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Previous Article | Next Article 
Molecular and Cellular Biology, April 2006, p. 2550-2559, Vol. 26, No. 7
0270-7306/06/$08.00+0 doi:10.1128/MCB.26.7.2550-2559.2006
Laboratory of Molecular Immunology, National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892,1 Laboratory of Molecular Pharmacology, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 208922
Received 26 September 2005/ Returned for modification 2 December 2005/ Accepted 11 January 2006
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ABSTRACT |
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INTRODUCTION |
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Mammalian DNA is organized in a highly ordered chromatin structure, with the nucleosome as its basic repeat unit. The local chromatin architecture affects the accessibility of regulatory sequences and, thus, the expression potential of a gene. Structural changes result from posttranslational histone modifications (1, 4, 20, 21, 37, 38, 44) and from the activity of ATP-dependent remodelers, prototyped by the SWI/SNF complex (9, 12, 14, 19, 27-29, 31, 32, 40). BRG1 is the essential ATPase subunit of the mammalian SWI/SNF-like BAF complexes (10, 15, 41). Mutations of BRG1 have been identified in numerous cancerous cell lines (43), and inactivation of BRG1 in mice is lethal (2). In cell lines devoid of BRG1, reexpression activates many genes, including the CSF1 gene (25, 45).
We have shown that the TG repeat sequence and a binding site for the NFI or CAAT-box transcription factor (CTF) in the CSF1 promoter are required for activation by BRG1 (25). After activation of the CSF1 gene by BRG1, a Z-DNA structure is detected in the promoter region (25), suggesting that Z-DNA formation may be involved in the activation process. However, there is no experimental evidence regarding the mechanism of Z-DNA function in the expression of the CSF1 gene. The extensive negative supercoiling generated by transcription from the promoter can stabilize Z-DNA conformation in cells (24, 42); thus, it is not clear whether Z-DNA formation is a result of transcriptional activation or whether it plays an active role in the chromatin remodeling and transcriptional activation. In this report, we provide critical evidence that formation of Z-DNA structure plays an active role in modulating inhibitory chromatin structure. We show that Z-DNA formation facilitates productive chromatin remodeling by the BRG1-containing complexes by promoting the transition from a transient and partial remodeling to a more complete disruption of the nucleosomal structure. We propose that formation of Z-DNA structure is an important mechanism in regulating chromatin structure.
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MATERIALS AND METHODS |
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Constructs. pREP7-BRG1 and its control vector pREP7 were described previously (13). The CSF1 promoter constructs WT, noTG, and 18GC have been described previously (25). The mTA construct was generated by introducing point mutations in the two TATA boxes by use of a QuickChange kit from Stratagene.
ChIP. SW-13 cells grown in 10-cm petri dishes were cotransfected with the CSF1 reporter constructs and pREP7-BRG1 or pREP7 by use of SuperFect reagent. The cells were selected with 50 µg/ml hygromycin B for 3 days before cross-linking with formaldehyde and harvesting for chromatin immunoprecipitation (ChIP) were performed as described previously (25). The antibody for BRG1 was described previously (25); the antibody for RNA polymerase II (Pol II) was from Santa Cruz Biotech.
Detection of Z-DNA structure by ZaaFOK digestion. SW-13 cells were transfected and selected as for the ChIP experiments. MG63 cells were treated with 40 µg/ml of actinomycin D for 6 h. The cells were cross-linked with 1% formaldehyde for 5 min at room temperature, followed by washing once with 1 M glycine and once with 1x phosphate-buffered saline (PBS). Following treatment with 0.5% Triton X-100 in 1x PBS for 2 min, the cells were resuspended in 100 µl of 1 x NEB4 (20 mM Tris-acetate [pH 7.9], 10 mM magnesium acetate, 50 mM potassium acetate, 1 mM dithiothreitol) and digested with 5 ng/µl of ZaaFOK for 20 min at 37°C. The reaction was stopped by addition of 100 µl of 1 mg/ml proteinase K and 1% sodium dodecyl sulfate in 1x Tris-EDTA buffer (TE). After reverse cross-linking at 65°C for 6 h, the DNA was purified and analyzed by ligation-mediated PCR (LM-PCR) using CSF1 promoter primers as described previously (23, 25, 30).
Nucleosome mapping. SW-13 cells grown in one 10-cm petri dish were cross-linked with 1% formaldehyde for 5 min at room temperature. Following two washes with 1x PBS, the cells were incubated in 5 ml of 1%Triton X-100 in 1x PBS for 2 min. The cells were harvested into two Eppendorf tubes and were resuspended in 100 µl of 1x TE in each tube followed by digestion with 0.05 and 0.01 U of micrococcal nuclease (Sigma) for 5 min at 37°C in 1 mM CaCl2 buffer. The reaction was stopped by addition of 100 µl of 1 mg/ml proteinase K—5 mM EGTA—1% sodium dodecyl sulfate in 1x TE. After reverse cross-linking at 65°C for 6 h, 10 µg of purified DNA was resolved on a 2% agarose gel in 1x TBE buffer followed by Southern blotting using a 32P-labeled CSF1 promoter probe (KpnI/NheI fragment).
For mapping nucleosome boundaries, dinucleosome-sized DNA was isolated from an agarose gel and subjected to LM-PCR analysis. A first round of PCR to amplify regions of interest was performed followed by primer extension with a labeled primer. Locations of the labeled primers used in primer extension are denoted in the figures. The primer sequence for forward PCR primer –361F was 5'-ACGATCATAGAGCGCCAGCACTGA-3', for forward labeling primer –318F was 5'-AAGGAAAGGGTCGGTCCGCAGA, for reverse PCR primer –40R was 5'-GGCATGTGGTTTATGGGAAAT, and for reverse labeling primer –58R was 5'-AATCACCCTGGCCAGGCGCCA (see Fig. 2B). The primer sequence for PCR primer A (–203R) was 5'-CTTCCTAGTCACCCTCTGTCTT, for labeling primer A (–225R) was 5'-CTGCGCTGCACTTCCAAGCCT, for PCR primer B (–39R) was 5'-GGCATGTGGTTTATGGGAA, for labeling primer B (–58R) was 5'-AATCACCCTGGCCAGGCGCCA, for PCR primer C (+127R) was 5'-GCTCGCTCGCTGGCTGCTGC, for labeling primer C (+107R) was 5'-GGAGTGGGCGCCCCGGCCGA, for PCR primer D (–223F) was 5'-GAAGACAGAGGGTGACTAGGAAGA, for labeling primer D (–161F) was 5'-GAGGCGGGGGAAGGCGGCTGA, for PCR primer E (–349F) was 5'-GAGCGCCAGCACTGAATCAGCCTGG, and for labeling primer E (–329F) was 5'-GAGAGCGCGGAAGGAAAGGGT (see Fig. 3A).
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RESULTS |
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) of the editing enzyme ADAR1 (16, 25). The ZaaFOK nuclease domain then makes cleavages within or around the Z-DNA region to which it is bound. The generation of cleavage sites in regions of interest can be detected by LM-PCR. We assayed the induction of Z-DNA structure at the CSF1 promoter in vivo by first formaldehyde cross-linking SW-13 cells transfected with either an expression vector for BRG1 for mutBRG1 or a control expression vector. The cells were then permeabilized and treated with ZaaFOK. The LM-PCR products with CSF1 promoter-specific primers are shown in Fig. 1B. Several additional bands were generated when BRG1 was expressed compared to the results seen with cells expressing mutBRG1 or no BRG1. These new cleavage sites are located in or near the region of the CSF1 promoter containing the TG repeat sequence. These data indicate that a B-DNA-to-Z-DNA transition was induced by BRG1 in the endogenous CSF1 promoter region, and this transition required the ATP-dependent chromatin remodeling activity. Importantly, the Z-DNA-forming TG repeat sequence is required for the BRG1-mediated activation (25). The Z-DNA-forming TG repeat sequences are assembled in a positioned nucleosome. To investigate the interplay between Z-DNA and chromatin remodeling in activation of the CSF1 gene, we first mapped the nucleosomal structure of the promoter. SW-13 nuclei or purified genomic DNA was briefly treated with micrococcal nuclease (MNase) followed by complete digestion with KpnI, which recognizes a site at –734 relative to the transcription start site (+1) in the CSF1 promoter. Southern blot analysis using a probe from KpnI to the NheI sites detected a ladder of nucleosome-sized DNA from nuclei but not from purified DNA samples (Fig. 2A; compare lanes 1 and 2 with lanes 3 and 4), indicating that the promoter region is organized into several positioned nucleosomes. The nucleosome boundaries for nuc-4 and nuc-3 were estimated to be at –764 to –624 and at –564 to –420, respectively. We mapped the nucleosome boundaries of nuc-1 and nuc-2 by LM-PCR analysis of MNase-digested SW-13 nuclei (Fig. 2B). Using a primer specific to the CSF1 promoter at –318, LM-PCR yielded three bands (Fig. 2B, lane 2). The approximate size of each product is indicated in the figure. To determine the location of the protected regions, we subtracted the size of the universal linker (25 bp) to yield 90-bp, 120-bp, and 270-bp products. The protected region of approximately 150 bp corresponds to a nucleosome (nuc-1) positioned between about –200 and –50 relative to the transcription start site. A reverse primer starting at –58 revealed another protected region corresponding to a second nucleosome (nuc-2) (Fig. 2B, lane 4). We can assign a general positioning estimate for the nuc-2 boundaries at –230 and –380. Although at this resolution we cannot precisely map the nucleosome boundaries, the data strongly indicate the presence of four nucleosomal regions within the indicated ranges. Interestingly, the long stretch of Z-DNA-forming TG repeat sequence (–125 to –76) lies within nuc-1. The results are summarized in Fig. 2C.
The TG-containing nucleosome was disrupted upon activation of the CSF1 gene by BRG1. The BRG1-containing BAF complexes use energy derived from ATP hydrolysis to disrupt the extensive DNA histone interactions in a nucleosome. This disruption allows for greater accessibility of nucleosomal DNA to transcription factors. Because the DNA sequence associated with nuc-1 in the CSF promoter is required for BRG1-mediated activation of the gene, a chromatin reorganization event likely occurs in this region. We therefore examined the fate of the two promoter proximal nucleosomes after expression of BRG1. We transfected BRG1-deficient SW-13 cells with a BRG1 or control expression vector. Nuclei were isolated and subjected to MNase digestion to yield mostly dinucleosome sized material. The dinucleosome bands were purified from an agarose gel and analyzed by LM-PCR. As a control for potential bias in MNase cleavage, DNA was used. Based on the coordinates generated from the MNase mapping data, CSF1 promoter-specific primers were designed to analyze boundary regions of nuc-1 and nuc-2. The linker regions around and between these two nucleosomes were accessible to MNase both in the absence and presence of BRG1 (Fig. 3A and B; linker regions are indicated by brackets which designate the region being mapped). Activation of the CSF1 gene by BRG1 did not significantly change the digestion pattern of nuc-2, as demonstrated by the LM-PCR products generated from a primer within nuc-2 and from a primer at the 3' edge of nuc-1 (Fig. 3A, lanes 1 and 2 [Primer A] and lanes 4 and 5 [Primer B]). However, the expression of BRG1 induced several new bands in the otherwise protected region of nuc-1 that were detected by two forward primers (Fig. 3B, lanes 10 and 11 [Primer D] and lanes 13 and 14 [Primer E]) as well as by the reverse primers B and C (Fig. 3A, lanes 4 and 5 and lanes 7 and 8). Interestingly, there were more digestions in the linker regions both upstream and downstream of nuc-1 in the presence of BRG1, which is consistent with an overall more accessible chromatin structure accompanying the remodeling of nuc-1 by BRG1. These results provide strong evidence for nucleosome disruption by BRG1 at nuc-1 but not at the more distal nuc-2.
To confirm the disruption of nuc-1 by BRG1, we employed an assay based on the increased accessibility of a disrupted nucleosome to nucleases such as MNase. The accessibility to MNase was determined by measuring the relative abundance of a specific DNA sequence using quantitative PCR after digestion by the enzyme. A decrease in the total amount of DNA would be expected when the nucleosome is disrupted at the site being analyzed. We isolated nuclei from SW-13 cells transfected with either a BRG1 or control expression vector and digested with MNase to mostly mononucleosome-sized DNA, which was then isolated from an agarose gel. The amount of nuc-2 and nuc-1 DNA contained within the isolated mononucleosomal DNA was measured by quantitative PCR using specific primers. As shown in Fig. 3C (left panel), expression of BRG1 did not allow for increased accessibility of the DNA contained within nuc-2 to MNase. In contrast, a significant decrease in the amount of nuc-1 DNA contained within the mononucleosome fraction was observed in the presence of BRG1. The quantified data are presented graphically in the lower panel of Fig. 3C. Normalization of the nuc-1 DNA band to the nuc-2 DNA band indicates that there is a roughly twofold decrease of the nuc-1 DNA level (Fig. 3C, right panel). The decrease of the nuc-1 DNA level in the mononucleosome fraction could be caused by a partial disruption or complete ejection of nuc-1 upon remodeling by BRG1. Though we cannot distinguish between the two mechanisms, the data provide further evidence that the nuc-2 nucleosome in the CSF1 promoter is stable upon activation by BRG1 whereas the nuc-1 nucleosome is disrupted.
BRG1 associated with transcriptionally inactive CSF1 promoters in vivo.
Previously, we employed a transient reporter assay to determine the role of the TG repeat sequence on transcription from the CSF1 promoter (25). Using this assay, we showed that the CSF1 promoter cloned in the episomal pREP4 vector adopts Z-DNA structure when transiently cotransfected with BRG1 into SW-13 cells. The TG repeat sequence was required for activation of the promoter by BRG1 (25) (shown in Fig. 4A [noTG]). Replacement of the TG repeat sequence with a GC repeat fully supported the activation of the promoter by BRG1 (25) (shown in Fig. 4A [18GC]). In this study, we have used the approach to assay the effect of active transcription on Z-DNA formation. We generated a construct that was transcriptionally inactive but still contained the TG repeats required for Z-DNA formation by mutating the two potential TATA boxes within the CSF1 promoter (Fig. 4A [mTA]). We first asked whether the BRG1-mediated transcription from the noTG or from the mTA promoter constructs was inhibited due to decreased binding of BRG1. We used ChIP assays to test this possibility. SW-13 cells were cotransfected with the BRG1 expression vector or a control and with one of the CSF1 promoter reporter constructs. After cross-linking with formaldehyde was performed, cells were sonicated to generate mostly mono- and dinucleosome-sized DNA fragments and an anti-BRG1 antibody was used for immunoprecipitation. The relative amount of CSF1 promoter associated with BRG1 was measured by PCR. As shown in Fig. 4B, the anti-BRG1 antibody pulled down the CSF1 promoter sequence from all three constructs in a BRG1-dependent manner (lanes 2, 4, and 6), indicating that BRG1 was associated with all three promoters in vivo. However, when ChIP was performed on the same cells by use of an anti-RNA polymerase II antibody, only the wild-type promoter was associated with RNA polymerase (Fig. 4B, lane 2 [-Pol II]). Furthermore, the endogenous CSF1 promoter was also associated with BRG1 and Pol II when activated in SW-13 cells by expressing BRG1 (data not shown). The loss of transcriptional activity at the mutant promoters is not due to decreased association of BRG1. There is, however, a decreased association with Pol II, even at the "noTG" construct which has an intact TATA box. These data indicate that the Z-DNA-forming potential is necessary for BRG1 activity, which is required for activation of the CSF1 gene.
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Chromatin remodeling- but not transcription-initiated Z-DNA formation. To understand how the Z-DNA-forming sequence facilitates chromatin remodeling by BRG1, we used ZaaFOK digestion to examine the induction of Z-DNA structure in the different promoter constructs in the absence and presence of BRG1. As shown in Fig. 5A, the expression of BRG1 resulted in several new ZaaFOK cleavage sites within or near the TG repeat sequence in the wild-type promoter (compare lanes 1 and 2), indicating the formation of Z-DNA structure. Similar results were observed for the promoter containing GC repeats in place of the TG repeats (compare lanes 7 and 8). However, no significant Z-DNA structure was detected in the promoter without TG repeats even in the presence of BRG1 (lanes 3 and 4). Interestingly, in the presence of BRG1, several bands were detected within the TG repeat region in the TATA-mutated promoter, indicating that Z-DNA structure was induced even in the absence of transcription. These results strongly support the idea that the two factors required for Z-DNA formation at the CSF1 promoter are BRG-1-mediated chromatin remodeling and DNA sequence which has the potential to form Z-DNA.
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To confirm that the induction of Z-DNA structure at the CSF1 promoter does not require active transcription, we used actinomycin D to inhibit transcription in a cell line (MG63) that constitutively expresses CSF1 (Fig. 5B) (25). We treated MG63 cells with 40 µg/ml of actinomycin D for 6 h, which reduced the amount of CSF1 mRNA to less than 10% of the results seen with untreated cells (Fig. 4C). The cells were then fixed, permeabilized, and subjected to ZaaFOK treatment. LM-PCR of the resulting cleaved DNA revealed that the Z-DNA structure persisted even in the presence of actinomycin D (Fig. 4D; compare lanes 1 and 2), providing further evidence that the formation of Z-DNA structure in the CSF1 promoter does not require transcriptional activity. However, the presence of actinomycin D did inhibit two bands (as indicated by the asterisks), suggesting that, as seen with the promoter reporter constructs, the extent of Z-DNA structure was more restricted in the absence of transcription.
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DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Z-DNA may aid in chromatin remodeling by stabilizing transient nucleosome intermediates. Nucleosomal structure is energetically stabilized by extensive DNA-histone interactions (26), which can be disrupted by chromatin remodeling enzymes by use of ATP-derived energy (9, 12, 14, 19, 27-29, 31, 32, 40). Extensive studies of the mechanisms of chromatin remodeling suggest that the BAF complexes modify nucleosomal structure by transiently converting a stable nucleosome to several unstable intermediates, as illustrated in Fig. 6 (reviewed in references 31 and 19). If the unstable intermediates revert to the original, more stable state, the remodeling is unproductive. This may be the most frequent case in living cells, as the remodeling enzymes scan through the genome for specific target sites or partner proteins. However, the transient nucleosome disruption may provide an opportunity for transcription factors and/or other chromatin-modifying enzymes to access their target sites (18). The binding of transcription factors or other enzymes may capture the less stable intermediates and stabilize the remodeled state to some degree, as illustrated by step A in Fig. 6.
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Does transcription play a role in the formation of Z-DNA structure in the CSF1 promoter? The negative superhelical torsion generated behind an actively transcribing RNA polymerase has been associated with Z-DNA formation (24, 42). The results presented in Fig. 5 are consistent with a transcription-independent induction of Z-DNA structure at the CSF1 promoter. However, the ZaaFOK cleavage patterns differ between the wild-type (active) and TATA-mutated (inactive) CSF1 promoters (Fig. 5A). The cleaved bands are only detected within the TG repeat region in the TATA-mutated promoter, while the cleavage appears more extensive at the wild-type promoter since bands are detected outside of the TG repeat region (Fig. 5A). These results suggest that the Z-DNA structure induced by the chromatin remodeling activity of BRG1 in the TATA-mutated promoter is less stable or spans a smaller region than in the wild-type promoter (step B in Fig. 6). More-extensive Z-DNA as seen in the wild-type promoter may result from more negative superhelical torsion generated by transcription (step C in Fig. 6). The zigzag structure may lead to a more extensive disruption of the nucleosomal structure or ejection of the nucleosome (step C in Fig. 6).
These data are consistent with a model suggesting that induction of a localized Z-DNA structure by chromatin remodeling is followed by formation of a more extensive Z-DNA structure promoted by active transcription. The more extensive Z-DNA can inhibit reformation of the canonical nucleosome, maintaining an open chromatin structure at the promoter. Therefore, the Z-DNA structure induced by chromatin remodeling and by transcription functions at different stages of promoter activation. Chromatin remodeling-induced Z-DNA formation functions early to promote the transition from a transient and partial remodeling to a more complete disruption of the nucleosomal structure, while the transcription-induced Z-DNA formation functions late to stabilize and maintain an open chromatin structure by inhibiting reformation of the nucleosome.
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ACKNOWLEDGMENTS |
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This work was supported by intramural grants to the National Heart, Lung, and Blood Institute, National Institutes of Health.
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FOOTNOTES |
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