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1 Division of Integrative Cell Biology, Institute of Molecular Embryology and Genetics, Kumamoto University, Kumamoto 860-0811, Japan
2 Division of Biological Functional Science, Graduate School of Life and Environmental Science, University of Tsukuba, Ibaraki 305-8572, Japan
3 Division of Stem Cell Regulation, The Institute of Medical Science, The University of Tokyo, Tokyo 108-8639, Japan
4 Department of Life Sciences, The University of Tokyo, Tokyo 153-8902, Japan
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Abstract |
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 Top Abstract IntroductionResultsDiscussionExperimental proceduresReferences |
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Introduction |
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The molecular mechanisms of action of Sall1 and SALL1 are, however, still obscure. Sall1 functions as a transcriptional repressor, as determined by luciferase reporter assays, and interacts physically with histone deacetylase (HDAC) and other components of the chromatin remodeling NuRD complex. However, it remains to be resolved whether the transcriptional repression is solely dependent on the HDAC activity (Netzer et al. 2001; Kiefer et al. 2002). Both human SALL1 and mouse Sall1 localize to pericentric heterochromatin (Netzer et al. 2001; Kiefer et al. 2002; Sato et al. 2004), a highly condensed chromatin structure that exists near the centromeric region (Horz & Altenburger 1981; Wong & Rattner 1988). In mouse cells, pericentric heterochromatin is mainly composed of a repetitive DNA sequence called the major satellite repeat, and some proteins, including heterochromatin protein 1 (HP1) that maintain the heterochromatin structure. The histones in this region are highly methylated, especially the 9th lysine of the histone H3 tail (H3K9) (Nielsen et al. 2001; Lachner et al. 2003; Peters et al. 2003). The centromere is known to function in the organization of correct chromosome segregation, whereas the functions of pericentric heterochromatin remain largely unknown. Recently, an association between transcriptional repression and SALL1 localization to heterochromatin was reported (Netzer et al. 2006), indicating that the functions of Sall1 may depend on its localization to heterochromatin.
Here, we focused on the zinc finger domains of Sall1, and determined the mechanism of its localization to heterochromatin. Sall1 encodes a protein that contains ten zinc finger motifs. The most N-terminal zinc finger is a single C2HC-type zinc finger that is only conserved in vertebrates (Drosophila sal does not have this N-terminal C2HC zinc finger). The other zinc fingers are of the C2H2-type and arranged as doublets (double zinc fingers) or triplets. We demonstrate that Sall1 requires its C-terminal double zinc finger domains for localization to heterochromatin, and that these domains recognize A/T-rich nucleotide sequences in the major satellite DNA of heterochromatin.
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Results |
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Over-expressed Sall1 protein has been reported to localize to pericentric heterochromatin in cultured cell lines (Netzer et al. 2001; Kiefer et al. 2002; Sato et al. 2004). However, the localization of endogenous Sall1 remains unclear. Since embryonic stem (ES) cells expressed abundant Sall1 protein as evaluated by Western blot analysis (data not shown), we examined the localization of endogenous Sall1 in ES cells using an anti-Sall1 antibody. Heterochromatin regions can be identified by bright spots of 4,6-diamidino-2-phenylindole (DAPI) staining (DAPI-dense clusters) in the nuclei of several mouse cell lines. We also used anti-HP1
and anti-tri-methylated H3K9 antibodies to visualize the regions of heterochromatin. The staining for Sall1 revealed a dotted pattern in the nucleus that overlapped with the staining for HP1 and tri-methylated H3K9 (Fig. 1A), indicating that endogenous Sall1 is localized to heterochromatin. NIH 3T3 cells do not express Sall1 and serve as a tool for assessing the distribution of exogenous Sall1. When wild-type or GFP-fused Sall1 was introduced into NIH3T3 cells, both proteins localized to heterochromatin (Fig. 1B and data not shown). Thus the Sall1-GFP fusion protein is likely to be useful for dissecting the localization mechanisms of Sall1 to heterochromatin.
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(Fig. 1D; data not shown), indicating that it is not localized to heterochromatin. Zn2-5mut, a similar mutant except for the N-terminal C2HC zinc finger motif, showed a similar localization pattern to Zn1-5mut. In contrast, a Zn1 mutant that retained all the zinc fingers except for the N-terminal one was localized to heterochromatin. These results indicate that four clusters of the C2H2 zinc finger domains of Sall1 (Zn2, Zn3, Zn4 and Zn5) are necessary for the localization, whereas Zn1 is not. Localization of Sall1 to heterochromatin requires the C-terminal double zinc fingers Zn4 and Zn5
To determine which zinc finger domains of Sall1 are essential for its localization to heterochromatin, we produced a series of mutants (Zn345mut, Zn245mut, Zn235mut and Zn234mut), which each returned one intact zinc finger domain cluster to the mutant Zn2-5mut (Fig. 2A). However, none of these single C2H2 double zinc finger domains completely restored the localization of Sall1 to heterochromatin. Among the mutants, Zn234mut, which had an intact Zn5, partially localized to heterochromatin (Fig. 2A, red box; Fig. 2D), suggesting that Zn5 may be the most responsible cluster for the Sall1 localization. Next, we prepared another series of mutants (Zn45mut, Zn35mut, Zn34mut, Zn25mut, Zn24mut and Zn23mut), which each returned two intact zinc finger clusters to the mutant Zn2-5mut (Fig. 2B). Zn25mut, which had mutations in both Zn2 and Zn5, still diffused uniformly throughout the nucleus except for the spots that were positive for DAPI or tri-methylated H3K9 (Fig. 2B, upper red box; Fig. 2D), while the other four mutants (Zn45mut, Zn35mut, Zn34mut and Zn24mut) were partially localized to heterochromatin. Strikingly, Zn23mut, which had intact Zn4 and Zn5 double zinc finger clusters, was completely localized to heterochromatin, similar to the case for wild-type Sall1-GFP (Fig. 2B, lower red box; Fig. 2D). Further disruption of Zn1 (Zn123mut) did not affect the localization. These results suggest that Zn4 and Zn5 are sufficient for heterochromatin localization. Next, we constructed mutants in which only one zinc finger cluster was mutated. Although mutations in Zn2 or Zn3 had no effects, mutations in either Zn4 or Zn5, especially Zn5, caused localization not only in heterochromatin but also in other nuclear regions that were negative for DAPI or tri-methylated H3K9 (Fig. 2C, red box; Fig. 2D). Thus the C-terminal two zinc finger clusters (Zn4 and Zn5), especially Zn5, are important for the heterochromatin localization of Sall1 protein.
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Next, we produced GFP-fusion proteins of all the other Sall family members, and introduced them into NIH3T3 cells. Sall1 and Sall3 have five zinc finger clusters (ten zinc fingers in total), while Sall2 and Sall4 have four clusters (eight fingers in total). Interestingly, Sall4 showed a similar localization pattern to Sall1 (Fig. 3A). Although a chick Sall3 homolog, csal-3, was reported to be localized in the cytoplasm (Sweetman et al. 2003), mouse Sall3 was localized in both the cytoplasm and the nucleus. Sall3 in the nucleus was localized to heterochromatin in a similar pattern to Sall1 and Sall4. Sall2 was localized in the nucleus, but showed a quite different pattern from the other Sall family members, since it diffused uniformly throughout the nuclear compartments except for the DAPI-positive spots (Fig. 3A). To examine whether these localization differences were due to the sequences of the zinc finger motifs, we aligned the amino acid sequences of all the C2H2 double zinc finger domains among the four Sall family proteins (Fig. 3B). C2H2 zinc finger motifs are comprised of a ß-strand and an -helix. The -helix (referred to as the recognition helix) contacts bases in the major groove of DNA, and four amino acids at positions –1, 2, 3 and 6 in the recognition -helix were reported to define the four nucleotides recognized by C2H2 zinc finger domains (Pabo et al. 2001). The amino acids at these positions in Sall1, Sall3 and Sall4 are highly conserved, and in particular, the most C-terminal zinc fingers (Zn5 of Sall1 and Sall3, Zn4 of Sall4) have the same amino acid residues at these positions. In contrast, the corresponding amino acids in Zn4 of Sall2 are quite different from those of the other family members (Fig. 3B). Therefore, the heterochromatin localization of other Sall family members may be dependent on the sequence of the recognition -helix of their most C-terminal double zinc finger. To confirm this hypothesis in Sall1, we changed the amino acid residues in the recognition -helix of each of the zinc finger clusters of Sall1 to those of Sall2 Zn4 (-helix mut; Fig. 3B, asterisks). As expected, the localization of the -helix mutants showed similar patterns to the equivalent cysteine residue mutants (Fig. 2, -helix mut). Considering the proposed role of the -helix in the binding to DNA, our results suggest that at least in Sall1, the double zinc finger motifs may recognize some specific nucleotide sequences that could exist in heterochromatin.
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To investigate whether the zinc fingers of Sall1 recognize specific nucleotide sequences, we performed the cyclic amplification and selection of targets (CASTing) technique as a binding site selection assay (Wright et al. 1991). For this assay, we used a bacterially-expressed truncate of Sall1 fused with glutathione-S-transferase (GST) (GST-Zn5; Fig. 4A). This truncate contained Zn5, which was identified as the most important zinc finger for heterochromatin localization (Fig. 2). The fusion protein was bound to glutathione-Sepharose beads (Fig. 4B) and then mixed with randomized double-stranded (ds) oligonucleotides. The dsDNA pulled down by GST-Zn5 was subjected to PCR amplification. After four cycles of the selection process, we carried out electrophoretic mobility shift assays (EMSAs) to examine the enhancement of the protein-DNA affinity. The dsDNA obtained after selection showed significantly higher affinity than the initial oligonucleotides (Fig. 4C), and no binding was detected when GST was used as a negative control. When the dsDNA was cloned into a vector and sequenced, A/T-rich sequences were obtained (Fig. 5A). From an alignment of these sequences, we propose the putative consensus sequences ATAA(A/T)(A/T) (Fig. 5B).
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To determine whether Sall1 associates with major satellite DNA, we performed EMSAs using nuclear extracts from HEK293 cells expressing Sall1. Sall1 bound strongly to major satellite DNA as a clear band, indicating that a Sall1-DNA complex was formed. In contrast, a control nuclear extract (mock) with GFP expression showed no complex formation (Fig. 6B, lanes 1, 2). To examine which region of the major satellite DNA Sall1 associates with, we divided the major satellite DNA into three fragments, designated as Major A, B and C (Fig. 6A). Sall1 associated with all three fragments, but the strongest interaction was observed with Major B (Fig. 6B, lanes 3–8). The band for the Sall1-DNA complex was supershifted upon addition of an anti-Sall1 antibody (-Sall1), and disappeared after the addition of excess cold major satellite DNA (Fig. 6C, lanes 1–5). These observations indicate that Sall1 specifically associates with major satellite DNA. To confirm this association, we performed EMSAs using nuclear extracts containing several zinc finger mutants. Zn23mut showing heterochromatin localization (Fig. 2B) associated with both full-length major satellite DNA (data not shown) and the Major B fragment, while Zn2-5mut and Zn45mut showing diffuse distributions in the nucleus (Fig. 2B) did not (Fig. 6C, lanes 6–8). These findings indicate that Sall1 binds to major satellite DNA via its C-terminal double zinc fingers.
Sall1 recognizes A/T-rich sequences present in major satellite DNA
Since Sall1 had a higher affinity for the Major B fragment than the other two fragments (Fig. 6B), we produced four deletion probes deleted from the 5'- or 3'-end of the Major B probe (B-1, B-2, B-3 and B-4; Fig. 7A), and performed EMSAs using these deletion probes. The deletion probes B-2, B-3 and B-4, which contained multiple A/T-rich sequences similar to the Zn5 consensus sequences, had stronger affinities for Sall1 than probe B-1, which contained one copy of the A/T-rich sequence (Fig. 7B).
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Discussion |
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Mutations in Townes–Brocks syndrome are often found in the region between Zn1 and Zn2 of Sall1 (Kohlhase 2000). In addition, mice expressing C-terminally-truncated Sall1 mimic the symptoms of Townes–Brocks syndrome, while Sall1-null mice do not. These data may be explained by a dominant-negative effect of the truncated Sall1 protein. Since the C-terminal zinc fingers are responsible for the heterochromatin localization, the C-terminally truncated Sall1 protein could be mislocalized from heterochromatin and may bind to other Sall family proteins through its N-terminal dimerization domain, as reported previously (Kiefer et al. 2003; Sweetman et al. 2003). Since Sall2 is not localized to heterochromatin and Sall2-deficient mice show no obvious abnormalities (Sato et al. 2003), mislocalization of Sall3 and Sall4 may be responsible for Townes–Brocks syndrome-like phenotypes in mice. Indeed, we recently showed that mice heterozygous for both Sall1 and Sall4 exhibit some of these abnormalities and that Sall1 and Sall4 form heterodimers (Sakaki-Yumoto et al. 2006). Since truncated Sall1 caused mislocalization of Sall4 from the heterochromatin, some of the Townes–Brocks syndrome-like phenotypes may result from Sall4 inhibition by truncated Sall1, functioning in a dominant-negative manner.
If this holds true in humans, the onset mechanisms of Townes–Brocks syndrome may be explained by mislocalization of SALL family members to non-heterochromatic regions via truncated SALL1. However, species differences in the pathogenesis of the disease should be taken into account. Some patients with deletion of the whole SALL1 gene on one allele still exhibit the disease (Borozdin et al. 2006). Thus SALL1 haploinsufficiency in humans can cause a mild Townes–Brocks syndrome phenotype, while Sall1 heterozygous mice show no unusual phenotypes. However, the reported phenotype is milder than that of the classical mutations, suggesting that truncated SALL1 resulting from the classical mutations may inhibit other SALL family members, thereby leading to more severe phenotypes. Regardless of the initial genetic event, the phenotypes should be caused by impairment of SALL functions, and humans may simply be more sensitive to dosage reduction of SALL1 than mice. Therefore, it remains an open question whether SALL functions are relevant to its localization to heterochromatin.
Although we have revealed the mechanism of the heterochromatin localization of Sall1, the functions of Sall1 at this nuclear region still remain unknown. Sall1 may be involved in the formation or maintenance of heterochromatin. Sall1 was reported to associate with HDAC and several components of the NuRD chromatin remodeling complex (MTA1, MTA2 and RbAp46/48) (Kiefer et al. 2002; Lauberth & Rauchman 2006). By binding to the major satellite DNA and recruiting these remodeling factors to heterochromatin, Sall1 could participate in the formation or maintenance of chemical modifications of histones or DNA in heterochromatin. Alternatively, Sall1 may function as a transcriptional regulator similar to Ikaros, another C2H2 zinc finger protein that also localizes to heterochromatin and associates with major satellite DNA. Sall1, as well as Ikaros, may regulate gene expression by repositioning target gene loci to heterochromatin (Brown et al. 1997; Cobb et al. 2000; Koipally et al. 2002). To test this hypothesis, it is necessary to identify the direct downstream targets and demonstrate the association of these loci with Sall1. Recently, it was reported that Sall4, in cooperation with Tbx5, up-regulates Fgf10 in developing forelimbs and both up-regulates Gja5 and down-regulates Nppa in the developing heart (Koshiba-Takeuchi et al. 2006). In zebrafish, sall1a, the ortholog of Sall1, functions with sall4 and regulates fgfr2 and fgf10 expressions in pectoral fin development (Harvey & Logan 2006). Thus it is necessary to test whether the gene loci of Fgf10 and other potential Sall1 targets in mammals are associated with heterochromatin. Finally, Sall1 may bind to A/T-rich sequences in the promoter region of target genes that do not associate with heterochromatin. Although our data suggest that this is less likely, it remains possible that a small amount of Sall1 protein exists in non-heterochromatic regions and regulates gene expression as a transcription factor. In any case, it is necessary to identify the direct target genes of Sall1 and the molecules it associates with. Taken together, our data have revealed the heterochromatin localization mechanism of Sall1, and further elucidation of Sall1 functions at this site would lead to better understanding of organ development and the mechanisms of Townes–Brocks syndrome.
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Experimental procedures |
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A Sall1-GFP expression vector (pCAGEN-Sall1-GFP) and N-terminal HA-tagged Sall1 expression vector (pCAGEN-HA-Sall1) were produced as described previously (Sato et al. 2004) from pCAGEN, a mammalian expression vector driven by the CAG promoter (Niwa et al. 1991). Sall1 zinc finger mutants were produced by replacing each of the zinc finger clusters of pCAGEN-Sall1-GFP with mutated fragments. Site-directed mutagenesis was performed by PCR using a Sall1 cDNA as a template and primers designed to alter the cysteine residues of each zinc finger to glycine, producing Zn1-5mut. In -helix Zn2-5 mut, -helix amino acids were changed to the Sall2 form. All PCR reactions were carried out using Pfu DNA polymerase (Promega). Each zinc finger mutant fragment amplified by PCR was cloned into pCR-Blunt II-TOPO (Invitrogen) and verified by sequencing. The Zn1 point mutant fragment was excised by EcoRI digestion, the Zn2 mutant by SacI and ApaI digestion, the Zn3 mutant by ApaI and XhoI digestion, the Zn4 mutant by XhoI and SpeI digestion and the Zn5 mutant by SpeI and AgeI digestion, followed by in-frame replacement of wild-type Sall1 with each mutant fragment. pCAGEN-HA-tagged Zn2-5mut, Zn23mut and Zn45mut without GFP, which were used for EMSAs, were generated by replacing the EcoRI-SfiI fragment of pCAGEN-HA-Sall1 with fragments of pCAGEN-Zn2-5mut-GFP, pCAGEN-Zn23mut-GFP and pCAGEN-Zn45mut-GFP, respectively.
To generate a GST-Zn5 bacterial expression vector, the SpeI-NotI fragment of Sall1 cDNA (Sato et al. 2004) was inserted in-frame into the SmaI site of pGEX-4T-3 (GE Healthcare).
Cell culture and transfection
For immunocytochemistry, E14.1 ES cells were plated on 4-well Lab-TekII chamber slides (Nunc) at a density of 5 x 103 cells/well. For localization of GFP-fused Sall1 mutants, NIH3T3 cells were plated on 4-well Lab-TekII chamber slides at a density of 1 x 104 cells/well, and 1 µg of each plasmid was transiently introduced using Fugene 6 (Roche). For the nuclear extracts used in EMSAs, 3 x 106 HEK293 cells were plated in 100-mm dishes.
Analysis of protein localization in cells
At 24 h after transfection of GFP-fused mutants, NIH3T3 cells were fixed in phosphate-buffered saline (PBS) containing 2% paraformaldehyde, 0.1% Triton X-100 and 2 µg/mL DAPI at 4 °C for 20 min, and then washed for 3 x 5 min in PBS at room temperature. For immunocytochemistry, we used a monoclonal anti-Sall1 antibody (Sato et al. 2004), an anti-HP1 antibody (Catalog No. 07-346; Upstate Biotechnology) and an anti-tri-methylated H3K9 antibody (Catalog No. 07-523; Upstate Biotechnology). After fixation, the cells were blocked with 10% goat serum for 30 min at room temperature, incubated with each primary antibody diluted 1 : 100 in PBS containing 1% goat serum for 1 h at room temperature, rinsed with PBS and detected using an Alexa Fluor 488-conjugated anti-mouse IgG secondary antibody (Molecular Probes) or rhodamine-conjugated anti-rabbit IgG secondary antibody (Chemicon). The localization of the labeled proteins was detected using a confocal microscope (Radiance 2100; Bio-Rad).
Binding site selection
For binding site selection, we employed the CASTing technique as previously described (Morinaga et al. 2005) with some modifications. Single-stranded oligonucleotides containing a 20-bp random sequence flanked by a 20-bp PCR primer annealing site were synthesized (5'-GCTCTGGAACTAGTGAGTCC-N20-CGATTCTGTCGACCTCGAAG-3'). A dsDNA library was generated via extension by Taq polymerase primed with a reverse primer (5'-CTTCGAGGTCGACAGAATCG-3'). An aliquot (2 ng) of the dsDNA library was mixed with an excess of GST-Zn5 bound to glutathione-Sepharose beads (GE Healthcare) in 100 µL of CASTing binding buffer (20 mM Tris–HCl pH 8.0, 150 mM NaCl, 2 mM MgCl2, 25 µM ZnCl2, 0.2 mM EDTA, 10% glycerol, 0.1% NP-40, 1 mM dithiothreitol (DTT), 1 mg/mL bovine serum albumin (BSA), 100 µg/µL poly[dI-dC] and 1% (v/v) protease inhibitor cocktail for mammalian cell extracts (Sigma)) and incubated at 4 °C for 30 min with continual rotation. After six washes with binding buffer (without BSA or poly[dI-dC]), the DNA-protein complexes were eluted by incubating the beads with elution buffer (5 mM reduced glutathione, 100 mM Tris–HCl pH 8.0 and 150 mM NaCl). Selected oligonucleotides were recovered by phenol extraction and ethanol precipitation, and amplified by 20 cycles of PCR with a forward primer (5'-GCTCTGGAACTAGTGAGTCC-3') and the above-described reverse primer. The same procedure was repeated for three additional rounds using 15 cycles of PCR for each. The final PCR products were subcloned into pCRII-TOPO (Invitrogen) and sequenced.
EMSAs
Bacterially-expressed GST-Zn5 was purified from bacterial lysates by binding to glutathione-Sepharose beads, followed by competitive elution with reduced glutathione. Aliquots (200 ng) of the protein were mixed with 32P-labeled DNA probes (20 000 cpm) in 15 µL of CASTing binding buffer (without BSA), incubated at 30 °C for 30 min and then resolved by 4% polyacrylamide gel electrophoresis in 0.5 x TBE and 5% glycerol. The gels were dried and autoradiographed. To prepare 32P-labeled probes, the final CASTing PCR products and dsDNA library before CASTing were labeled by PCR using [-32P]dCTP. Individual cloned sequences (probes 1, 2, 3, 4, 4-1 and 4-2) were excised from the cloned plasmids by EcoRI digestion, labeled using the Klenow large fragment (Takara) and [-32P]dATP, and purified using MicroSpin G-25 columns (GE Healthcare).
For nuclear extract preparation, HEK293 cells were transiently transfected with 6 µg of each plasmid. After 48 h, the cells were washed with PBS, lysed with 600 µL of buffer A (10 mM HEPES–KOH pH 7.8, 10 mM KCl, 1.5 mM MgCl2, 0.05% NP-40, 0.5 mM DTT and 5% (v/v) protease inhibitor cocktail for mammalian cell extracts) for 10 min on ice and centrifuged at 2 300 g for 1 min. The pellet was suspended in 300 µL of buffer B (20 mM HEPES–KOH pH 7.8, 500 mM NaCl, 1.5 mM MgCl2, 0.5 mM DTT and 5% v/v protease inhibitor cocktail for mammalian cell extracts), rotated at 4 °C for 30 min and centrifuged at 20 000 g for 30 min. The supernatant was diluted with an equal volume of buffer B containing 50% glycerol. The relative amount of each Sall1 protein was determined by Western blot analysis using the anti-Sall1 monoclonal antibody. The Sall1 protein level was normalized by dilution, with reference to nuclear extracts from cells transfected with the empty pCAG-EGFP vector.
The major satellite DNA probes were amplified by PCR using Pfu DNA polymerase, genomic DNA extracted from C57BL/6 mice and the following primers: Major satellite: forward, 5'-GGAATTCGGACCTGGAAATAGGCG-3' and reverse, 5'-GGAATTCTTCAGTGTGCATTTCTCATTTTTCACG-3'; Major A:
forward, 5'-GAATTCGGACCTGGAAATAGGCGAGAAAAC-3' and
reverse, 5'-AGTTTTCCTCGCCATATTTCACGTCCTAAA -3'; Major B:
forward, 5'-GAGGAAAACTGAAAAAGGTGGAAAATTTAG-3' and
reverse, 5'-TCTCATTTTCCATAATTATTCAGTTTTCTT-3'; and Major C:
forward, 5'-GAAAATGAGAAACATCCACTTGAAGACTTG-3' and
reverse, 5'-GAATTCTTCAGTGTGCATTTCTCATTT-3'.
Each fragment amplified by PCR was cloned into pCR-Blunt II-TOPO. Major B mutant fragments were derived from synthetic ds oligonucleotides and also cloned into pCR-Blunt II-TOPO.
For EMSAs, 3-µL aliquots of the nuclear extracts were diluted with 12 µL of EMSA reaction buffer (50 mM HEPES–KOH pH 7.8, 1.5 mM MgCl2, 0.5 mM DTT, 0.05 mM ZnCl2, 1% (v/v) protease inhibitor cocktail for mammalian cell extracts, 20% (v/v) glycerol, radioisotope-labeled probe (5 x 104 cpm)), incubated at 30 °C for 30 min, and then separated by polyacrylamide gel electrophoresis in 4% gels containing 15% glycerol with 0.5 x TBE as the electrophoresis buffer.
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Acknowledgements |
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Footnotes |
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aThese two authors contributed equally to this work.
* Correspondence: E-mail: ryuichi{at}kaiju.medic.kumamoto-u.ac.jp |
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Received: 26 April 2006
Accepted: 30 October 2006
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3'). The positions of the point mutations are shown in red. (B) EMSAs using Major B deletion mutants. (C) EMSAs showing the affinity attenuation for B-3mt (lanes 1–6). Excess amounts of non-labeled B-3 and B-3mt were added to show the binding specificity (lanes 7 and 8, respectively).