGGACATCCTGCGCGAACTG-3' and 5'-CTCCAGCGCGTCATGCACC-3') and the GAPDH housekeeping gene (human GAPDH: 5'-GAGGTGAAGGTCGGAGTC-3' and 5'-GAAGATGGTGATGGGATTTC-3'; murine GAPDH: 5'-ACCACAGTCCATGCCATCAC-3' and 5'-TCCACCACCCTGTTGCTGTA-3'). Primers for GATA-4, GATA-5 and GATA-6 were used as described previously (28–30).
Generation of transgenic mice
A construct containing 2917 bp of the human T-catenin promoter, the T-catenin exon 1 sequence and 153 bp of the intron 1 sequence was designed to direct expression of the LacZ reporter gene. To obtain this, the luciferase coding sequence from the pGL3Basic vector was exchanged with the ?-galactosidase coding sequence. The plasmid was digested with SalI to reduce the amount of vector sequences, leaving intact a 6681 bp fragment containing the above-mentioned T-catenin gene fragment and the ?-galactosidase coding sequence. This fragment was purified by agarose gel electrophoresis followed by two consecutive ethanol precipitations, and injected into the pronuclei of fertilized eggs from FVB mice. The manipulated oocytes were transferred into the oviducts of pseudopregnant foster mothers. The transgenic mice were generated in collaboration with Thromb-X nv (Leuven, Belgium).
Transgenics in the progeny were identified by PCR analysis of genomic DNA isolated from tail tissue. The primer set used was specific for the transgene, with one primer located in the T-catenin promoter region (5'-TGTAACTCTGTGACCACCAAGAAG-3') and the other in the LacZ coding sequence (5'-TTGAGGGGACGACGACAGTA-3'). Southern-blot analysis with a ?-galactosidase-specific probe was performed to verify transgene integration and the absence of rearrangements of the transgene.
Analysis of ?-galactosidase expression
The expression of the transgene was analyzed in the F1 progeny, derived from backcrosses with non-transgenic FVB mice. Tissues (and organs) were dissected and fixed for 2 h at 4°C in 0.1 M sodium phosphate (pH 7.3) containing 2% paraformaldehyde, 0.2% glutaraldehyde, 0.01% sodium deoxycholate and 0.02% NP-40. After three 30 min washes in a 0.1 M sodium phosphate buffer (pH 7.3) containing 2 mM MgCl2, 0.01% sodium deoxycholate and 0.02% NP-40, tissues were stained overnight at 30°C in X-gal staining solution . To examine the distribution of transgene expression histologically, organs were dissected, fixed, transferred to 15 and 30% sucrose in the PBS for 4 and 16 h, respectively, and embedded in OCT freezing medium. Sections of 10 μm were cut on a cryostat, washed in 0.1 M sodium phosphate and incubated in X-gal staining solution.
?-Galactosidase enzyme activity was measured using the Galacto-star kit (Tropix Inc.). The tissues were homogenized in lysis buffer (0.1 M potassium phosphate, pH 7.8, containing 0.2% Triton X-100, 1 mM DTT, 0.2 mM phenylmethylsulfonyl fluoride and 5 μg/ml leupeptin). After two freeze–thaw cycles, tissue debris was removed by brief centrifugation. Protein concentrations were measured by the Bio-Rad assay, and equal amounts of tissues were used for the determination of enzymatic activity.
Immunohistochemical analysis
Cryosections of 10 μm were prepared from paraformaldehyde–glutaraldehyde-fixed tissues. The sections were washed in PBS and pre-incubated with 10% goat serum for 10 min. They were then incubated with primary antibody for 45 min at room temperature. The primary antibody used was a polyclonal T-catenin specific antibody (11), diluted in PBS with 1% BSA. The secondary antibody used was a biotin-labeled goat-anti-mouse Ig (Dako, Carpinteria, CA), which was linked to a streptavidin–biotin complex coupled to horseradish peroxidase. Detection was carried out by incubation with the chromogenic peroxidase substrate diaminobenzidine for 5 min (Biogenex, San Remon, CA).
RESULTS
The T-catenin promoter contains putative binding sites for several cardiac- and muscle-specific transcription factors
Sequence analysis of the human T-catenin promoter region and a search in the transcription factor binding site database (Transfac database; www.genomatix.de; www.biobase.de) (31) revealed several putative binding sites for cardiac- and muscle-specific transcription factors. A MEF2C consensus site, several GATA-binding sites, an Nkx2.5 site and a binding site for helix–loop–helix transcription factors eHAND and dHAND were identified. All these regulatory elements were within 300 bp upstream of the transcription initiation site. Parts of the mouse and rat promoter regions were identified by BLAST analysis using the mouse ortholog of the human T-catenin. Sequence alignments showed high sequence conservation only in the proximal T-catenin promoter region. Interestingly, two GATA boxes, the MEF2C site and the HAND binding site, are conserved in the three different species (Figure 1).
Figure 1. Alignment of the 5' flanking sequences of the human, mouse and rat T-catenin genes. The numbering is relative to the transcription initiation site (indicated by arrow). The putative regulatory regions as predicted by the Matinspector Software are indicated by gray boxes.
MEF2C and GATA-4 transactivate the T-catenin promoter synergistically
We used co-transfection studies to assess the contribution of the above-mentioned cis-acting elements and their respective transcription factors to the activation of the T-catenin gene (Figure 2A). Co-transfection into P19 cells of a MEF2C expression plasmid with an T-catenin promoter luciferase construct activated the promoter on average 12-fold (Figure 2B). Mutation of the MEF2C consensus site abolished this activating effect almost completely. Also GATA-4 was able to transactivate the T-catenin promoter after co-transfection into P19 cells, although only about 3- to 4-fold (Figure 2C). Only mutagenesis of the GATA box2 element could destroy this activation effect. Remarkably, the mutation of this GATA box2 site also diminished the effect of MEF2C on the promoter (Figure 2B, 5). Indeed, mutation of both the MEF2C site and the GATA box2 element is needed to completely destroy the activating effect of MEF2C on the T-catenin promoter (Figure 2B, 6).
Figure 2. MEF2C and GATA-4 transactivate the T-catenin promoter in P19 cells. Schematic diagram of the region of the T-catenin gene near the start of transcription showing the wild-type and mutant constructs tested for promoter activity (A). Promoter constructs were co-transfected with MEF2C (B) and GATA-4 (C) transcription factors in P19 embryonal carcinoma cells. Values represent x-fold induction by MEF2C or GATA-4 and are the average of three independent experiments.
Co-transfection of GATA-4 and low amounts of MEF2C with the promoter constructs led to the synergistic activation of the T-catenin promoter, pointing to cooperation between MEF2C and GATA-4 in activating the T-catenin promoter (Figure 3).
Figure 3. MEF2C and GATA-4 transcription factors transactivate the T-catenin promoter synergistically. Different amounts of MEF2C and GATA-4 expression vectors were co-transfected with the T-catenin promoter reporter construct into P19 cells.
Heterodimers of eHAND or dHAND and E47 transcription factors could also transactivate the T-catenin promoter in a co-transfection promoter reporter assay, while the Nkx2.5 transcription factor had no effect on the T-catenin promoter (data not shown).
GATA box2 is required for T-catenin promoter activity in HL-1 cardiac muscle cells
The T-catenin promoter was tested for tissue-specific activity by the transfection of a reporter construct in cell lines of a different origin and its activity was compared with the SV40 promoter activity (Figure 4A). HL-1 is a mouse cardiac-muscle cell line that can be repeatedly passaged in culture without losing its cardiac-specific phenotype (26). Using RT–PCR and western-blot analysis with an T-catenin-specific polyclonal antibody (11), we found that HL-1 cells express T-catenin (Figure 4B), while all other cell lines tested did not express T-catenin. Hence, HL-1 cells should have all the necessary transcription factors to confer expression of the T-catenin gene. Indeed, the activity of the T-catenin promoter in these cells was demonstrated by transient transfection with T-catenin promoter luciferase constructs (Figure 4A). In contrast, the T-catenin promoter showed no activity after transfection in the mouse embryonal carcinoma cell line P19, in the PC3 prostate carcinoma cell line or in the MCF7/AZ epithelial cell line, pointing to a requirement for tissue-specific regulatory factors. To determine whether the MEF2C site and the GATA boxes are actually required for T-catenin promoter activity in HL-1 cells, we mutated the respective core binding sequences (Figure 4C). Mutation of the GATA box2 element reduced the T-catenin promoter activity in HL-1 cells to background levels. Mutation of the MEF2C site, however, had no effect on the T-catenin promoter activity, although HL-1 cells do express the MEF2C transcription factor (data not shown). These results show that the GATA box2 element is absolutely required for the expression of T-catenin in cultured cardiac muscle cells.
Figure 4. (A) Tissue specific activity of the T-catenin promoter in vitro. An T-catenin promoter construct was transfected in MCF7/AZ, PC3, P19 and HL-1 cells and its activity is represented as a fold of the SV40 promoter activity. The light gray bars represent background values measured by the empty pGL3Basic vector. (B) Tissue- and cell-line-specific expression of T-catenin as shown by RT–PCR. (C) Functional role of T-catenin promoter regulatory elements in cardiac muscle specific activity. Wild-type and mutant T-catenin promoter constructs were transfected in cardiac HL-1 cells. Values represent percentage of activity compared to the promoter activity of the wild-type promoter construct (100%). The values are the average of three independent experiments.
The GATA-4 and GATA-6 transcription factors specifically bind the GATA box2 in HL-1 cells
We examined whether the GATA box2 sequence is indeed bound by GATA proteins in cardiac muscle cells. Semiquantitative RT–PCR expression analysis of HL-1 cells for all the six known GATA factors revealed that GATA factor 4 is most prominently expressed, whereas GATA-3, GATA-5 and GATA-6 are weakly to moderately expressed (data not shown). No expression of GATA-1 or GATA-2 was detected. In an EMSA using wild-type and mutant probes encompassing the GATA box2 element (Table 1), the labeled GATA box2 probe formed a sequence-specific DNA–protein complex when incubated with nuclear extracts from HL-1 cells (Figure 5A). The formation of this complex was prevented by addition of a 100-fold excess of the unlabeled probe whereas the competition was far less extensive when using excess cold mutated GATA-box2 oligonucleotide or excess of a probe containing the MEF2C site. To determine if GATA-4 was involved in the shifted complex, we performed a supershift analysis with a GATA-4-specific antibody. As can be seen in Figure 5B, addition of this antibody to the binding reaction resulted in a weak supershifted band. To further confirm binding of GATA-4 to the GATA box2 site, a DNA precipitation assay was performed. A biotinylated wild-type GATA box2 oligonucleotide was able to extract GATA-4 from HL-1 nuclear extracts, which was not possible with the mutated oligonucleotide (Figure 5C). Moreover, ChIP analysis revealed that a proximal T-catenin promoter region containing the GATA-box2 is bound in vivo by transcription factors GATA-4 and GATA-6 but not by GATA-3 (Figure 5D). No binding could be detected to a distal T-catenin promoter region.
Figure 5. Site specificity of transcription factor binding to the GATA box2 site in the T-catenin promoter in cardiac muscle cells. (A) Nuclear extracts from cardiac HL-1 cells were mixed with labeled double-stranded GATA box2 oligonucleotide (Table 1). One complex was shifted, as indicated by an arrow. This complex was removed by the addition of excess cold oligo, but not by an excess mutant oligonucleotide or MEF2C oligonucleotide. (B) Addition of a GATA-4 specific antibody supershifted the formed complex. (C) Wild-type and mutant biotin-labeled GATA box2 oligonucleotides were mixed with HL-1 nuclear extracts, and bound proteins were collected by streptavidin beads and analyzed on western blot with a GATA-4 specific antibody. (D) In vivo binding of proximal and distal T-catenin promoter sequences to transcription factors GATA-3, GATA-4 and GATA-6 in HL-1 cells, as determined by ChIP analysis. Association of transcription factors was quantified by real-time PCR and is depicted as the fold increase in association of factors detected with specific anti-GATA antibodies, compared with the levels detected for negative control IgG antibody.
In vitro binding assays were also performed to test binding of MEF2C to the T-catenin promoter, but here we could not detect any specific shifted complexes (data not shown).
Transgenic mice show tissue-specific activity of the T-catenin promoter
To test tissue specificity of the cloned T-catenin promoter region in vivo, we constructed transgenic mice in which the T-catenin promoter drives the expression of the LacZ reporter gene. Six founder lines were identified by PCR and Southern-blot hybridization. No rearrangements of the transgene were detected. Expression of the transgene was detected initially by soaking dissected tissues in solutions containing the ?-galactosidase substrate X-gal. Blue staining was clearly detected in the brain, with prominent staining in the cerebellum, and testis, and also in discrete areas of the heart and skeletal muscle (Figure 6). In some transgenic lines the transgene was not present in the germline, while other lines did not show any expression of the transgene.
Figure 6. Wholemount X-gal staining of testis, heart, brain and skeletal muscle of wild-type (A) and transgenic (B) mice. Tissues were removed and stained overnight in X-gal solution. Note the variegated expression pattern in heart and skeletal muscle.
Transgenic line #10 showed high expression levels of the transgene and was further analyzed in detail. The tissue specificity of the reporter gene determined by a ?-galactosidase enzyme activity assay is shown in Figure 7. High to moderate levels of enzyme activity were detected only in brain, heart, skeletal muscle and testis. In all other tissues tested, no activity could be detected above the background. This tissue specificity is fully concordant with that found for T-catenin expression by RT–PCR analysis of mouse tissues (11). To test this further, transgene expression and T-catenin expression were analyzed at the cellular level, by X-gal staining and immunohistochemistry on cryosections.
Figure 7. The T-catenin promoter shows tissue-specific activity in vivo. Extracts were prepared from tissues from two transgenic and two wild-type mice and assayed for ?-galactosidase activity. Values for each tissue represent the ?-galactosidase activity in transgenic mice compared to wild-type mice, and are the means of three measurements.
X-gal staining on 10 μm cryosections of testis showed that ?-galactosidase localizes to peritubular myoid cells and to the Sertoli cells (Figure 8B). In the Sertoli cells, the ?-galactosidase staining was seen to be perinuclear, resulting in a dotted staining pattern on the outside of the testis tubule. The small blue deposits seen more centrally in the tubule represent staining in the cytoplasm of the Sertoli cells. Localization of the endogenous T-catenin was visualized by incubation with an T-catenin-specific antibody. In contrast to previous findings where T-catenin was only found localized to the peritubular myoid cells of the human testis (11), in mouse we also found T-catenin expression inside the tubule, apparently on Sertoli-cell–germ-cell contacts (Figure 8C).
Figure 8. Cellular localization of ?-galactosidase and T-catenin expression in testis (A–C) and cerebellum (D–F). Cryosections from wild-type mice were stained with X-gal followed by counterstaining with hematoxylin to reveal tissue details (A and D). Cryosections of transgenic mice were stained with X-gal (B and E) or immunostained with an T-catenin specific antibody (C and F). (B) Blue deposits are found in Sertoli cells (arrows) and peritubular myoid cells (arrowheads). Mo, Molecular layer; Pu, Purkinje cell layer; Gr, Granular cell layer.
In the brain, ?-galactosidase staining mainly localized to the cell bodies of neurons in the granular cell layer of the cerebellum (Figure 8E). In this cerebellum, T-catenin localized in the molecular layer, adjacent to the granular layer, probably in the axons of the neurons whose cell bodies are housed in the granular layer of the cerebellum (Figure 8F).
In heart and skeletal muscle, myocytes expressed ?-galactosidase, but only in a variegated pattern reflecting that the transgene was expressed in some but not all cells.
DISCUSSION
To understand the mechanism of tissue-specific expression of the T-catenin gene in testis, brain, heart and skeletal muscle, we cloned the 5' flanking promoter region of the human gene CTNNA3. Sequence alignments of the human, rat and mouse promoter regions show very high sequence conservation of the first 300 bp upstream of the transcription initiation site. Within these short 5' flanking sequences, a number of conserved regulatory elements were identified, including binding sites for GATA factors, MEF2C and E47/HAND. These binding sites are conserved in both sequence and mutual spacing. We demonstrated that these transcription factors readily transactivate the T-catenin promoter in a specific manner, as mutation of the core binding site interfered with the transactivating potential of the respective transcription factors.
Further, we showed that GATA-4 is directly involved in regulating T-catenin expression in cardiac muscle cells. The GATA-4 transcription factor specifically binds the GATA box2 sequence in the T-catenin promoter as demonstrated by EMSA and DNA precipitations. Furthermore, ChIP analysis showed the in vivo binding of GATA-4 only to a proximal T-catenin promoter region that contains the GATA box2 sequence, while no binding was observed to a distal promoter region. This GATA box2 sequence is critical for cardiac expression of the T-catenin gene, as mutation of this GATA box2 sequence completely abolished T-catenin promoter activity in transfected cardiac HL-1 cells. All these data together show that GATA-4 is an essential factor in regulating T-catenin expression. GATA-4 is a member of the GATA family of zinc finger transcription factors and shows high expression in both the atrium and the ventricle throughout development . This GATA-4 transcription factor is known to regulate the expression of a number of cardiac-specific genes, among them cardiac troponin (32), corin (33), atrial natriuretic factor (34), cardiac sarcolemmal Na+–Ca+ exhanger (35) and B-type natriuretic peptide (36). Interestingly, GATA-4 also emerged as a transcriptional activator of the N-cadherin gene in cardiac muscle cells (37). It thus appears that one of the roles of GATA-4 in cardiac morphogenesis is the establishment of a functional cadherin/catenin complex in cardiomyocytes.
Expression analysis of other GATA factors in HL-1 cells revealed that in addition to GATA-4 there is also expression of GATA-3, GATA-5 and GATA-6 in this cardiac muscle cell line. We assessed in particular the activating role of GATA-4 on the T-catenin promoter as this is the GATA factor expressed most prominently in HL-1 cells. But other GATA factors could have similar effects, as they all seem to recognize the same target sequence. Indeed, we showed in HL-1 cells that in addition to GATA-4, GATA-6 also binds the endogenous T-catenin promoter, whereas GATA-3 does not. GATA-6 is known to colocalize with GATA-4 in the nucleus of cardiac myocytes (32) and hence can together with GATA-4 play a role in activation of transcription from the T-catenin promoter in cardiac muscle cells. Cooperation of GATA-4 and GATA-6 transcription factors in controlling tissue-specific transcription has been reported previously (38).
The MEF2C transcription factor was able to transactivate the T-catenin promoter in co-transfection studies. Involvement of this factor in regulating cardiac expression of T-catenin expression is less clear. Mutation of the MEF2C site had no effect on T-catenin promoter activity in cardiac HL-1 cells. However, it has been described that MEF2C can exert its effect via GATA boxes as well (23). This is also clear from our co-transfection studies in P19 cells: mutation of the MEF2C site only partially abolished MEF2C transactivation and mutation of the GATA box2 site was also necessary to completely destroy the activating effect of MEF2C on the T-catenin promoter. On the other hand, we have been unable to detect MEF2C in a complex bound to the GATA box2 sequence by DNA precipitations or EMSA supershift analysis (data not shown). So far, we have no evidence for a direct role for MEF2C in cardiac expression of T-catenin, but we certainly cannot exclude it.
Another major finding of this study is that the cloned T-catenin promoter region can direct tissue-specific expression of a reporter gene both in vitro, and in vivo in transgenic mice. In the transgenic mice the ?-galactosidase transgene showed expression in heart, testis, brain and skeletal muscle, which exactly concurs with endogenous T-catenin expression. Apparently most information required to direct spatial T-catenin gene activation is comprised within the cloned promoter sequence. Our in vitro studies have shown that GATA-4 is involved in activating the T-catenin promoter in cardiac muscle cells, but this transcription factor is also a key regulator of gonadal gene transcription (24). In testis GATA-4 is expressed in the Sertoli cells (39) and here it transactivates a number of testis-specific genes (24). Our in vivo studies with transgenic mice showed T-catenin promoter activity in Sertoli cells, strongly suggesting that GATA-4 is also involved in regulating testis-specific expression of T-catenin. T-catenin expression in Sertoli cells was also somewhat less restricted than in previous findings, where T-catenin was only detected in the peritubular myoid cells of human testis (11). This difference in localization may be due to species differences between man and mouse. Nonetheless, we successfully used the human T-catenin promoter for mouse transgenesis as the human and mouse promoters show very high sequence identity and conservation of the regulatory elements.
The T-catenin promoter is also highly active in the brain in vivo, which is consistent with endogenous T-catenin protein expression. The factors involved in regulating brain-specific expression of T-catenin could not be studied in vitro, as we found no in vitro system for culturing neurally derived cells expressing T-catenin. Our in vitro data showed that MEF2C can transactivate the T-catenin promoter. Apart from its expression in the heart, the MEF2C transcription factor also shows high expression in postmitotic neurons in different regions of the brain (40), and is therefore a good candidate for regulating brain-specific T-catenin expression. Future experiments will aim at verifying this.
In heart and skeletal muscle we could detect only a variegated expression of the LacZ transgene. Variegated expression is frequently observed in transgenic mice and reflects transgene silencing in a fraction of cells . The mechanism is poorly understood, but it is believed to involve epigenetic modification of the transgene due to chromosomal position effects (42). A remarkable finding is that the position effect variegation in our transgenics was only evident in cardiac and skeletal muscle cells, while in brain and testis transgene expression was apparently not influenced by its chromosomal position. The 2917 bp T-catenin promoter region used for transgenesis may lack an insulator element or locus control region that mitigates variegating position effects in heart and skeletal muscle. Altogether, our studies show that transgene expression driven by the 2917 bp T-catenin promoter sequence occurs exclusively in the heart, testis, brain and skeletal muscle.
In conclusion, we report on the cloning of the promoter region of the T-catenin gene and show that it is responsible for the observed tissue-restricted expression of this -catenin. We demonstrate a functional role for the GATA-4 transcription factor and a possible role for MEF2C in this tissue-restricted expression of T-catenin.
ACKNOWLEDGEMENTS
We thank Dr B.Sekkali, Dr B.Janssens and Dr P.Saunders for helpful discussions. Acknowledgements for pronuclear injections to Dr L.Schoonjans (Thromb-X nv). G.V. was supported by the ‘Vlaams Instituut voor de Bevordering van het Wetenschappelijk-technologisch onderzoek in de Industrie’ and the ‘Centrum voor gezwelziekten’ Gent. G.B. is a postdoctoral Fellow with the Fund for Scientific Research, Flanders (FWO). The research by the authors has been supported by the Association for International Cancer Research (AICR-UK), the Geconcerteerde Onderzoeksacties of Ghent University, FWO-Flanders, Fortis verzekeringen (Belgium) and Interuniversitaire attractiepolen (Belgian Government).
REFERENCES
Shimoyama,Y., Nagafuchi,A., Fujita,S., Gotoh,M., Takeichi,M., Tsukita,S. and Hirohashi,S. ( (1992) ) Cadherin dysfunction in a human cancer cell line: possible involvement of loss of alpha-catenin expression in reduced cell–cell adhesiveness. Cancer Res., , 52, , 5770–5774.
Morton,R.A., Ewing,C.M., Nagafuchi,A., Tsukita,S. and Isaacs,W.B. ( (1993) ) Reduction of E-cadherin levels and deletion of the alpha-catenin gene in human prostate cancer cells. Cancer Res., , 53, , 3585–3590.
Breen,E., Clarke,A., Steele,G. and Mercurio,A.M. ( (1993) ) Poorly differentiated colon carcinoma cell lines deficient in alpha-catenin expression express high levels of surface E-cadherin but lack Ca2+-dependent cell–cell adhesion. Cell Adhes. Commun., , 1, , 239–250.
Roe,S., Koslov,E.R. and Rimm,D.L. ( (1998) ) A mutation in alpha-catenin disrupts adhesion in Clone A cells without perturbing its actin and beta-catenin binding activity. Cell Adhes. Commun., , 5, , 283–296.
Vermeulen,S.J., Nollet,F., Teugels,E., Vennekens,K.M., Malfait,F., Phillippé,J., Speleman,F., Bracke,M.E., van Roy,F.M. and Mareel,M.M. ( (1999) ) The E-catenin gene (CTNNA1) acts as an invasion-suppressor gene in human colon cancer cells. Oncogene, , 18, , 905–916.
Bullions,L.C., Notterman,D.A., Chung,L.S. and Levine,A.J. ( (1997) ) Expression of wild-type alpha-catenin protein in cells with a mutant alpha-catenin gene restores both growth regulation and tumor suppressor activities. Mol. Cell. Biol., , 17, , 4501–4508.
Watabe,M., Nagafuchi,A., Tsukita,S. and Takeichi,M. ( (1994) ) Induction of polarized cell–cell association and retardation of growth by activation of the E-cadherin catenin adhesion system in a dispersed carcinoma line. J. Cell Biol., , 127, , 247–256.
Ewing,C.M., Ru,N., Morton,R.A., Robinson,J.C., Wheelock,M.J., Johnson,K.R., Barrett,J.C. and Isaacs,W.B. ( (1995) ) Chromosome 5 suppresses tumorigenicity of PC3 prostate cancer cells: correlation with re-expression of alpha-catenin and restoration of E-cadherin function. Cancer Res., , 55, , 4813–4817.
van Hengel,J., Gohon,L., Bruyneel,E., Vermeulen,S., Cornelissen,M., Mareel,M. and van Roy,F. ( (1997) ) Protein kinase C activation upregulates intercellular adhesion of -catenin-negative human colon cancer cell variants via induction of desmosomes. J. Cell Biol., , 137, , 1103–1116.
Vasioukhin,V., Bauer,C., Degenstein,L., Wise,B. and Fuchs,E. ( (2001) ) Hyperproliferation and defects in epithelial polarity upon conditional ablation of alpha-catenin in skin. Cell, , 104, , 605–617.
Janssens,B., Goossens,S., Staes,K., Gilbert,B., van Hengel,J., Colpaert,C., Bruyneel,E., Mareel,M. and van Roy,F. ( (2001) ) T-catenin: a novel tissue-specific ?-catenin binding protein mediating strong cell–cell adhesion. J. Cell Sci., , 114, , 3177–3188.
Luo,Y. and Radice,G. ( (2003) ) Cadherin-mediated adhesion is essential for myofibril continuity across the plasma membrane but not for assembly of the contractile apparatus. J. Cell Sci., , 15, , 1471–1479.
Ferreira-Cornwell,M.C., Luo,Y., Narula,N., Lenox,J.M., Lieberman,M. and Radice,G.L. ( (2002) ) Remodeling the intercalated disc leads to cardiomyopathy in mice misexpressing cadherins in the heart. J. Cell Sci., , 115, , 1623–1634.
Bowles,N.E., Bowles,K.R. and Towbin,J.A. ( (2000) ) The ‘final common pathway’ hypothesis and inherited cardiovascular disease. The role of cytoskeletal proteins in dilated cardiomyopathy. Herz, , 25, , 168–175.
Janssens,B., Mohapatra,B., Vatta,M., Goossens,S., Vanpoucke,G., Kools,P., Montoye,T., van Hengel,J., Bowles,N.E., van Roy,F. et al. ( (2003) ) Assessment of the CTNNA3 gene encoding human alpha T-catenin regarding its involvement in dilated cardiomyopathy. Hum. Genet., , 112, , 227–236.
Towbin,J.A. and Bowles,N.E. ( (2002) ) The failing heart. Nature, , 415, , 227–233.
Busby,V., Goossens,S., Nowotny,P., Hamilton,G., Smemo,S., Harold,D., Turic,D., Jehu,L., Myers,A., Womick,M. et al. ( (2004) ) AlphaT-catenin is expressed in human brain and interacts with the Wnt signalling pathway but is not responsible for linkage to chromosome 10 in Alzheimer's disease. Neuromol. Med., , 5, , 133–146.
Ertekin-Taner,N., Ronald,J., Asahara,H., Younkin,L., Hella,M., Jain,S., Gnida,E., Younkin,S., Fadale,D., Ohyagi,Y. et al. ( (2003) ) Fine mapping of the alpha-T catenin gene to a quantitative trait locus on chromosome 10 in late-onset alzheimer's disease pedigrees. Hum. Mol. Genet., , 12, , 3133–3143.
Brand,T. ( (2003) ) Heart development: molecular insights into cardiac specification and early morphogenesis. Dev. Biol., , 258, , 1–19.
Molkentin,J.D. ( (2000) ) The zinc finger-containing transcription factors GATA-4, -5, and -6. Ubiquitously expressed regulators of tissue-specific gene expression. J. Biol. Chem., , 275, , 38949–38952.
Black,B.L. and Olson,E.N. ( (1998) ) Transcriptional control of muscle development by myocyte enhancer factor-2 (MEF2) proteins. Annu. Rev. Cell Dev. Biol., , 14, , 167–196.
Skerjanc,I.S., Petropoulos,H., Ridgeway,A.G. and Wilton,S. ( (1998) ) Myocyte enhancer factor 2C and Nkx2-5 up-regulate each other's expression and initiate cardiomyogenesis in P19 cells. J. Biol. Chem., , 273, , 34904–34910.
Morin,S., Charron,F., Robitaille,L. and Nemer,M. ( (2000) ) GATA-dependent recruitment of MEF2 proteins to target promoters. EMBO J., , 19, , 2046–2055.
Tremblay,J.J. and Viger,R.S. ( (2001) ) GATA factors differentially activate multiple gonadal promoters through conserved GATA regulatory elements. Endocrinology, , 142, , 977–986.
Molkentin,J.D., Kalvakolanu,D.V. and Markham,B.E. ( (1994) ) Transcription factor GATA-4 regulates cardiac muscle-specific expression of the alpha-myosin heavy-chain gene. Mol. Cell. Biol., , 14, , 4947–4957.
Claycomb,W.C., Lanson,N.A.,Jr, Stallworth,B.S., Egeland,D.B., Delcarpio,J.B., Bahinski,A. and Izzo,N.J.,Jr ( (1998) ) HL-1 cells: a cardiac muscle cell line that contracts and retains phenotypic characteristics of the adult cardiomyocyte. Proc. Natl Acad. Sci. USA, , 95, , 2979–2984.
Hata,A., Seoane,J., Lagna,G., Montalvo,E., Hemmati-Brivanlou,A. and Massague,J. ( (2000) ) OAZ uses distinct DNA- and protein-binding zinc fingers in separate BMP-Smad and Olf signaling pathways. Cell, , 100, , 229–240.
Meyer,N., Jaconi,M., Landopoulou,A., Fort,P. and Puceat,M. ( (2000) ) A fluorescent reporter gene as a marker for ventricular specification in ES-derived cardiac cells. FEBS Lett., , 478, , 151–158.
Molkentin,J.D., Tymitz,K.M., Richardson,J.A. and Olson,E.N. ( (2000) ) Abnormalities of the genitourinary tract in female mice lacking GATA5. Mol. Cell. Biol., , 20, , 5256–5260.
Molkentin,J.D., Lin,Q., Duncan,S.A. and Olson,E.N. ( (1997) ) Requirement of the transcription factor GATA4 for heart tube formation and ventral morphogenesis. Genes Dev., , 11, , 1061–1072.
Quandt,K., Frech,K., Karas,H., Wingender,E. and Werner,T. ( (1995) ) MatInd and MatInspector—new fast and versatile tools for detection of consensus matches in nucleotide sequence data. Nucleic Acids Res., , 23, , 4878–4884.
Bhavsar,P.K., Dellow,K.A., Yacoub,M.H., Brand,N.J. and Barton,P.J. ( (2000) ) Identification of cis-acting DNA elements required for expression of the human cardiac troponin I gene promoter. J. Mol. Cell. Cardiol., , 32, , 95–108.
Pan,J., Hinzmann,B., Yan,W., Wu,F., Morser,J. and Wu,Q. ( (2002) ) Genomic structures of the human and murine corin genes and functional GATA elements in their promoters. J. Biol. Chem., , 277, , 38390–38398.
Lee,Y., Shioi,T., Kasahara,H., Jobe,S.M., Wiese,R.J., Markham,B.E. and Izumo,S. ( (1998) ) The cardiac tissue-restricted homeobox protein Csx/Nkx2.5 physically associates with the zinc finger protein GATA4 and cooperatively activates atrial natriuretic factor gene expression. Mol. Cell. Biol., , 18, , 3120–3129.
Nicholas,S.B. and Philipson,K.D. ( (1999) ) Cardiac expression of the Na(+)/Ca(2+) exchanger NCX1 is GATA factor dependent. Am. J. Physiol., , 277, , H324–330.
Charron,F., Paradis,P., Bronchain,O., Nemer,G. and Nemer,M. ( (1999) ) Cooperative interaction between GATA-4 and GATA-6 regulates myocardial gene expression. Mol. Cell. Biol., , 19, , 4355–4365.
Zhang,H., Toyofuku,T., Kamei,J. and Hori,M. ( (2003) ) GATA-4 regulates cardiac morphogenesis through transactivation of the N-cadherin gene. Biochem. Biophys. Res. Commun., , 312, , 1033–1038.
Fluck,C.E. and Miller,W.L. ( (2004) ) GATA-4 and GATA-6 modulate tissue-specific transcription of the human gene for P450c17 by direct interaction with Sp1. Mol. Endocrinol., , 18, , 1144–1157.
Ketola,I., Anttonen,M., Vaskivuo,T., Tapanainen,J.S., Toppari,J. and Heikinheimo,M. ( (2002) ) Developmental expression and spermatogenic stage specificity of transcription factors GATA-1 and GATA-4 and their cofactors FOG-1 and FOG-2 in the mouse testis. Eur. J. Endocrinol., , 147, , 397–406.
Leifer,D., Golden,J. and Kowall,N.W. ( (1994) ) Myocyte-specific enhancer binding factor 2C expression in human brain development. Neuroscience, , 63, , 1067–1079.
Dillon,N. and Festenstein,R. ( (2002) ) Unravelling heterochromatin: competition between positive and negative factors regulates accessibility. Trends Genet., , 18, , 252–258.
Ramirez,A., Milot,E., Ponsa,I., Marcos-Gutierrez,C., Page,A., Santos,M., Jorcano,J. and Vidal,M. ( (2001) ) Sequence and chromosomal context effects on variegated expression of keratin 5/lacZ constructs in stratified epithelia of transgenic mice. Genetics, , 158, , 341–350.(Griet Vanpoucke, Steven Goossens1, Bram )
* To whom correspondence should be addressed. Tel: +32 9 3313740/3313600; Fax: +32 9 3313609; Email: geert.berx@dmbr.Ugent.be
Present address: Griet Vanpoucke, MRC—Human Reproductive Sciences Unit, 49, Little France Crescent, Edinburgh EH16 4SB, UK
DDBJ/EMBL/GenBank accession no. AF361938
ABSTRACT
T-catenin is a recently identified member of the -catenin family of cell–cell adhesion molecules. Its expression is restricted mainly to cardiomyocytes, although it is also expressed in skeletal muscle, testis and brain. Like other -catenins, T-catenin provides an indispensable link between a cadherin-based adhesion complex and the actin cytoskeleton, resulting in strong cell–cell adhesion. We show here that the tissue-specificity of T-catenin expression is controlled by its promoter region. By in silico analysis, we found that the T-catenin promoter contains several binding sites for cardiac and muscle-specific transcription factors. By co-transfection studies in P19 embryonal carcinoma cells, we demonstrated that MEF2C and GATA-4 each have an activating effect on the T-catenin promoter. Transfections with wild-type and mutant promoter constructs in cardiac HL-1 cells indicated that one GATA box is absolutely required for high T-catenin promoter activity in these cells. Furthermore, we showed that the GATA-4 transcription factor specifically binds and activates the T-catenin promoter in vivo in cardiac HL-1 cells. In vivo promoter analysis in transgenic mice revealed that the isolated T-catenin promoter region could direct the tissue-specific expression of a LacZ reporter gene in concordance with endogenous T-catenin expression.
INTRODUCTION
-Catenins are key molecules of the E-cadherin-mediated cell–cell adhesion complex, because they make the indispensable link to the actin cytoskeleton. The importance of this link to confer strong and functional cell–cell adhesion is illustrated by tumor cells that have lost a functional E-catenin protein, a change that is associated with the loss of cell–cell aggregation and the gain of invasive capacity (1–6). Re-introduction of exogenous functional -catenin results in the restoration of cell–cell adhesion and the inhibition of the invasive capacity in vitro and in vivo (3,6–9). Besides its role as an invasion-suppressor molecule, -catenin has a tumor-growth suppressive capacity. This was recently demonstrated by a conditional knock-out of E-catenin in the epidermis (10). Ablation of E-catenin expression in the skin results in a sustained activation of the Ras-MAPK pathway leading to hyperproliferation of the epidermal cells (10).
Recently, a new member of the -catenin family was identified and termed T-catenin. This novel -catenin shows very high sequence homology to both E- and N-catenins (11). T-catenin expression is restricted to certain tissues. It was first discovered in testis, but also found in cardiac and skeletal muscle and in the brain (11). The distinct expression pattern of T-catenin contrasts with the ubiquitous expression of the closely related E-catenin. In some cell types E- and T-catenin are co-expressed, as is found at the intercalated discs of cardiac muscle cells, whereas in other tissues, for instance in human testis, they are differentially localized (11).
Not much is known so far about the function of this recently identified T-catenin. It has been shown that T-catenin can bind strongly to ?-catenin in heart and testis tissues, and in vitro data show that T-catenin can function as a genuine -catenin by providing a link between a cadherin-mediated cell–cell adhesion complex and the actin cytoskeleton (11). N-cadherin-mediated adhesion is critical for proper myofibril organization in cardiomyocytes (12), and altered cadherin expression in the myocardium leads to dilated cardiomyopathy (DCM) (13). DCM is a ‘cytoskeletalopathy’, as most of the DCM genes identified so far have an influence on actin cytoskeleton organization (14). Interestingly, the gene encoding human T-catenin, CTNNA3, was mapped to chromosome 10q21 (15), a region that has been linked to certain cases of dominantly inherited autosomally DCM (16). In view of its high expression in the heart and its presumed role in anchoring the actin cytoskeleton to the cadherin-mediated cell–cell adhesion complex, T-catenin is considered as a good candidate gene for inherited or sporadic DCM. However, so far no final evidence for such causative link has been found (15). Besides their structural role in cell–cell adhesion, -catenins have also been implicated in signaling events. Similar to other -catenins, T-catenin is also able to inhibit Wnt signaling (17). Based on this finding and on the chromosomal position of T-catenin, a link between T-catenin gene variation and late-onset of Alzheimer disease has been proposed (17,18).
The close relation of -catenin genes and their partly overlapping expression patterns makes understanding the transcriptional regulation of these genes of considerable interest. Moreover, due to the particular expression pattern of T-catenin being restricted to a very limited number of tissues, its promoter must be very tightly regulated and is thus an interesting object to study.
Extensive study of the cardiac-specific gene regulatory network has shown that MEF2C, Nkx2.5 and GATA proteins are key transcriptional regulators in the heart (19). GATA factors are Cys4 zinc finger transcription factors with GATA-4, -5 and -6 playing roles in the development and proper function of the heart . Members of the MEF2 family of MADS-box transcription factors are expressed at high levels in all muscle cells, and MEF2C is essential for proper heart development . Overexpressed MEF2C can initiate cardiomyogenesis in embryonic carcinoma cells (22). In cardiomyocytes, MEF2C acts together with GATA factors to induce gene transcription (23). Apart from their role in heart-specific gene regulation, certain GATA factors and MEF2C are also expressed in other tissues exhibiting T-catenin expression. MEF2C is found in skeletal muscle cells, where it strongly potentiates the transcriptional activity of MyoD, and in post-mitotic neurons in the brain, whereas GATA-4 is involved in gene regulation in the testis (24).
In the present study, we analyze the human T-catenin promoter region both in vitro and in vivo and we determine the role of MEF2C and GATA-factors in the tissue-restricted expression of the T-catenin gene.
MATERIALS AND METHODS
Cloning and sequencing of the human T-catenin promoter
A human bacterial artificial chromosome (BAC) library (Genome Systems Inc.) was screened by a PCR specific for the first exon of the CTNNA3 gene (5'-TGTCATCTGCCTCTCAATTTG-3', 5'-ATGCTGCCTTTCTGTTTCTTC-3'). One positive BAC clone was obtained. Fragments containing exon 1 were identified by Southern hybridization with an exon-1-specific primer and subcloned into the pGEM11 vector to generate pGEM11-hTctnprom. Clones of interest were identified by colony hybridization and sequenced. A sequence of 3412 bp of the human T-catenin promoter was deposited in the GenBank database (GenBank accession no. AF361938 ).
Luciferase reporter plasmids and site-directed mutagenesis
A 3266 bp SacI–SpeI fragment from the pGEM11-hTctnprom construct was blunt-ended and cloned into the blunt-ended HindIII site of the pGL3-Basic Luciferase reporter vector (Promega, Madison, WI) to generate plasmid Tctnprom-luc1.
Site-directed mutations of two potential GATA-binding sites and one putative MEF2C-binding site were introduced into the Tctnprom-luc1 plasmid using the QuickChange site-directed mutagenesis kit (Stratagene). The following primers were used to generate the mutated constructs (mutations are in boldface and are underlined): 5'-TAACCTCCCCTTTCTTTCTTAGGCTGGGTGAACAACGCT-3' and 5'-GAGCGTTGTTCACCCAGCCTAAGAAAGAAAGGGGAGGTTA-3' for GATA box1. 5'-ACTAGCGGTTCAGGATTACCTACCACCCACCCTGGCTTG-3' and 5'-CAAGCCACCGTGGGTGGTAGGTAATCCTGAACCGCTAGT-3' for GATA box2. 5'-GTCTGGCCTTTTTCAACGAAGCACGCTCAGTAACAAGTTGTCAG-3' and 5'-CTGACAACTTGTTACTGAGCGTGCTTCGTTGAAAAAGGCCAGAC-3' for the MEF2C-binding site.
Eukaryotic expression vectors
A GATA-4 expression vector (pcDNA1/GATA-4) and an Nkx2.5 expression vector (pGK-Nkx2.5) were obtained from Dr J.Molkentin (25) and Dr I.Skerjanc (22), respectively. The MEF2C coding sequence was PCR amplified from human heart cDNA (BD Biosciences Clontech, Palo Alto, CA) using primers 5'-GGGACTATGGGGAGAAAAAAGA-3' and 5'-TGTTGCCCATCCTTCAGAAAGT-3', and cloned into the pEF6Ahismyc expression vector (Invitrogen, San Diego, CA).
Cell culture
P19 embryonal carcinoma cells were cultured in -minimal essential medium (Invitrogen) containing 5% newborn calf serum, 5% fetal calf serum, 2 mM L-glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin. HL-1 cardiac cells were cultured as described previously (26).
Transfections and reporter gene assays
Co-transfections in P19 cells were performed using FuGENE 6 Reagent (Invitrogen). These co-transfections included 0.5 μg of the wild-type or mutant T-catenin promoter luciferase reporter constructs combined with 0.5 μg of the pcDNA1/GATA-4, pGK-Nkx2.5 or pEF6Ahismyc-MEF2C expression vectors, in various combinations. Total DNA for each transfection was corrected by the addition of empty expression vector. In each transfection, 0.5 μg of a CMV driven LacZ expression vector (pUT651; Eurogentec) was added as a control for the transfection efficiency. HL-1 cells were transfected using calcium phosphate precipitation in the presence of serum. The calcium phosphate precipitate contained 2 μg of the luciferase reporter plasmid and 1 μg of the pUT651 LacZ expression vector as a control for transfection efficiency. Fresh medium was added to the cells 12 h after transfection.
Forty-eight hours after transfection, the cells were lysed and luciferase and ?-galactosidase activities were measured by adding a luciferase substrate buffer and a ?-galactosidase substrate (Galactostar kit, Tropix Inc., Bedford, MA), respectively.
Nuclear extract preparation and EMSA analysis
Nuclear extracts of HL-1 cells were prepared by controlled NP-40 lysis. Briefly, confluent cell monolayers in 75 mm2 dishes were washed twice with ice-cold phosphate-buffered saline (PBS), harvested by scraping in PBS, and centrifuged at 1500 g for 5 min. The pellets were washed once with PBS and pelleted again as before. The cell pellets were suspended in buffer 1 and the cells were allowed to swell on ice for 15 min. The cells were then lysed by adding 0.6% NP-40 in buffer 1 and vortex-mixed briefly. Lysates were centrifuged at full speed for 15 min and the cytoplasmic fraction was discarded. Nuclei pellets were resuspended in buffer 2 (20 mM HEPES, pH 7.5, 1 mM MgCl2, 400 mM NaCl, 10 mM KCl, 20% glycerol, 0.5 mM EDTA, 0.1 mM EGTA, 2 mM Pefabloc, 0.5 mM DTT and 0.15 IU/ml aprotinin) containing 1% NP-40, vortex-mixed well and centrifuged for 15 min at full speed. The supernatant (nuclear extract) was recovered and dialyzed overnight against 1.6x EMSA binding buffer (20 mM HEPES, pH 7.9, 100 mM KCl, 2 mM DTT, 1 mM EDTA and 20% glycerol) with one buffer change.
Oligonucleotide probes used for EMSA analysis are listed in Table 1. Each reaction mixture (20 μl) contained 12 mM HEPES, pH 7.9, 60 mM KCl, 1.2 mM DTT, 0.6 mM EDTA, 12% glycerol, 0.1% NP-40, 0.05% BSA, 1 μg poly(dI–dC), 2–10 μg nuclear extract and 50 fmol 32P end-labeled probe (10 000 c.p.m.). The mixture was incubated at 30°C for 30 min and loaded directly onto a 6% non-denaturing polyacrylamide gel in 0.5x Tris–borate-EDTA buffer. For competition experiments, cold competitors were added to the reaction mixture simultaneously with the labeled probe. For supershift assays, a GATA-4 antibody (Santa Cruz Biotechnology Inc., Santa Cruz, CA) was added to the DNA/nuclear extract mix, and the reactions were allowed to proceed for 20 min before loading the samples.
Table 1. Wild-type and mutant oligonucleotides for EMSA analysis and DNA precipitations
Biotinylated oligonucleotide precipitation assay
DNA precipitations were carried out as described previously (27). The biotinylated double-stranded wild-type and mutant oligonucleotides were the same as those used for the EMSA assays (Table 1). DNA bound proteins were collected with streptavidin–agarose beads (Pierce Biotechnology, Rockford, IL) and analyzed by western blotting using antibodies specific for GATA-4 and MEF2C (Santa Cruz Biotechnology).
Chromatin immunoprecipitation (ChIP)
We used the ChIP assay kit from Active Motif (Carlsbad, CA). Briefly, sample cell lysates were sonicated with a Sonics Vibra Cell (Newtown, CT), equipped with four 3 mm stepped microtips at 25% power, in this way simultaneously shearing four samples of 350 μl in 1.5 ml Eppendorf tubes. Samples were sonicated 15 times for 15 s with intermittent cooling on ice water for 30 s. The chromatin–antibody protein complex was eluted from the protein-G–Sepharose beads with freshly prepared elution buffer (0.2 M NaHCO3 and 1% SDS). NaCl was added to each ChIP eluate to a final concentration of 200 mM before heating the mixture at 65°C for 4 h to reverse the cross link. The samples were digested with proteinase K for 2 h at 42°C. The DNA from the samples was purified using DNA minicolumns and eluted in water. The GATA-3, GATA-4 and GATA-6 antibodies used were from Santa Cruz Biotechnology.
Quantitative real-time PCR
Primers for PCR analysis for ChIP of the mouse T-catenin promoter were designed using primer Express software (Perkin-Elmer Applied Biosystems). Sequences of primers for the murine proximal T-catenin promoter sequence (–146 to –15) were 5'-TTATTTAGAGCTTCCGAAGA-3' (forward), 5'-GGTGGGTGGTATCTAATCC-3' (reverse); primers for a distal region of the T-catenin promoter (–3373 to –3324), which we designated the distal promoter, were 5'-AATCCTGGGCCATACATGTT-3' (forward) and 5'-TGTGCTGGCATCCATAAGAC-3' (reverse). To detect the amplified promoter regions the SyBr green I reagent mix (Eurogentec, Belgium) was used. The average threshold cycle (Ct) values of triplicate measurements were used for all subsequent calculations on the basis of the delta Ct method.
RT–PCR analysis
RNA from cell lines and tissues was prepared using the RNeasy Mini Kit (Qiagen) according to the manufacturer's instructions. RNA was reverse transcribed to cDNA using pSuperscript Reverse Transcriptase (Invitrogen) and the residual RNA template was removed by RNAse H treatment. Primer sets were developed to specifically amplify T-catenin (human T-catenin: 5'-CTGCCTCTCAATTTGGTACT-3' and 5'-AGGGGTCATCTGTAAATCTC-3'; murine T-catenin: 5'-CCCCTTTCTCTCTTATCCTGAG-3' and 5'-GCTGCCAGCTCTTCCTTTAAA-3'), the different mouse GATA factors (GATA-1: 5'-GCCCAAGAAGCGAATGATTGT-3' and 5'-GCCAGATGCCTTGCGGTTCCT-3'; GATA-2: 5'-TGAAGACATGGAGGCGTTTG-3' and 5'-TCTAGGGCTGCGTCAGGG-3'; GATA-3: 5'-GGGACATCCTGCGCGAACTG-3' and 5'-CTCCAGCGCGTCATGCACC-3') and the GAPDH housekeeping gene (human GAPDH: 5'-GAGGTGAAGGTCGGAGTC-3' and 5'-GAAGATGGTGATGGGATTTC-3'; murine GAPDH: 5'-ACCACAGTCCATGCCATCAC-3' and 5'-TCCACCACCCTGTTGCTGTA-3'). Primers for GATA-4, GATA-5 and GATA-6 were used as described previously (28–30).
Generation of transgenic mice
A construct containing 2917 bp of the human T-catenin promoter, the T-catenin exon 1 sequence and 153 bp of the intron 1 sequence was designed to direct expression of the LacZ reporter gene. To obtain this, the luciferase coding sequence from the pGL3Basic vector was exchanged with the ?-galactosidase coding sequence. The plasmid was digested with SalI to reduce the amount of vector sequences, leaving intact a 6681 bp fragment containing the above-mentioned T-catenin gene fragment and the ?-galactosidase coding sequence. This fragment was purified by agarose gel electrophoresis followed by two consecutive ethanol precipitations, and injected into the pronuclei of fertilized eggs from FVB mice. The manipulated oocytes were transferred into the oviducts of pseudopregnant foster mothers. The transgenic mice were generated in collaboration with Thromb-X nv (Leuven, Belgium).
Transgenics in the progeny were identified by PCR analysis of genomic DNA isolated from tail tissue. The primer set used was specific for the transgene, with one primer located in the T-catenin promoter region (5'-TGTAACTCTGTGACCACCAAGAAG-3') and the other in the LacZ coding sequence (5'-TTGAGGGGACGACGACAGTA-3'). Southern-blot analysis with a ?-galactosidase-specific probe was performed to verify transgene integration and the absence of rearrangements of the transgene.
Analysis of ?-galactosidase expression
The expression of the transgene was analyzed in the F1 progeny, derived from backcrosses with non-transgenic FVB mice. Tissues (and organs) were dissected and fixed for 2 h at 4°C in 0.1 M sodium phosphate (pH 7.3) containing 2% paraformaldehyde, 0.2% glutaraldehyde, 0.01% sodium deoxycholate and 0.02% NP-40. After three 30 min washes in a 0.1 M sodium phosphate buffer (pH 7.3) containing 2 mM MgCl2, 0.01% sodium deoxycholate and 0.02% NP-40, tissues were stained overnight at 30°C in X-gal staining solution . To examine the distribution of transgene expression histologically, organs were dissected, fixed, transferred to 15 and 30% sucrose in the PBS for 4 and 16 h, respectively, and embedded in OCT freezing medium. Sections of 10 μm were cut on a cryostat, washed in 0.1 M sodium phosphate and incubated in X-gal staining solution.
?-Galactosidase enzyme activity was measured using the Galacto-star kit (Tropix Inc.). The tissues were homogenized in lysis buffer (0.1 M potassium phosphate, pH 7.8, containing 0.2% Triton X-100, 1 mM DTT, 0.2 mM phenylmethylsulfonyl fluoride and 5 μg/ml leupeptin). After two freeze–thaw cycles, tissue debris was removed by brief centrifugation. Protein concentrations were measured by the Bio-Rad assay, and equal amounts of tissues were used for the determination of enzymatic activity.
Immunohistochemical analysis
Cryosections of 10 μm were prepared from paraformaldehyde–glutaraldehyde-fixed tissues. The sections were washed in PBS and pre-incubated with 10% goat serum for 10 min. They were then incubated with primary antibody for 45 min at room temperature. The primary antibody used was a polyclonal T-catenin specific antibody (11), diluted in PBS with 1% BSA. The secondary antibody used was a biotin-labeled goat-anti-mouse Ig (Dako, Carpinteria, CA), which was linked to a streptavidin–biotin complex coupled to horseradish peroxidase. Detection was carried out by incubation with the chromogenic peroxidase substrate diaminobenzidine for 5 min (Biogenex, San Remon, CA).
RESULTS
The T-catenin promoter contains putative binding sites for several cardiac- and muscle-specific transcription factors
Sequence analysis of the human T-catenin promoter region and a search in the transcription factor binding site database (Transfac database; www.genomatix.de; www.biobase.de) (31) revealed several putative binding sites for cardiac- and muscle-specific transcription factors. A MEF2C consensus site, several GATA-binding sites, an Nkx2.5 site and a binding site for helix–loop–helix transcription factors eHAND and dHAND were identified. All these regulatory elements were within 300 bp upstream of the transcription initiation site. Parts of the mouse and rat promoter regions were identified by BLAST analysis using the mouse ortholog of the human T-catenin. Sequence alignments showed high sequence conservation only in the proximal T-catenin promoter region. Interestingly, two GATA boxes, the MEF2C site and the HAND binding site, are conserved in the three different species (Figure 1).
Figure 1. Alignment of the 5' flanking sequences of the human, mouse and rat T-catenin genes. The numbering is relative to the transcription initiation site (indicated by arrow). The putative regulatory regions as predicted by the Matinspector Software are indicated by gray boxes.
MEF2C and GATA-4 transactivate the T-catenin promoter synergistically
We used co-transfection studies to assess the contribution of the above-mentioned cis-acting elements and their respective transcription factors to the activation of the T-catenin gene (Figure 2A). Co-transfection into P19 cells of a MEF2C expression plasmid with an T-catenin promoter luciferase construct activated the promoter on average 12-fold (Figure 2B). Mutation of the MEF2C consensus site abolished this activating effect almost completely. Also GATA-4 was able to transactivate the T-catenin promoter after co-transfection into P19 cells, although only about 3- to 4-fold (Figure 2C). Only mutagenesis of the GATA box2 element could destroy this activation effect. Remarkably, the mutation of this GATA box2 site also diminished the effect of MEF2C on the promoter (Figure 2B, 5). Indeed, mutation of both the MEF2C site and the GATA box2 element is needed to completely destroy the activating effect of MEF2C on the T-catenin promoter (Figure 2B, 6).
Figure 2. MEF2C and GATA-4 transactivate the T-catenin promoter in P19 cells. Schematic diagram of the region of the T-catenin gene near the start of transcription showing the wild-type and mutant constructs tested for promoter activity (A). Promoter constructs were co-transfected with MEF2C (B) and GATA-4 (C) transcription factors in P19 embryonal carcinoma cells. Values represent x-fold induction by MEF2C or GATA-4 and are the average of three independent experiments.
Co-transfection of GATA-4 and low amounts of MEF2C with the promoter constructs led to the synergistic activation of the T-catenin promoter, pointing to cooperation between MEF2C and GATA-4 in activating the T-catenin promoter (Figure 3).
Figure 3. MEF2C and GATA-4 transcription factors transactivate the T-catenin promoter synergistically. Different amounts of MEF2C and GATA-4 expression vectors were co-transfected with the T-catenin promoter reporter construct into P19 cells.
Heterodimers of eHAND or dHAND and E47 transcription factors could also transactivate the T-catenin promoter in a co-transfection promoter reporter assay, while the Nkx2.5 transcription factor had no effect on the T-catenin promoter (data not shown).
GATA box2 is required for T-catenin promoter activity in HL-1 cardiac muscle cells
The T-catenin promoter was tested for tissue-specific activity by the transfection of a reporter construct in cell lines of a different origin and its activity was compared with the SV40 promoter activity (Figure 4A). HL-1 is a mouse cardiac-muscle cell line that can be repeatedly passaged in culture without losing its cardiac-specific phenotype (26). Using RT–PCR and western-blot analysis with an T-catenin-specific polyclonal antibody (11), we found that HL-1 cells express T-catenin (Figure 4B), while all other cell lines tested did not express T-catenin. Hence, HL-1 cells should have all the necessary transcription factors to confer expression of the T-catenin gene. Indeed, the activity of the T-catenin promoter in these cells was demonstrated by transient transfection with T-catenin promoter luciferase constructs (Figure 4A). In contrast, the T-catenin promoter showed no activity after transfection in the mouse embryonal carcinoma cell line P19, in the PC3 prostate carcinoma cell line or in the MCF7/AZ epithelial cell line, pointing to a requirement for tissue-specific regulatory factors. To determine whether the MEF2C site and the GATA boxes are actually required for T-catenin promoter activity in HL-1 cells, we mutated the respective core binding sequences (Figure 4C). Mutation of the GATA box2 element reduced the T-catenin promoter activity in HL-1 cells to background levels. Mutation of the MEF2C site, however, had no effect on the T-catenin promoter activity, although HL-1 cells do express the MEF2C transcription factor (data not shown). These results show that the GATA box2 element is absolutely required for the expression of T-catenin in cultured cardiac muscle cells.
Figure 4. (A) Tissue specific activity of the T-catenin promoter in vitro. An T-catenin promoter construct was transfected in MCF7/AZ, PC3, P19 and HL-1 cells and its activity is represented as a fold of the SV40 promoter activity. The light gray bars represent background values measured by the empty pGL3Basic vector. (B) Tissue- and cell-line-specific expression of T-catenin as shown by RT–PCR. (C) Functional role of T-catenin promoter regulatory elements in cardiac muscle specific activity. Wild-type and mutant T-catenin promoter constructs were transfected in cardiac HL-1 cells. Values represent percentage of activity compared to the promoter activity of the wild-type promoter construct (100%). The values are the average of three independent experiments.
The GATA-4 and GATA-6 transcription factors specifically bind the GATA box2 in HL-1 cells
We examined whether the GATA box2 sequence is indeed bound by GATA proteins in cardiac muscle cells. Semiquantitative RT–PCR expression analysis of HL-1 cells for all the six known GATA factors revealed that GATA factor 4 is most prominently expressed, whereas GATA-3, GATA-5 and GATA-6 are weakly to moderately expressed (data not shown). No expression of GATA-1 or GATA-2 was detected. In an EMSA using wild-type and mutant probes encompassing the GATA box2 element (Table 1), the labeled GATA box2 probe formed a sequence-specific DNA–protein complex when incubated with nuclear extracts from HL-1 cells (Figure 5A). The formation of this complex was prevented by addition of a 100-fold excess of the unlabeled probe whereas the competition was far less extensive when using excess cold mutated GATA-box2 oligonucleotide or excess of a probe containing the MEF2C site. To determine if GATA-4 was involved in the shifted complex, we performed a supershift analysis with a GATA-4-specific antibody. As can be seen in Figure 5B, addition of this antibody to the binding reaction resulted in a weak supershifted band. To further confirm binding of GATA-4 to the GATA box2 site, a DNA precipitation assay was performed. A biotinylated wild-type GATA box2 oligonucleotide was able to extract GATA-4 from HL-1 nuclear extracts, which was not possible with the mutated oligonucleotide (Figure 5C). Moreover, ChIP analysis revealed that a proximal T-catenin promoter region containing the GATA-box2 is bound in vivo by transcription factors GATA-4 and GATA-6 but not by GATA-3 (Figure 5D). No binding could be detected to a distal T-catenin promoter region.
Figure 5. Site specificity of transcription factor binding to the GATA box2 site in the T-catenin promoter in cardiac muscle cells. (A) Nuclear extracts from cardiac HL-1 cells were mixed with labeled double-stranded GATA box2 oligonucleotide (Table 1). One complex was shifted, as indicated by an arrow. This complex was removed by the addition of excess cold oligo, but not by an excess mutant oligonucleotide or MEF2C oligonucleotide. (B) Addition of a GATA-4 specific antibody supershifted the formed complex. (C) Wild-type and mutant biotin-labeled GATA box2 oligonucleotides were mixed with HL-1 nuclear extracts, and bound proteins were collected by streptavidin beads and analyzed on western blot with a GATA-4 specific antibody. (D) In vivo binding of proximal and distal T-catenin promoter sequences to transcription factors GATA-3, GATA-4 and GATA-6 in HL-1 cells, as determined by ChIP analysis. Association of transcription factors was quantified by real-time PCR and is depicted as the fold increase in association of factors detected with specific anti-GATA antibodies, compared with the levels detected for negative control IgG antibody.
In vitro binding assays were also performed to test binding of MEF2C to the T-catenin promoter, but here we could not detect any specific shifted complexes (data not shown).
Transgenic mice show tissue-specific activity of the T-catenin promoter
To test tissue specificity of the cloned T-catenin promoter region in vivo, we constructed transgenic mice in which the T-catenin promoter drives the expression of the LacZ reporter gene. Six founder lines were identified by PCR and Southern-blot hybridization. No rearrangements of the transgene were detected. Expression of the transgene was detected initially by soaking dissected tissues in solutions containing the ?-galactosidase substrate X-gal. Blue staining was clearly detected in the brain, with prominent staining in the cerebellum, and testis, and also in discrete areas of the heart and skeletal muscle (Figure 6). In some transgenic lines the transgene was not present in the germline, while other lines did not show any expression of the transgene.
Figure 6. Wholemount X-gal staining of testis, heart, brain and skeletal muscle of wild-type (A) and transgenic (B) mice. Tissues were removed and stained overnight in X-gal solution. Note the variegated expression pattern in heart and skeletal muscle.
Transgenic line #10 showed high expression levels of the transgene and was further analyzed in detail. The tissue specificity of the reporter gene determined by a ?-galactosidase enzyme activity assay is shown in Figure 7. High to moderate levels of enzyme activity were detected only in brain, heart, skeletal muscle and testis. In all other tissues tested, no activity could be detected above the background. This tissue specificity is fully concordant with that found for T-catenin expression by RT–PCR analysis of mouse tissues (11). To test this further, transgene expression and T-catenin expression were analyzed at the cellular level, by X-gal staining and immunohistochemistry on cryosections.
Figure 7. The T-catenin promoter shows tissue-specific activity in vivo. Extracts were prepared from tissues from two transgenic and two wild-type mice and assayed for ?-galactosidase activity. Values for each tissue represent the ?-galactosidase activity in transgenic mice compared to wild-type mice, and are the means of three measurements.
X-gal staining on 10 μm cryosections of testis showed that ?-galactosidase localizes to peritubular myoid cells and to the Sertoli cells (Figure 8B). In the Sertoli cells, the ?-galactosidase staining was seen to be perinuclear, resulting in a dotted staining pattern on the outside of the testis tubule. The small blue deposits seen more centrally in the tubule represent staining in the cytoplasm of the Sertoli cells. Localization of the endogenous T-catenin was visualized by incubation with an T-catenin-specific antibody. In contrast to previous findings where T-catenin was only found localized to the peritubular myoid cells of the human testis (11), in mouse we also found T-catenin expression inside the tubule, apparently on Sertoli-cell–germ-cell contacts (Figure 8C).
Figure 8. Cellular localization of ?-galactosidase and T-catenin expression in testis (A–C) and cerebellum (D–F). Cryosections from wild-type mice were stained with X-gal followed by counterstaining with hematoxylin to reveal tissue details (A and D). Cryosections of transgenic mice were stained with X-gal (B and E) or immunostained with an T-catenin specific antibody (C and F). (B) Blue deposits are found in Sertoli cells (arrows) and peritubular myoid cells (arrowheads). Mo, Molecular layer; Pu, Purkinje cell layer; Gr, Granular cell layer.
In the brain, ?-galactosidase staining mainly localized to the cell bodies of neurons in the granular cell layer of the cerebellum (Figure 8E). In this cerebellum, T-catenin localized in the molecular layer, adjacent to the granular layer, probably in the axons of the neurons whose cell bodies are housed in the granular layer of the cerebellum (Figure 8F).
In heart and skeletal muscle, myocytes expressed ?-galactosidase, but only in a variegated pattern reflecting that the transgene was expressed in some but not all cells.
DISCUSSION
To understand the mechanism of tissue-specific expression of the T-catenin gene in testis, brain, heart and skeletal muscle, we cloned the 5' flanking promoter region of the human gene CTNNA3. Sequence alignments of the human, rat and mouse promoter regions show very high sequence conservation of the first 300 bp upstream of the transcription initiation site. Within these short 5' flanking sequences, a number of conserved regulatory elements were identified, including binding sites for GATA factors, MEF2C and E47/HAND. These binding sites are conserved in both sequence and mutual spacing. We demonstrated that these transcription factors readily transactivate the T-catenin promoter in a specific manner, as mutation of the core binding site interfered with the transactivating potential of the respective transcription factors.
Further, we showed that GATA-4 is directly involved in regulating T-catenin expression in cardiac muscle cells. The GATA-4 transcription factor specifically binds the GATA box2 sequence in the T-catenin promoter as demonstrated by EMSA and DNA precipitations. Furthermore, ChIP analysis showed the in vivo binding of GATA-4 only to a proximal T-catenin promoter region that contains the GATA box2 sequence, while no binding was observed to a distal promoter region. This GATA box2 sequence is critical for cardiac expression of the T-catenin gene, as mutation of this GATA box2 sequence completely abolished T-catenin promoter activity in transfected cardiac HL-1 cells. All these data together show that GATA-4 is an essential factor in regulating T-catenin expression. GATA-4 is a member of the GATA family of zinc finger transcription factors and shows high expression in both the atrium and the ventricle throughout development . This GATA-4 transcription factor is known to regulate the expression of a number of cardiac-specific genes, among them cardiac troponin (32), corin (33), atrial natriuretic factor (34), cardiac sarcolemmal Na+–Ca+ exhanger (35) and B-type natriuretic peptide (36). Interestingly, GATA-4 also emerged as a transcriptional activator of the N-cadherin gene in cardiac muscle cells (37). It thus appears that one of the roles of GATA-4 in cardiac morphogenesis is the establishment of a functional cadherin/catenin complex in cardiomyocytes.
Expression analysis of other GATA factors in HL-1 cells revealed that in addition to GATA-4 there is also expression of GATA-3, GATA-5 and GATA-6 in this cardiac muscle cell line. We assessed in particular the activating role of GATA-4 on the T-catenin promoter as this is the GATA factor expressed most prominently in HL-1 cells. But other GATA factors could have similar effects, as they all seem to recognize the same target sequence. Indeed, we showed in HL-1 cells that in addition to GATA-4, GATA-6 also binds the endogenous T-catenin promoter, whereas GATA-3 does not. GATA-6 is known to colocalize with GATA-4 in the nucleus of cardiac myocytes (32) and hence can together with GATA-4 play a role in activation of transcription from the T-catenin promoter in cardiac muscle cells. Cooperation of GATA-4 and GATA-6 transcription factors in controlling tissue-specific transcription has been reported previously (38).
The MEF2C transcription factor was able to transactivate the T-catenin promoter in co-transfection studies. Involvement of this factor in regulating cardiac expression of T-catenin expression is less clear. Mutation of the MEF2C site had no effect on T-catenin promoter activity in cardiac HL-1 cells. However, it has been described that MEF2C can exert its effect via GATA boxes as well (23). This is also clear from our co-transfection studies in P19 cells: mutation of the MEF2C site only partially abolished MEF2C transactivation and mutation of the GATA box2 site was also necessary to completely destroy the activating effect of MEF2C on the T-catenin promoter. On the other hand, we have been unable to detect MEF2C in a complex bound to the GATA box2 sequence by DNA precipitations or EMSA supershift analysis (data not shown). So far, we have no evidence for a direct role for MEF2C in cardiac expression of T-catenin, but we certainly cannot exclude it.
Another major finding of this study is that the cloned T-catenin promoter region can direct tissue-specific expression of a reporter gene both in vitro, and in vivo in transgenic mice. In the transgenic mice the ?-galactosidase transgene showed expression in heart, testis, brain and skeletal muscle, which exactly concurs with endogenous T-catenin expression. Apparently most information required to direct spatial T-catenin gene activation is comprised within the cloned promoter sequence. Our in vitro studies have shown that GATA-4 is involved in activating the T-catenin promoter in cardiac muscle cells, but this transcription factor is also a key regulator of gonadal gene transcription (24). In testis GATA-4 is expressed in the Sertoli cells (39) and here it transactivates a number of testis-specific genes (24). Our in vivo studies with transgenic mice showed T-catenin promoter activity in Sertoli cells, strongly suggesting that GATA-4 is also involved in regulating testis-specific expression of T-catenin. T-catenin expression in Sertoli cells was also somewhat less restricted than in previous findings, where T-catenin was only detected in the peritubular myoid cells of human testis (11). This difference in localization may be due to species differences between man and mouse. Nonetheless, we successfully used the human T-catenin promoter for mouse transgenesis as the human and mouse promoters show very high sequence identity and conservation of the regulatory elements.
The T-catenin promoter is also highly active in the brain in vivo, which is consistent with endogenous T-catenin protein expression. The factors involved in regulating brain-specific expression of T-catenin could not be studied in vitro, as we found no in vitro system for culturing neurally derived cells expressing T-catenin. Our in vitro data showed that MEF2C can transactivate the T-catenin promoter. Apart from its expression in the heart, the MEF2C transcription factor also shows high expression in postmitotic neurons in different regions of the brain (40), and is therefore a good candidate for regulating brain-specific T-catenin expression. Future experiments will aim at verifying this.
In heart and skeletal muscle we could detect only a variegated expression of the LacZ transgene. Variegated expression is frequently observed in transgenic mice and reflects transgene silencing in a fraction of cells . The mechanism is poorly understood, but it is believed to involve epigenetic modification of the transgene due to chromosomal position effects (42). A remarkable finding is that the position effect variegation in our transgenics was only evident in cardiac and skeletal muscle cells, while in brain and testis transgene expression was apparently not influenced by its chromosomal position. The 2917 bp T-catenin promoter region used for transgenesis may lack an insulator element or locus control region that mitigates variegating position effects in heart and skeletal muscle. Altogether, our studies show that transgene expression driven by the 2917 bp T-catenin promoter sequence occurs exclusively in the heart, testis, brain and skeletal muscle.
In conclusion, we report on the cloning of the promoter region of the T-catenin gene and show that it is responsible for the observed tissue-restricted expression of this -catenin. We demonstrate a functional role for the GATA-4 transcription factor and a possible role for MEF2C in this tissue-restricted expression of T-catenin.
ACKNOWLEDGEMENTS
We thank Dr B.Sekkali, Dr B.Janssens and Dr P.Saunders for helpful discussions. Acknowledgements for pronuclear injections to Dr L.Schoonjans (Thromb-X nv). G.V. was supported by the ‘Vlaams Instituut voor de Bevordering van het Wetenschappelijk-technologisch onderzoek in de Industrie’ and the ‘Centrum voor gezwelziekten’ Gent. G.B. is a postdoctoral Fellow with the Fund for Scientific Research, Flanders (FWO). The research by the authors has been supported by the Association for International Cancer Research (AICR-UK), the Geconcerteerde Onderzoeksacties of Ghent University, FWO-Flanders, Fortis verzekeringen (Belgium) and Interuniversitaire attractiepolen (Belgian Government).
REFERENCES
Shimoyama,Y., Nagafuchi,A., Fujita,S., Gotoh,M., Takeichi,M., Tsukita,S. and Hirohashi,S. ( (1992) ) Cadherin dysfunction in a human cancer cell line: possible involvement of loss of alpha-catenin expression in reduced cell–cell adhesiveness. Cancer Res., , 52, , 5770–5774.
Morton,R.A., Ewing,C.M., Nagafuchi,A., Tsukita,S. and Isaacs,W.B. ( (1993) ) Reduction of E-cadherin levels and deletion of the alpha-catenin gene in human prostate cancer cells. Cancer Res., , 53, , 3585–3590.
Breen,E., Clarke,A., Steele,G. and Mercurio,A.M. ( (1993) ) Poorly differentiated colon carcinoma cell lines deficient in alpha-catenin expression express high levels of surface E-cadherin but lack Ca2+-dependent cell–cell adhesion. Cell Adhes. Commun., , 1, , 239–250.
Roe,S., Koslov,E.R. and Rimm,D.L. ( (1998) ) A mutation in alpha-catenin disrupts adhesion in Clone A cells without perturbing its actin and beta-catenin binding activity. Cell Adhes. Commun., , 5, , 283–296.
Vermeulen,S.J., Nollet,F., Teugels,E., Vennekens,K.M., Malfait,F., Phillippé,J., Speleman,F., Bracke,M.E., van Roy,F.M. and Mareel,M.M. ( (1999) ) The E-catenin gene (CTNNA1) acts as an invasion-suppressor gene in human colon cancer cells. Oncogene, , 18, , 905–916.
Bullions,L.C., Notterman,D.A., Chung,L.S. and Levine,A.J. ( (1997) ) Expression of wild-type alpha-catenin protein in cells with a mutant alpha-catenin gene restores both growth regulation and tumor suppressor activities. Mol. Cell. Biol., , 17, , 4501–4508.
Watabe,M., Nagafuchi,A., Tsukita,S. and Takeichi,M. ( (1994) ) Induction of polarized cell–cell association and retardation of growth by activation of the E-cadherin catenin adhesion system in a dispersed carcinoma line. J. Cell Biol., , 127, , 247–256.
Ewing,C.M., Ru,N., Morton,R.A., Robinson,J.C., Wheelock,M.J., Johnson,K.R., Barrett,J.C. and Isaacs,W.B. ( (1995) ) Chromosome 5 suppresses tumorigenicity of PC3 prostate cancer cells: correlation with re-expression of alpha-catenin and restoration of E-cadherin function. Cancer Res., , 55, , 4813–4817.
van Hengel,J., Gohon,L., Bruyneel,E., Vermeulen,S., Cornelissen,M., Mareel,M. and van Roy,F. ( (1997) ) Protein kinase C activation upregulates intercellular adhesion of -catenin-negative human colon cancer cell variants via induction of desmosomes. J. Cell Biol., , 137, , 1103–1116.
Vasioukhin,V., Bauer,C., Degenstein,L., Wise,B. and Fuchs,E. ( (2001) ) Hyperproliferation and defects in epithelial polarity upon conditional ablation of alpha-catenin in skin. Cell, , 104, , 605–617.
Janssens,B., Goossens,S., Staes,K., Gilbert,B., van Hengel,J., Colpaert,C., Bruyneel,E., Mareel,M. and van Roy,F. ( (2001) ) T-catenin: a novel tissue-specific ?-catenin binding protein mediating strong cell–cell adhesion. J. Cell Sci., , 114, , 3177–3188.
Luo,Y. and Radice,G. ( (2003) ) Cadherin-mediated adhesion is essential for myofibril continuity across the plasma membrane but not for assembly of the contractile apparatus. J. Cell Sci., , 15, , 1471–1479.
Ferreira-Cornwell,M.C., Luo,Y., Narula,N., Lenox,J.M., Lieberman,M. and Radice,G.L. ( (2002) ) Remodeling the intercalated disc leads to cardiomyopathy in mice misexpressing cadherins in the heart. J. Cell Sci., , 115, , 1623–1634.
Bowles,N.E., Bowles,K.R. and Towbin,J.A. ( (2000) ) The ‘final common pathway’ hypothesis and inherited cardiovascular disease. The role of cytoskeletal proteins in dilated cardiomyopathy. Herz, , 25, , 168–175.
Janssens,B., Mohapatra,B., Vatta,M., Goossens,S., Vanpoucke,G., Kools,P., Montoye,T., van Hengel,J., Bowles,N.E., van Roy,F. et al. ( (2003) ) Assessment of the CTNNA3 gene encoding human alpha T-catenin regarding its involvement in dilated cardiomyopathy. Hum. Genet., , 112, , 227–236.
Towbin,J.A. and Bowles,N.E. ( (2002) ) The failing heart. Nature, , 415, , 227–233.
Busby,V., Goossens,S., Nowotny,P., Hamilton,G., Smemo,S., Harold,D., Turic,D., Jehu,L., Myers,A., Womick,M. et al. ( (2004) ) AlphaT-catenin is expressed in human brain and interacts with the Wnt signalling pathway but is not responsible for linkage to chromosome 10 in Alzheimer's disease. Neuromol. Med., , 5, , 133–146.
Ertekin-Taner,N., Ronald,J., Asahara,H., Younkin,L., Hella,M., Jain,S., Gnida,E., Younkin,S., Fadale,D., Ohyagi,Y. et al. ( (2003) ) Fine mapping of the alpha-T catenin gene to a quantitative trait locus on chromosome 10 in late-onset alzheimer's disease pedigrees. Hum. Mol. Genet., , 12, , 3133–3143.
Brand,T. ( (2003) ) Heart development: molecular insights into cardiac specification and early morphogenesis. Dev. Biol., , 258, , 1–19.
Molkentin,J.D. ( (2000) ) The zinc finger-containing transcription factors GATA-4, -5, and -6. Ubiquitously expressed regulators of tissue-specific gene expression. J. Biol. Chem., , 275, , 38949–38952.
Black,B.L. and Olson,E.N. ( (1998) ) Transcriptional control of muscle development by myocyte enhancer factor-2 (MEF2) proteins. Annu. Rev. Cell Dev. Biol., , 14, , 167–196.
Skerjanc,I.S., Petropoulos,H., Ridgeway,A.G. and Wilton,S. ( (1998) ) Myocyte enhancer factor 2C and Nkx2-5 up-regulate each other's expression and initiate cardiomyogenesis in P19 cells. J. Biol. Chem., , 273, , 34904–34910.
Morin,S., Charron,F., Robitaille,L. and Nemer,M. ( (2000) ) GATA-dependent recruitment of MEF2 proteins to target promoters. EMBO J., , 19, , 2046–2055.
Tremblay,J.J. and Viger,R.S. ( (2001) ) GATA factors differentially activate multiple gonadal promoters through conserved GATA regulatory elements. Endocrinology, , 142, , 977–986.
Molkentin,J.D., Kalvakolanu,D.V. and Markham,B.E. ( (1994) ) Transcription factor GATA-4 regulates cardiac muscle-specific expression of the alpha-myosin heavy-chain gene. Mol. Cell. Biol., , 14, , 4947–4957.
Claycomb,W.C., Lanson,N.A.,Jr, Stallworth,B.S., Egeland,D.B., Delcarpio,J.B., Bahinski,A. and Izzo,N.J.,Jr ( (1998) ) HL-1 cells: a cardiac muscle cell line that contracts and retains phenotypic characteristics of the adult cardiomyocyte. Proc. Natl Acad. Sci. USA, , 95, , 2979–2984.
Hata,A., Seoane,J., Lagna,G., Montalvo,E., Hemmati-Brivanlou,A. and Massague,J. ( (2000) ) OAZ uses distinct DNA- and protein-binding zinc fingers in separate BMP-Smad and Olf signaling pathways. Cell, , 100, , 229–240.
Meyer,N., Jaconi,M., Landopoulou,A., Fort,P. and Puceat,M. ( (2000) ) A fluorescent reporter gene as a marker for ventricular specification in ES-derived cardiac cells. FEBS Lett., , 478, , 151–158.
Molkentin,J.D., Tymitz,K.M., Richardson,J.A. and Olson,E.N. ( (2000) ) Abnormalities of the genitourinary tract in female mice lacking GATA5. Mol. Cell. Biol., , 20, , 5256–5260.
Molkentin,J.D., Lin,Q., Duncan,S.A. and Olson,E.N. ( (1997) ) Requirement of the transcription factor GATA4 for heart tube formation and ventral morphogenesis. Genes Dev., , 11, , 1061–1072.
Quandt,K., Frech,K., Karas,H., Wingender,E. and Werner,T. ( (1995) ) MatInd and MatInspector—new fast and versatile tools for detection of consensus matches in nucleotide sequence data. Nucleic Acids Res., , 23, , 4878–4884.
Bhavsar,P.K., Dellow,K.A., Yacoub,M.H., Brand,N.J. and Barton,P.J. ( (2000) ) Identification of cis-acting DNA elements required for expression of the human cardiac troponin I gene promoter. J. Mol. Cell. Cardiol., , 32, , 95–108.
Pan,J., Hinzmann,B., Yan,W., Wu,F., Morser,J. and Wu,Q. ( (2002) ) Genomic structures of the human and murine corin genes and functional GATA elements in their promoters. J. Biol. Chem., , 277, , 38390–38398.
Lee,Y., Shioi,T., Kasahara,H., Jobe,S.M., Wiese,R.J., Markham,B.E. and Izumo,S. ( (1998) ) The cardiac tissue-restricted homeobox protein Csx/Nkx2.5 physically associates with the zinc finger protein GATA4 and cooperatively activates atrial natriuretic factor gene expression. Mol. Cell. Biol., , 18, , 3120–3129.
Nicholas,S.B. and Philipson,K.D. ( (1999) ) Cardiac expression of the Na(+)/Ca(2+) exchanger NCX1 is GATA factor dependent. Am. J. Physiol., , 277, , H324–330.
Charron,F., Paradis,P., Bronchain,O., Nemer,G. and Nemer,M. ( (1999) ) Cooperative interaction between GATA-4 and GATA-6 regulates myocardial gene expression. Mol. Cell. Biol., , 19, , 4355–4365.
Zhang,H., Toyofuku,T., Kamei,J. and Hori,M. ( (2003) ) GATA-4 regulates cardiac morphogenesis through transactivation of the N-cadherin gene. Biochem. Biophys. Res. Commun., , 312, , 1033–1038.
Fluck,C.E. and Miller,W.L. ( (2004) ) GATA-4 and GATA-6 modulate tissue-specific transcription of the human gene for P450c17 by direct interaction with Sp1. Mol. Endocrinol., , 18, , 1144–1157.
Ketola,I., Anttonen,M., Vaskivuo,T., Tapanainen,J.S., Toppari,J. and Heikinheimo,M. ( (2002) ) Developmental expression and spermatogenic stage specificity of transcription factors GATA-1 and GATA-4 and their cofactors FOG-1 and FOG-2 in the mouse testis. Eur. J. Endocrinol., , 147, , 397–406.
Leifer,D., Golden,J. and Kowall,N.W. ( (1994) ) Myocyte-specific enhancer binding factor 2C expression in human brain development. Neuroscience, , 63, , 1067–1079.
Dillon,N. and Festenstein,R. ( (2002) ) Unravelling heterochromatin: competition between positive and negative factors regulates accessibility. Trends Genet., , 18, , 252–258.
Ramirez,A., Milot,E., Ponsa,I., Marcos-Gutierrez,C., Page,A., Santos,M., Jorcano,J. and Vidal,M. ( (2001) ) Sequence and chromosomal context effects on variegated expression of keratin 5/lacZ constructs in stratified epithelia of transgenic mice. Genetics, , 158, , 341–350.(Griet Vanpoucke, Steven Goossens1, Bram )