©1996 by The American Society for Biochemistry and Molecular Biology, Inc.
Regulation of Tissue-specific Expression of the Skeletal Muscle Ryanodine Receptor Gene (*)

(Received for publication, August 22, 1995; and in revised form, December 12, 1995)

Sabine Schmoelzl (1)(§) Tosso Leeb (1) Heinrich Brinkmeier (2) Gottfried Brem (3) Bertram Brenig (1)(¶)

From the  (1)Institute of Veterinary Medicine, University of Göttingen, 37073 Göttingen, Federal Republic of Germany, the (2)Department of General Physiology, University of Ulm, 89069 Ulm, Federal Republic of Germany, and the (3)Institute of Animal Breeding and Genetics, University of Vienna, 1030 Vienna, Austria

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

The ryanodine receptors (RYR) are a family of calcium release channels that are expressed in a variety of tissues. Three genes, i.e. ryr1, ryr2, and ryr3, have been identified coding for a skeletal muscle, cardiac muscle, and brain isoform, respectively. Although, the skeletal muscle isoform (RYR1) was shown to be expressed predominantly in skeletal muscle, expression was also detected in the esophagus and brain. To analyze the transcriptional regulation of the RYR1 gene, we have constructed chimeric genes composed of the upstream region of the RYR1 gene and the bacterial chloramphenicol acetyltransferase (CAT) gene and transiently transfected them into primary cultured porcine myoblasts, myotubes, and fibroblasts. A 443-base pair region upstream from the transcription start site was sufficient to direct CAT activity without tissue specificity. Deletion of a 61-base pair fragment from the 5`-end of the promoter resulted in a marked reduction of CAT activity in all three tissue types. A similar reduction of expression was observed when using a construct with the first intron in antisense orientation upstream from the promoter. In contrast, the first intron in sense orientation enhanced expression only in myotubes, while expression was repressed in fibroblasts and myoblasts. Gel retardation analyses showed DNA binding activity in nuclear extracts for two upstream DNA sequence elements. Our data suggest that (i) RYR1 gene expression is regulated by at least two novel transcription factors (designated RYREF-1 and RYREF-2), and (ii) tissue specificity results from a transcriptional repression in nonmuscle cells mediated by the first intron.


INTRODUCTION

The skeletal muscle ryanodine receptor (RYR1) is a member of a family of calcium release channels that are expressed in different tissues. Three isoforms, RYR1, RYR2, and RYR3, have been isolated and characterized so far. The skeletal muscle isoform was shown to be expressed in skeletal muscle, brain, and esophagus by reverse-transcription-polymerase chain reaction analysis(1) . In skeletal muscle, RYR1 regulates calcium release from the terminal cisternae of the sarcoplasmic reticulum (2, 3, 4, 5, 6) . The physiological release of calcium is controlled by a voltage-gated calcium channel in the membrane of the transverse tubule (dihydropyridine receptor)(7) . To understand the complex mechanisms of excitation-contraction coupling, both calcium channels have been studied in detail. In the last few years, the ryanodine receptor I has gained special interest because it was shown to be involved in malignant hyperthermia, a pharmacogenetic disorder in a variety of species(8, 9) . The porcine RYR1 gene has been assigned to chromosome 6q12(10, 11) . It was shown that a single point mutation in the 5`-region of the RYR1 gene accounts for all cases of malignant hyperthermia in swine(12, 13) .

However, so far, nothing is known about the transcriptional regulation of the skeletal muscle ryanodine receptor gene. In general, activation of muscle-specific genes during skeletal muscle determination and differentiation is regulated by a number of cell type-specific and ubiquitous factors(14, 15) . Most of these factors auto- or transactivate themselves or interact and cooperate with one another in a very complex manner. In principle, there are two groups of transcription factors that are involved in muscle-specific gene expression: the MyoD family of transcription factors, i.e. MyoD, myogenin, Myf-5, and MRF-4 (herculin, Myf-6), and a second group of more ubiquitous factors mainly regulating gene expression in differentiated myocytes. An A/T-rich DNA motif (CTA(A/T)(4)TAG) is present in most promoters or enhancers of muscle-specific genes and is recognized predominantly by the myocyte-enhancing factor 2 (MEF-2). (^1)From a variety of studies, it is known that MEF-2 is essential but not sufficient for muscle-specific expression(16) . In most promoters, additional transcription factor-binding sites are necessary for transactivation. The MEF-3/MAF1 motif (TCAGGTT(A/T)C(A/T)), for example, has been reported to be an important regulatory site in the rat aldolase A gene distal promoter, the mouse myogenin gene promoter, and the skeletal muscle enhancer of the murine cardiac troponin C gene (17, 18, 19) . The primary site of regulation in most muscle-specific genes lies within a relatively short stretch of DNA upstream from the transcription start site. However, regulatory elements have also been identified several kilobase pairs upstream and downstream(20, 21, 22, 23) . Furthermore, the first intron of several genes contains enhancer-like elements that control gene expression(19, 24) . We have been analyzing the genomic structure of the porcine RYR1 gene and have previously reported about the cloning and partial structure of a 80-kilobase pair group of overlapping clones of the porcine ryanodine receptor I locus(25, 26, 27) .

In this study, we have used transient transfection and gel shift assays to identify cis-acting sequences that regulate the expression of the porcine RYR1 gene. We tested the expression of different promoter deletion constructs with or without the first intron, linked to the chloramphenicol acetyltransferase (CAT) gene in primary cultured muscle and nonmuscle cells. The results of these experiments led to the identification of two novel DNA binding activities in nuclear extracts from porcine skeletal muscle. Furthermore, we were able to show that the first intron in sense orientation controls RYR1 gene expression by repressing transcription in nonmuscle cells.


EXPERIMENTAL PROCEDURES

Cloning of the RYR1 Gene Promoter, Plasmid Construction, and Primer Extension Analysis

The construction of the porcine genomic liver DNA library has been described previously(25, 26) . A total of 1.2 times 10^6 recombinant EMBL3A phages were plated, and two replicas were made of each plate. The nitrocellulose filters were hybridized with a [alpha-P]dCTP-labeled fragment of the porcine RYR1 gene. Labeling reactions were performed using a random primer labeling kit (Life Technologies, Inc.). After hybridization, filters were washed twice for 30 min at room temperature in 2 times SSC, 0.1% SDS and twice for 20 min at 68 °C in 0.1 times SSC, 0.1% SDS. Filters were exposed to x-ray films (Eastman Kodak XAR5) at -80 °C for 18 h with intensifying screens (Quanta III, DuPont). Positive clones were isolated and enriched.

Isolated recombinant EMBL3A DNA was digested with different restriction enzymes, and the resulting fragments were subcloned into the polylinker of pGEM-4Z (Promega). Recombinant pGEM-4Z was used to transform Escherichia coli XL1-Blue (Stratagene). For sequence analysis, plasmid DNA was prepared from 10-50-ml liquid cultures using QIAGEN columns according to the supplier's instructions. Fragments were sequenced using a T7 sequencing kit (Pharmacia Biotech Inc.) and alpha-S-dATP as a label. Sequencing reactions were separated on 0.4-mm 8.3 M urea, 5% polyacrylamide sequencing gels. Sequencing gels were processed on a Sparc station 2 (Sun Microsystems) and BioImage (Millipore Corp.). Sequence data were analyzed with MacMolly 3.5 (Soft Gene GmbH). Genomic sequences were aligned with EMBL Nucleotide Sequence Data Bank (Release 38.0) accession numbers X62880 and M91492. The positions of fragments harboring only intron regions were also determined by Southern blotting and hybridization. The complete DNA sequence of the 5`-region of phage RYR56-15.6.2-5 has been assigned to EMBL Nucleotide Sequence Data Bank accession number Z49778 (ID SSCRCRYR1).

For the construction of the RYR1 reporter gene constructs, we used the pGCAT-A and pGCAT-C vectors(28) . Promoter deletion fragments were generated by restriction enzyme digestions using EcoRI (pGCAT-a/E), BglI (pGCAT-a/B), XhoII (pGCAT-a/X), and SacII (pGCAT-a/S). Fragments were isolated after agarose gel electrophoresis, blunted with T4 DNA polymerase according to standard protocols(29) , and cloned into the SmaI site of pGCAT-A or pGCAT-C. A deletion at the 3`-end of the promoter was generated using exonuclease III and nuclease S1 (pGCAT-a/+8). Two constructs were generated by cloning 2356 bp of the first intron in sense (pGCAT-a/Vi+) and antisense (pGCAT-a/Vi-) orientation upstream from the promoter of the construct harboring the complete promoter (pGCAT-a/V).

For primer extension analysis, total RNA was prepared from porcine skeletal muscle according to standard protocols(29) . 5 µg (10 µg) of total RNA was incubated for 5 min at 25 °C and then for 60 min at 42 °C in a 10-µl reaction mixture containing 0.05 M Tris-HCl (pH 8.3), 75 mM KCl, 5 mM MgCl(2), 1 mM dNTPs, 0.3% Nonidet P-40, 10 mM dithiothreitol, 10 fmol of primer, 10 units of RNasin, 100 units of Moloney murine leukemia virus reverse transcriptase, and 10 µCi of [alpha-P]dATP. After reverse transcription, Moloney murine leukemia virus reverse transcriptase was inactivated by incubation at 94 °C for 5 min. RNA was digested with 10 µg of ribonuclease A for 30 min at 37 °C. Reaction mixtures were extracted with phenol and precipitated, and 25,000 cpm were separated on a 8.3 M urea, 5% polyacrylamide sequencing gel.

Cell Culture, DNA Transfection, and CAT Assay

Skeletal muscle tissue (Musculus gluteus superficialis) excised from 4-week-old piglets was used to prepare primary cultures as described previously (30) . Briefly, muscle tissue was cut into small pieces (1 mm^3) using a scalpel, and 500 mg of tissue was incubated in a shaker at 80 rpm and 37 °C for 60-80 min in 5 ml of dissociation buffer (Ham's F-12 medium, 1.5 mg/ml collagenase, 2 mg/ml protease, 2 mM HEPES/NaOH (pH 7.2)). After the tissue was dissociated, the suspension was passed through two layers of nylon gauze with a 50-µm pore size and one layer with a 20-µm pore size. The nylon gauze was washed with phosphate-buffered saline-d, and the filtrates were pooled and centrifuged at 1200 rpm for 5 min to pellet the cells. The cell pellet was resuspended in growth medium (Ham's F-12 medium, 15% fetal calf serum, 2% chicken embryo extract), and 10^6 cells were plated in 250-cm^2 culture flasks and incubated at 37 °C under 5% CO(2) for 20 min. The supernatant containing myoblasts was transferred into a new flask. Fibroblasts were cultured in Dulbecco's modified Eagle's medium supplemented with 5% fetal calf serum. Myoblasts were transfected when reaching half-confluence and then incubated for an additional 24 h. For differentiation, myoblasts were transferred into new culture flasks coated with 5 µg of collagen/cm^2. After reaching half-confluence, myoblasts were transfected and cultured in differentiation medium (Dulbecco's modified Eagle's medium, 5% horse serum). After myotubes were formed, cells were harvested. Cells were transfected by the calcium phosphate precipitation method(31, 32) . 20 µg of plasmid DNA/plate was used in each case. All transfection experiments were done in triplicate. After 4 h of incubation at 37 °C, the medium containing the precipitates was removed, and the cells were incubated for 3 min at 37 °C in 2 ml of 140 mM NaCl, 0.5 mM KCl, 0.55 mM Na(2)HPO(4), 5 mM glucose, 20 mM HEPES/NaOH (pH 7.15) containing 10% glycerol. Cells were harvested and lysed by repeated freezing in ethanol/dry ice and thawing at 37 °C for 5 min. Protein concentrations were determined using the Bio-Rad protein assay based on the Bradford dye-binding procedure(33) .

As an internal standard, the simian virus 40-beta-galactosidase vector pCH110 was used. beta-Galactosidase activity was determined according to (34) using o-nitrophenyl-beta-D-galactopyranoside as substrate. Transcriptional activities of the reporter gene constructs were determined by detection of CAT concentrations using the CAT enzyme-linked immunosorbent assay (Boehringer Mannheim).

Preparation of Nuclear Extracts and Gel Retardation Assay

Nuclear extracts from porcine skeletal muscle were prepared by the procedure described previously(35) . Pelleted nuclei from 10-15 g of muscle tissue were lysed in 40 ml of lysis buffer (20 mM HEPES/NaOH (pH 7.6), 100 mM KCl, 1.5 mM MgCl(2), 0.2 mM EDTA, 0.5 mM dithiothreitol, 0.5 mM phenylmethylsulfonyl fluoride, 0.5 mM spermidine, 0.15 mM spermine, 2 µg/ml leupeptin, 2 µg/ml aprotinin, and 25% glycerol) using a glass Dounce homogenizer. Samples of crude extracts were stored at -80 °C. For further purification, nuclear extracts were fractionated through DEAE- and heparin-Sepharose columns.

Gel retardation assays were performed essentially as described(36) . Double-stranded DNA fragments or chemically synthesized oligonucleotides that were obtained by renaturation were radiolabeled at the 5`-end by polynucleotide kinase and [-P]ATP and used for the binding assay. 10-50 fmol of radiolabeled DNA and nuclear extracts were incubated at 4 °C in binding buffer (50 mM HEPES/NaOH (pH 7.9), 10 mM MgCl(2), 150-300 mM NaCl, 1 mM dithiothreitol, 1 mM EDTA, 10% glycerol) in the presence of 1-2 µg of poly[d(I-C)] and 10-20 µg of bovine serum albumin. After incubation, 0.1 volume of loading dye (625 mM HEPES/NaOH (pH 7.9), 100 mM dithiothreitol, 25% glycerol, 0.05% bromphenol blue, 0.05% xylene cyanol) was added, and the reaction mixtures were electrophoresed through 7% nondenaturing polyacrylamide gels (30:0.8 acrylamide/bisacrylamide) in 0.1 M Tris, 83 mM boric acid, 1 mM EDTA for 4 h. After electrophoresis, the gels were dried, and protein-DNA complexes were visualized by autoradiography.


RESULTS

Isolation and Analysis of the 5`-Region of the RYR1 Gene

The 5`-region of the RYR1 gene was isolated after screening a genomic liver DNA library using a 750-bp genomic fragment of the RYR1 gene. This fragment harbors exons 8 and 9 between positions +635 and +728 and positions +729 and +803, respectively(26) . The DNA sequence of the insert of the recombinant phage (RYR56-15.6.2-5) was determined (Table 1). The genomic structure of the 5`-region of the RYR1 gene is shown in Fig. 1A. The RYR1 promoter lacks a TATA box-like sequence (Fig. 1B). Near the transcription start site, there are several regions that are rich in G and C residues and show homology to the Sp1-binding site. One of these regions is located downstream of the transcription start. A CAAT box motif is present between positions -92 and -88. In addition to these basal sequence elements, an A/T-rich region is located between positions -342 and -328, sharing 79% homology with a known MEF-2-binding site in antisense orientation(37) . This motif was considered as a potential binding site for the myocyte enhancer factor MEF-2. Another DNA motif (GTGCCAGGGATC) 68 bp upstream from the MEF-2 site shows 83% homology to the recently published consensus sequence of MEF-3, which has been found in the neighborhood of MEF-2 in some genes(18, 19) . Within the DNA sequence of the first intron are at least 11 E box motifs. Notably, six of these sites occur as three pairs with a distance of 23 or 24 bp between members of each pair (Fig. 1C).




Figure 1: Genomic organization of porcine ryanodine receptor (RYR1) gene coding region 1-540 and DNA sequence of the promoter region. A, the diagram outlines the genomic structure, sequencing strategy, positions of subclones, and partial restriction map of RYR56-15.6.2-5. Exons are numbered (1-6) and are shown as boxes. Introns are shown as solid lines. Thin lines represent the lengths and locations of subclones. B, the nucleotide sequence of the 5`-untranslated region, the first exon, and 5 base pairs of the first intron of the RYR1 gene are shown. Homologies to transcription factor and CAAT box-binding consensus sites are boxed. The positions of fragments B and D and oligonucleotides A1, A2, A3, A4, A5, and MEF-2 are underlined. +1 denotes the transcription start site at the GT dinucleotide. The location of primer RPr30 is shown. C, the diagram illustrates the positions of potential E boxes within the first intron of the RYR1 gene. The first exon is shown as a closed box. E boxes are shown as hatched boxes.



To determine the transcription start site, we performed a primer extension analysis. Total RNA was extracted from porcine skeletal muscle. A primer (RPr30, 5`-CTC ATC TTC GCC CTC TCC TC-3`) corresponding to positions +11 to +30 of the coding region of the first exon (Fig. 1B) was annealed to the RNA and extended with Moloney murine leukemia virus reverse transcriptase. The elongated product was separated on a polyacrylamide sequencing gel along with a sequencing reaction using the same primer and the isolated genomic DNA fragment as template. As shown in Fig. 2, there is a single band detectable 109/110 bp upstream from the ATG translation initiation codon of the first exon. The extended product ends at a GT dinucleotide.


Figure 2: Determination of the transcription start site by primer extension analysis. Primer extension of the 5`-end of the RYR1 mRNA was carried out with the 20-nucleotide oligonucleotide RPr30 (nucleotides +11 to +30) to prime the reverse transcription. The sequencing reaction on the left side of the autoradiogram was carried out using RPr30 on a DNA template. Lanes 1 and 2 show primer extensions carried out with 5 and 10 µg of total skeletal muscle RNA, respectively. Note that the DNA sequence is that of the noncoding DNA strand. The direction of transcription is indicated on the left side.



Transfection of RYR1 CAT Genes into Skeletal Muscle and Nonmuscle Cells

To identify transcriptionally important regulatory regions of the RYR1 promoter, we cloned CAT reporter gene constructs and transiently transfected them into porcine primary cultures of fibroblasts, myoblasts, and myotubes. A full-length 537-bp BamHI/AvaI promoter fragment was isolated and cloned into the SmaI site of pGCAT-A. The AvaI site is located 15 bp upstream from the ATG translation initiation codon of the first exon. Deletion mutants were generated employing restriction enzymes as described above. The orientation of the inserts was determined by DNA sequencing. The resulting plasmids are shown in Fig. 3. The construct harboring the antisense-oriented full-length promoter fragment pGCAT-a/V- was used as a negative control. CAT activities were normalized to transfection efficiency as determined by o-nitrophenyl-beta-D-galactopyranoside assay and related to pGCAT-a/V-. As shown in Fig. 3A, the full-length construct pGCAT-a/V was sufficient to activate CAT expression without directing tissue specificity. Activation compared with the negative control was 13-fold in fibroblasts, 10-fold in myoblasts, and 30-fold in myotubes. Relative to the cytomegalovirus promoter, the level of activation was 16% in fibroblasts, 18% in myoblasts, and 13% in myotubes. A deletion between positions +9 and +108 (pGCAT-a/+8) reduced the CAT activity almost to background levels. This region harbors several GC-rich elements (Fig. 3A). A deletion of 61 bp from the 5`-end of the fragment (pGCAT-a/E) reduced CAT expression to background levels in fibroblasts, myoblasts, and myotubes. Further deletions of the promoter had only marginal influence on expression levels. A relative increase in CAT activity was observed in fibroblasts when using the pGCAT-a/S construct (Fig. 3A).


Figure 3: Transient transfection experiments with promoter constructs in primary cultures of porcine myoblasts, myotubes, and fibroblasts. The diagrams outline the structure of the different promoter deletion constructs. Putative DNA-binding motifs are indicated with boxes. +1 denotes the transcription start site. 20 µg of the indicated constructs was cotransfected with pCH110 DNA. Transcriptional activities of the reporter gene constructs were determined by detection of CAT concentrations using the CAT enzyme-linked immunosorbent assay. CAT activity is expressed as a percentage of activity of cytomegalovirus-CAT, used as a positive control, following normalization for DNA uptake and as described under ``Experimental Procedures.'' CAT activity was normalized to 1. Each value represents the mean ± S.D. for three independent transfection experiments.



To analyze the influence of the first intron containing 11 E box motifs, we transfected two constructs in which 2.3 kilobases of the first intron had been cloned in opposite orientations upstream from the RYR1 promoter (pGCAT-a/Vi+ and pGCAT-a/Vi-). When using pGCAT-a/Vi+ in transfections, CAT expression was reduced 80-fold in fibroblasts and 15-fold in myoblasts. In contrast, expression of pGCAT-a/Vi+ in myotubes was identical to that of the full-length promoter construct pGCAT-a/V (Fig. 3B). When we transfected the construct pGCAT-a/Vi-, carrying the intron fragment in antisense orientation, no CAT expression was detectable in any of the three cell types (Fig. 3B).

Identification of Nuclear Factor-binding Elements

From our transfection studies, we concluded that the region between positions -443 and -383 is important for transcriptional regulation of the RYR1 promoter. To identify protein-binding elements in the RYR1 promoter, we performed a series of gel shift assays. We prepared two RYR1 promoter fragments (B and D) by digestion with appropriate restriction enzymes. Protein binding to both fragments was tested with nuclear and cellular extracts from porcine skeletal muscle because they contain the putative transcription factor-binding motifs and therefore were expected to form protein-DNA complexes. To reduce nonspecific DNA binding, we purified the nuclear extracts on DEAE- and heparin-Sepharose. Fragment D formed too many protein-DNA complexes to distinguish any particular interaction. With fragment B, at least four protein-DNA complexes were formed in the presence of poly[d(I-C)] and the heterologous fragment D (Fig. 4). One complex was completely inhibited when using unlabeled fragment B as homologous competitor, whereas the others partly remained and were therefore seen as nonspecific. To analyze whether the specific protein-DNA complex formed with fragment B was due to MEF-2 binding, we used a synthetic oligonucleotide harboring the MEF-2 motif within the RYR1 promoter for gel mobility shift analysis (Fig. 1B). A complex formed that was inhibited by homologous competition in a 40-fold surplus of unlabeled oligonucleotide. Surprisingly, formation of the protein-DNA complex was not inhibited in the presence of unlabeled fragment B, which includes the sequence of oligonucleotide MEF-2 (data not shown).


Figure 4: Gel retardation assay with skeletal muscle nuclear extracts and fragment B. Extracts were prepared from skeletal muscle tissue, and 20 µl was analyzed for binding to 50 fmol of P-labeled fragment B containing the MEF-2-binding motif. 0.5 µg of poly[d(I-C)] was added to each reaction. 200 fmol of unlabeled fragment B was used as specific competitor (lane 3). Nonspecific competition was done with 200 fmol of unlabeled fragment D (lane 4). The specific protein-DNA complex is indicated with an arrow. The protein-DNA complexes were resolved on 7% nondenaturing polyacrylamide gels. FP, free probe.



To localize protein-binding sequence elements in the modulating region of the 5`-region of the RYR1 promoter, we used overlapping oligonucleotides spanning region -442 to -321 in gel mobility shift assays (Fig. 1B). As shown in Fig. 5, a complex was formed with oligonucleotide A4 that was inhibited in the presence of 0.5 µg of nonspecific competitor poly[d(I-C)] and 200 fmol of specific competitor A4. When using 0.5 µg of poly[d(A-T)] as nonspecific competitor, complex formation was not inhibited. Oligonucleotide A4 does not contain any known protein-binding sequence elements. When using oligonucleotide A5 in gel mobility shift assays, a specific protein-DNA complex was formed that was not inhibited in the presence of 0.5 µg of nonspecific competitor poly[d(I-C)] or poly[d(A-T)], but was inhibited in the presence of 200 fmol of unlabeled oligonucleotide A5. Complex formation was slightly enhanced in the presence of 200 fmol of unlabeled oligonucleotide MEF-2 (Fig. 6, lane 6). Oligonucleotides A1, A2 (harboring the putative MEF-3 consensus motif), and A3 did not form protein-DNA complexes with nuclear or whole cell extracts.


Figure 5: Gel retardation assay with skeletal muscle extracts and oligonucleotide A4. Cellular extracts were prepared from skeletal muscle tissue, and 25 µl was analyzed for binding to 50 fmol of P-labeled oligonucleotide A4. 0.5 µg of poly[d(I-C)] was added to the reactions in lanes 3 and 5, and 0.5 µg of poly[d(A-T)] was added to the reaction in lane 4. 200 fmol of unlabeled oligonucleotide A4 was used as specific competitor (lane 5). The specific protein-DNA complex is indicated with an arrow. The protein-DNA complexes were resolved on 7% nondenaturing polyacrylamide gels. FP, free probe.




Figure 6: Gel retardation assay with skeletal muscle extracts and oligonucleotide A5. Cellular extracts were prepared from skeletal muscle tissue, and 10 µl was analyzed for binding to 50 fmol of P-labeled oligonucleotide A5. 0.5 µg of poly[d(I-C)] was added to the reactions in lanes 3, 5, and 6. 0.5 µg of poly[d(A-T)] was added to the reaction in lane 4. 200 fmol of unlabeled oligonucleotide A5 was used as specific competitor (lane 5). Heterologous competition was done with 200 fmol of unlabeled oligonucleotide MEF-2 (lane 6). The specific protein-DNA complex is indicated with an arrow. The protein-DNA complexes were resolved on 7% nondenaturing polyacrylamide gels. FP, free probe.




DISCUSSION

In this report, we describe the isolation and transcriptional analysis of the porcine skeletal muscle ryanodine receptor (RYR1) gene promoter. The RYR1 promoter does not contain a TATA box, which is in agreement with the human and rabbit RYR1 gene promoters. Promoters lacking a TATA box have been first described in housekeeping genes and more recently found in various genes as in the human slow twitch skeletal muscle troponin I gene (38) as well as in nonmuscle-specific receptor genes such as the low density lipoprotein receptor-related protein/alpha(2)-macroglobulin receptor gene(19, 39) . In promoters lacking a TATA box, GC-rich elements play an important role in transcriptional activation. There are several GC-rich elements in the RYR1 promoter region that share homology with the Sp1 consensus motif in sense and antisense orientation. A GC-rich region that is situated between the transcription and translation start sites is crucial in the RYR1 gene promoter. In our experiments, we were able to show that the deletion of this GC-rich region abolishes CAT expression in fibroblasts, myoblasts, and myotubes. Whether these GC-rich elements are directly involved in the crucial role of this region within the RYR1 gene promoter or whether it is merely a positional effect due to the requirement of correct spacing between the transcription and translation start sites has not been investigated.

Similar to the aldolase A gene, the RYR1 gene promoter contains a single putative MEF-2 consensus site, but no E box(18) . In contrast to other promoters, the MEF-2 consensus site within the RYR1 promoter is in antisense orientation(18, 21, 37, 40, 41) . In our transfection studies, we found CAT expression in fibroblasts, myoblasts, and myotubes when using the construct pGCAT-a/V, which harbors the putative MEF-2 consensus site. Therefore, we conclude that the putative MEF-2 site within the 552-bp promoter region is not sufficient to direct tissue-specific expression. This is in agreement with other studies that have shown that activation of muscle-specific expression by MEF-2 is mediated by the interaction of different muscle-specific and ubiquitous factors with MEF-2(16, 17, 18, 19, 21) . So far, only one promoter has been described in which a single MEF-2 site was sufficient for muscle-specific expression(40) . Furthermore, when deleting region -442 to -382 upstream from the MEF-2 consensus site, no expression was detectable in fibroblasts, myoblasts, or myotubes. Thus, the presence of the putative MEF-2 consensus site alone was not sufficient to confer transcriptional activation. However, in gel mobility shift assays, we found formation of a specific protein-DNA complex with an oligonucleotide harboring the putative MEF-2 motif of the RYR1 promoter. Interestingly, competition with the RYR1 promoter fragment B harboring the same motif did not inhibit formation of the protein-DNA complex. Therefore, we addressed the question whether there are additional factors binding to fragment B that could exhibit cooperative binding effects.

Using a set of overlapping oligonucleotides spanning region -442 to -321, we found specific DNA binding activity to oligonucleotide A5, which is derived from the sequence immediately upstream from the putative MEF-2 motif. In the presence of unlabeled oligonucleotide MEF-2, complex formation was increased, indicating cooperative binding of MEF-2 and a so far unknown protein, similar to previous findings for MEF-2(16, 17, 18, 19, 21) . In addition, we found protein binding activity with oligonucleotide A4, which was derived from the RYR1 sequence upstream from oligonucleotide A5 and which bridges the region between pGCAT-a/V and pGCAT-a/E. No other DNA binding activity was detectable in whole cell and nuclear extracts in the farther upstream region covered by oligonucleotides A1, A2, and A3. In particular, there was no protein-DNA complex formed with oligonucleotide A2, containing the putative MEF-3-binding motif. It has been reported that the sequence requirements for MEF-3 binding are not very specific(18) . Therefore, we conclude that there is no MEF-3-like binding affinity for this region of the promoter. Combining the results from the transfection experiments and the gel retardation analysis, we conclude that there are DNA-binding factors recognizing so far uncharacterized sequence elements in the promoter region between positions -395 and -363 (A4) and positions -370 and -340 (A5). With the partial deletion of region A4 in construct pGCAT-a/E, one of the potential binding sites was removed, resulting in the loss of promoter activity. For the factor binding to A5, we concluded from our gel mobility experiments cooperative binding with MEF-2. Therefore, these factors possibly interact by protein/protein contacts with MEF-2 and are required for transcriptional activity of the RYR1 gene promoter. As there are no known protein-binding elements contained in the sequences of A4 and A5, we designated the binding factors as RYREF-1 and RYREF-2, respectively.

The 552-bp promoter fragment used in pGCAT-a/V was not sufficient to direct muscle-specific expression. Although in most muscle-specific genes reported so far, a relatively short stretch of DNA upstream from the transcription start site is sufficient for tissue-specific expression, there are some examples where far distant upstream regions influence muscle specificity(21, 22, 42) . In other muscle-specific genes, elements located in the first intron contribute to tissue specificity(19, 24) . For the RYR1 gene, we found that a construct harboring 2.3 kilobases of the first intron in addition to the 552-bp promoter fragment was able to direct tissue specificity by completely repressing expression in fibroblasts and myoblasts. In contrast, expression in the differentiated myotubes was found at the same levels as with the promoter fragment alone. This effect was only seen when the fragment of the first intron was sense-orientated. In contrast, the antisense-oriented first intron fragment repressed the expression in fibroblasts and myoblasts as well as in myotubes. The only known sequence motifs in the 2.3 kilobases of the first intron are 11 E box motifs, which occur in three pairs with even spacing and five separate motifs. Although E box motifs have a palindromic sequence and therefore no orientation, it is known that the flanking sequences can influence the binding of basic helix-loop-helix proteins, which bind in their transcriptional activating form as heterodimers. In fact, the importance of sequence orientation of E boxes was discussed recently by Ma et al.(43) . Furthermore, it has been shown in vitro that myogenin interacts in cooperative DNA binding with MEF-2 (44) and that an E box-binding factor functionally interacts with MEF-2 in the desmin gene promoter(21) . Thus, although we cannot exclude the possibility that other cis-acting elements exist farther upstream, our experiments show that the E box-containing fragment of the first intron is able to direct gene expression specifically in differentiated muscle cells. However, further analysis will be necessary to confirm our findings and to characterize the novel factors RYREF-1 and RYREF-2. Furthermore, the regulation of the RYR1 gene might be even more complicated because it is expressed not only in skeletal muscle, but also in restricted areas of the brain(1) .

In conclusion, 552 bp of the 5`-untranslated region of the RYR1 promoter are sufficient to direct expression without tissue specificity. The region from positions -442 to -382 is absolutely required for expression. A putative MEF-3 motif in this region seems to be insignificant for transcriptional regulation as determined by gel retardation analysis. Our data suggest the existence of two additional factors (one of which seems to interact directly with MEF-2) binding upstream from the MEF-2 motif to a region that is required for RYR1 gene promoter activity. Tissue specificity is likely to be conferred by the presence of the first intron in sense orientation presumably via interaction of myogenic bHLH factors that bind to E boxes within the first intron.

Only recently has the influence of cAMP, protein kinase C, and neural factors on the expression of RYR1 and the dihydropyridine receptor been reported(45) . These data suggest that RYR1 transcription in rat skeletal muscle is regulated by so far unidentified neural factors. It will be interesting to determine whether the two novel factors suggested by our studies, RYREF-1 and RYREF-2, might represent or be related to these neural factors.


FOOTNOTES

*
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Present address: Dept. of Biomedical Sciences, University of Guelph, Guelph, Ontario N1G2W1, Canada.

To whom correspondence should be addressed: Inst. of Veterinary Medicine, Groner Landstrasse 2, 37073 Göttingen, Germany. Tel.: 49-551-39-3383; Fax: 49-551-39-3399; :bbrenig{at}gwdg.de.

(^1)
The abbreviations used are: MEF-2, myocyte-enhancing factor 2; CAT, chloramphenicol acetyltransferase; bp, base pair(s).


ACKNOWLEDGEMENTS

We are grateful to E.-L. Winnacker for providing excellent working conditions in his laboratory. Thanks are extended to K. Barnewitz for expert technical assistance.


REFERENCES

  1. Ledbetter, M. W., Preiner, J. K., Louis, C. F., and Mickelson, J. R. (1994) J. Biol. Chem. 269, 31544-31551 [Abstract/Free Full Text]
  2. Hymel, L., Inui, M., Fleischer, S., and Schindler, H. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 441-445 [Abstract]
  3. Inui, M., Saito, A., and Fleischer, S. (1987) J. Biol. Chem. 262, 15637-15642 [Abstract/Free Full Text]
  4. Inui, M., Saito, A., and Fleischer, S. (1987) J. Biol. Chem. 262, 1740-1747 [Abstract/Free Full Text]
  5. Marks, A. R., Tempst, P., Hwang, K. S., Taubman, M. B., Inui, M., Chadwick, C., Fleischer, S., and Nadal, G. B. (1989) Proc. Natl. Acad. Sci. U. S. A. 86, 8683-8687 [Abstract]
  6. Pessah, I. N., Francini, A. O., Scales, D. J., Waterhouse, A. L., and Casida, J. E. (1986) J. Biol. Chem. 261, 8643-8648 [Abstract/Free Full Text]
  7. Yuan, S. H., Arnold, W., and Jorgensen, A. O. (1991) J. Cell Biol. 112, 289-301 [Abstract]
  8. Fill, M., Coronado, R., Mickelson, J. R., Vilven, J., Ma, J. J., Jacobson, B. A., and Louis, C. F. (1990) Biophys. J. 57, 471-475 [Abstract]
  9. MacLennan, D. H., and Phillips, M. S. (1992) Science 256, 789-794 [Medline] [Order article via Infotrieve]
  10. Harbitz, I., Chowdhary, B., Thomsen, P. D., Davies, W., Kaufmann, U., Kran, S., Gustavsson, I., Christensen, K., and Hauge, J. G. (1990) Genomics 8, 243-248 [Medline] [Order article via Infotrieve]
  11. Chowdhary, B. P., Thomson, P. D., Harbitz, I., Landset, M., and Gustavsson, I. (1994) Cytogenet. Cell Genet. 76, 211-214
  12. Fujii, J., Otsu, K., Zorzato, F., deLeon, S., Khanna, V. K., Weiler, J. E., O'Brien, P. J., and MacLennan, D. H. (1991) Science 253, 448-451 [Medline] [Order article via Infotrieve]
  13. Otsu, K., Khanna, V. K., Archibald, A. L., and MacLennan, D. H. (1991) Genomics 11, 744-750 [Medline] [Order article via Infotrieve]
  14. Lassar, A., and Münsterberg, A. (1994) Curr. Opin. Cell Biol. 6, 432-442 [Medline] [Order article via Infotrieve]
  15. Lassar, A. B., Skapek, S. X., and Novitch, B. (1994) Curr. Opin. Cell Biol. 6, 788-794 [Medline] [Order article via Infotrieve]
  16. Lee, K. J., Hickey, R., Zhu, H., and Chien, K. R. (1994) Mol. Cell. Biol. 14, 1220-1229 [Abstract]
  17. Edmondson, D. G., Cheng, T. C., Cserjesi, P., Chakraborty, T., and Olson, E. N. (1992) Mol. Cell. Biol. 12, 3665-3677 [Abstract]
  18. Hidaka, K., Yamamoto, I., Arai, Y., and Mukai, T. (1993) Mol. Cell. Biol. 13, 6469-6478 [Abstract]
  19. Parmacek, M. S., Ip, H. S., Jung, F., Shen, T., Martin, J. F., Vora, A. J., Olson, E. N., and Leiden, J. M. (1994) Mol. Cell. Biol. 14, 1870-1885 [Abstract]
  20. Biben, C., Kirschbaum, B. J., Garner, I., and Buckingham, M. (1994) Mol. Cell. Biol. 14, 3504-3513 [Abstract]
  21. Li, H., and Capetanaki, Y. (1994) EMBO J. 13, 3580-3589 [Abstract]
  22. van de Klundert, F., Jansen, H. J., and Bloemendal, H. (1994) J. Biol. Chem. 269, 220-225 [Abstract/Free Full Text]
  23. Dunwoodie, S. L., Joya, J. E., Arkell, R. M., and Hardeman, E. C. (1994) J. Biol. Chem. 269, 12212-12219 [Abstract/Free Full Text]
  24. Christensen, T. H., Prentice, H., Gahlmann, R., and Kedes, L. (1993) Mol. Cell. Biol. 13, 6752-6765 [Abstract]
  25. Brenig, B., and Brem, G. (1992) FEBS Lett. 298, 277-279 [CrossRef][Medline] [Order article via Infotrieve]
  26. Leeb, T., Schmoelzl, S., Brem, G., and Brenig, B. (1993) Genomics 18, 349-354 [CrossRef][Medline] [Order article via Infotrieve]
  27. Schmoelzl, S., Leeb, T., Brem, G., and Brenig, B. (1993) Life Sci. Adv. 12, 1-11
  28. Frebourg, T., and Brison, O. (1988) Gene (Amst.) 65, 315-318
  29. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K. (1990) Current Protocols in Molecular Biology , John Wiley & Sons, Inc., New York
  30. Brinkmeier, H., Seewald, M. J., Eichinger, H. M., and Rüdel, R. (1993) J. Am. Sci. 71, 1154-1160
  31. Graham, F. L., and van der Eb, A. J. (1973) Virology 52, 456-467 [Medline] [Order article via Infotrieve]
  32. Wigler, M., Pellicer, A., Silverstein, S., and Axel, R. (1978) Cell 14, 725-731 [Medline] [Order article via Infotrieve]
  33. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254 [CrossRef][Medline] [Order article via Infotrieve]
  34. Bignon, C., Daniel, N., and Djiane, J. (1993) BioTechniques 15, 243-245 [Medline] [Order article via Infotrieve]
  35. Mar, J. H., and Ordahl, C. P. (1990) Mol. Cell. Biol. 10, 4271-4283 [Medline] [Order article via Infotrieve]
  36. Schneider, R., Gander, I., Müller, U., Mertz, R., and Winnacker, E.-L. (1986) Nucleic Acids Res. 14, 1303-1317 [Abstract]
  37. Cserjesi, P., and Olson, E. N. (1991) Mol. Cell. Biol. 11, 4854-4862 [Medline] [Order article via Infotrieve]
  38. Corin, S. J., Juhasz, O., Zhu, L., Conley, P., Kedes, L., and Wade, R. (1994) J. Biol. Chem. 269, 10651-10659 [Abstract/Free Full Text]
  39. Kim, J. H., Johansen, F. E., Robertson, N., Catino, J. J., Prywes, R., and Kumar, C. C. (1994) J. Biol. Chem. 269, 13740-13743 [Abstract/Free Full Text]
  40. Nakatsuji, Y., Hidaka, K., Tsujino, S., Yamamoto, Y., Mukai, T., Yanagihara, T., Kishimoto, T., and Sakoda, S. (1992) Mol. Cell. Biol. 12, 4384-4390 [Abstract]
  41. Zhou, M.-D., Goswami, S. K., Martin, M. E., and Siddiqui, M. A. Q. (1993) Mol. Cell. Biol. 13, 1222-1231 [Abstract]
  42. Gekakis, N., and Sul, H. S. (1994) Biochemistry 33, 1771-1777 [Medline] [Order article via Infotrieve]
  43. Ma, P. C. M., Rould, M. A., Weintraub, H., and Pabo, C. O. (1994) Cell 77, 451-459 [Medline] [Order article via Infotrieve]
  44. Funk, W. D., and Wright, W. E. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 9484-9488 [Abstract]
  45. Ray, A., Kyselovic, J., Leddy, J. J., Wigle, J. T., Jasmin, B. J., and Tuana, B. S. (1995) J. Biol. Chem. 270, 25837-25844 [Abstract/Free Full Text]

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