Spatial Contacts and Nucleosome Step Movements Induced by the ...

THE JOURNAL

OF

BIOLOGICAL CHEMISTRY

Vol. 279, No. 38, Issue of September 17, pp. 39933–39941, 2004 Printed in U.S.A.

Spatial Contacts and Nucleosome Step Movements Induced by the S NURF Chromatin Remodeling Complex*□ Received for publication, June 1, 2004, and in revised form, July 6, 2004 Published, JBC Papers in Press, July 15, 2004, DOI 10.1074/jbc.M406060200

Ralf Schwanbeck‡, Hua Xiao, and Carl Wu§ From the Laboratory of Molecular Cell Biology, Center for Cancer Research, NCI, National Institutes of Health, Bethesda, Maryland 20892-4255

The nucleosome remodeling factor NURF is a foursubunit, ISWI-containing chromatin remodeling complex that catalyzes nucleosome sliding in an ATP-dependent fashion, thereby modulating the accessibility of the DNA. To elucidate the mechanism of nucleosome sliding, we have investigated by hydroxyl radical footprinting how NURF makes initial contact with a nucleosome positioned at one end of a DNA fragment. NURF binds to two separate locations on the nucleosome: a continuous stretch of linker DNA up to the nucleosome entry site and a region asymmetrically surrounding the nucleosome dyad within the minor grooves, close to residues of the histone H4 tail that have been implicated in the activation of ISWI activity. Kinetic analysis reveals that nucleosome sliding occurs in apparent increments or steps of 10 bp. Furthermore, single nucleoside gaps as well as nicks about two helical turns before the dyad interfere with sliding, indicating that structural stress at this region assists the relative movement of DNA. These findings support a sliding model in which the position-specific tethering of NURF forces a translocating ISWI ATPase to pump a DNA distortion over the histone octamer, thereby changing the translational position of the nucleosome.

The condensation of eukaryotic DNA in chromatin has a significant influence on gene function and metabolism. In the nucleosome core particle, the fundamental unit of chromatin compaction, the wrapping of 146 or 147 bp of DNA in ⬃1.7 superhelical turns over a histone octamer, occludes about half of the DNA surface (1, 2), rendering it poorly accessible to macromolecules such as DNA binding regulatory proteins and the transcriptional machinery. To counteract the constraints imposed by chromatin architecture at the nucleosome and higher levels of organization, cells employ two major classes of multiprotein enzymes, which post-translationally modify the nucleosome core histones or catalyze the mobilization of nucleosomes in an ATP-dependent fashion. These enzyme complexes, when targeted onto chromatin, catalyze the reorganization of chromatin structure, thereby affecting major DNA* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. □ S The on-line version of this article (available at http://www.jbc.org) contains a supplemental movie. ‡ Supported by a European Molecular Biology Organization Long Term Fellowship and a National Institutes of Health Visiting Fellowship. § To whom correspondence should be addressed: Laboratory of Molecular Cell Biology, Center for Cancer Research, NCI, NIH, Bldg. 37, Rm. 6068, Bethesda, MD 20892-4255. Tel.: 301-496-3029; Fax: 301-4353697; E-mail: [email protected]. This paper is available on line at http://www.jbc.org

dependent processes including transcription, DNA repair, and recombination (3– 8). The ATP-dependent class of chromatin remodeling complexes (referred to hereafter as the SWI/SNF family) is highly conserved from fungi to mammals and can be divided phylogenetically into four subgroups on the basis of the catalytic SWI2/ SNF2-like subunit, the SWI2/SNF2, ISWI, CHD1 (Mi-2), and INO80 ATPases (3, 7). Intensive genetic and biochemical studies of these complexes in a variety of eukaryotic systems have established their widespread functions as important elements of the transcription regulatory apparatus. All complexes of the SWI/SNF family use the energy of ATP hydrolysis to catalyze the mobilization of nucleosomes, in which nucleosomal DNA or the core histones are dislocated in relation to their canonical organization. Despite this general similarity, marked differences exist between family subgroups in the nature and extent of the catalyzed movements of nucleosomal DNA and histones (3, 6, 7). The ISWI complexes of the SWI/SNF family were initially identified in Drosophila (9 –11) and later in yeast (12) and vertebrates (13–15). Three distinct Drosophila ISWI chromatin remodeling complexes have been characterized: nucleosome remodeling factor (NURF)1 (9, 16), chromatin accessibility complex (CHRAC) (10), and ATP-dependent chromatin and remodeling gactor (ACF) (11). Recently, components of NURF have also been found in the TATA box binding protein-related factor 2 (TRF2) complex that controls promoter-selective expression (17). Analysis of Drosophila mutants for ISWI and the largest subunits of NURF and ACF/CHRAC indicate that these proteins function in vivo in transcription, chromatin assembly, and chromosome organization (8, 18 –20). NURF, CHRAC, and ACF are ATP-dependent enzymes in vitro, mediating the relative mobilization of nucleosomal DNA without disruption or irretrievable displacement of the histone octamer; this type of nucleosome mobilization is referred to as “nucleosome sliding” (21, 22). In addition, ACF also displays nucleosome assembly activity in vitro (11). The nucleosome sliding activities of these complexes are critically dependent on the ISWI ATPase (10, 11, 16, 21–23) and modulated by other subunits that facilitate interactions with the nucleosome substrate and DNA-binding regulators (21, 22, 24 –26). Moreover, the ATPase and nucleosome sliding activities of ISWI complexes are strikingly dependent on specific residues of the N-terminal histone H4 tail, indicating that interaction with histone H4 is a key regulatory step (27–30). However, despite substantial knowledge of physiological functions and biochemical activities, important questions regarding structure and mechanism of action of the ISWI complexes remain outstanding. 1 The abbreviations used are: NURF, nucleosome remodeling factor; CHRAC, chromatin accessibility complex; ACF, ATP-dependent chromatin and remodeling factor; BSA, bovine serum albumin; MOPS, 4-morpholinepropanesulfonic acid; PDB, protein data bank.

39933

39934

Mechanism of Nucleosome Remodeling by NURF

NURF is a four-subunit complex that contains, in addition to the ISWI ATPase, NURF301, a large multifunctional polypeptide (24), NURF55, a WD-repeat protein also found in other chromatin-associated factors (31), and NURF38, an inorganic pyrophosphatase (32). We have previously reconstituted NURF by co-expression of individual subunits in a baculovirus expression system and shown that ISWI and NURF301 are minimally required for reconstitution of high level nucleosome sliding activity (24). Here, we report how NURF assembled with the natural complement of four subunits makes close contact with specific sites of a positioned nucleosome prior to ATP-dependent nucleosome sliding and show that nucleosome mobilization occurs in discrete increments or steps of 10 bp. These findings provide insight into the mechanism of NURF-induced nucleosome sliding. EXPERIMENTAL PROCEDURES

End-labeled DNA—For reconstitution of nucleosomes a 194-bp fragment comprising bp 34 –227 of the 601 clone (282 bp) from Lowary and Widom (33) was used as a strong nucleosome positioning sequence. The 194-bp fragment was generated by PCR amplification using the primers 5⬘-TCACACCGAGTTCATCCCTTATGT-3⬘ (top strand) and 5⬘-ACAGGATGTATATATCTGACACGT-3⬘ (bottom strand). All oligonucleotides were obtained from MWG-Biotech (Munich, Germany). For double end labeling of the DNA fragment the same primer modified at the 5⬘ end with Bodipy 530/550 (top strand) and Bodipy 493/503 (bottom strand) were used. The PCR contained 2 ng of 194-bp fragments as a template, 0.6 mM dNTPs (BD Bioscience), a 5 ␮M concentration of both primers, and 2 ␮l of Titanium TaqDNA Polymerase (BD Bioscience) in 1⫻ Titanium TaqPCR buffer (BD Bioscience) in a 100-␮l reaction volume. After an initial 2-min denaturation at 95 °C, 25 thermal cycles were performed (30s 95 °C, 30 s 58 °C, 1 min 68 °C) with an additional 5-min extension step at 68 °C. For large scale preparation 20 PCRs were run in parallel, and products were concentrated in Microcon YM-30 concentrators (Millipore) and gel-purified on a native 6% PAGE in 0.5 ⫻ TBE (Tris borate-EDTA) using the Mini-Prep Cell (Bio-Rad). Elution was tracked by a flow-through UV monitor (UV-1, Amersham Biosciences) using an HR-10 flow cell. Recombinant Histones, Octamer Assembly, and Nucleosome Reconstitution—Recombinant core histones from Drosophila melanogaster were expressed in Escherichia coli as described previously (27). Histone octamers were assembled and purified as described (34). Briefly, 90 ␮mol of each histone in SAU-1000 buffer (7 M urea, 20 mM sodium acetate, pH 5.2, 1 M NaCl, 5 mM ␤-mercaptoethanol, 1 mM EDTA) were mixed at room temperature for 30 min, and buffer was slowly dialyzed against refolding buffer (2 M NaCl, 10 mM Tris/HCl, 1 mM EDTA, 5 mM ␤-mercaptoethanol) as decribed (34). 500 –1000 ␮g of octamer were purified on a Superdex 200 PC3.2/30 gel filtration column at 4 °C at a flow rate of 40 ␮l refolding buffer using the SMART System (Amersham Biosciences). Octamer concentration was determined photometrically using 1 A280 ⫽ 2.80 mg/ml. In a typical nucleosome reconstitution reaction 80 ␮g of the doubly end-labeled 194-bp DNA were incubated in a 1:1 ratio (w/w) with the octamer in 100 ␮l of 2 M NaCl, 10 mM Tris/HCl, pH 7.5, and 100 ng/␮l BSA for 10 min at 37 °C. The reaction was diluted stepwise to 1.5, 1.0, and 0.5 M NaCl using dilution buffer (10 mM Tris/HCl, 1 mM EDTA, 100 ng/␮l BSA) and incubating each step for 10 min at 37 °C. The reconstitution reaction was dialyzed against TE (10 mM Tris/HCl, 1 mM EDTA) for 2 h at room temperature, concentrated with Microcon YM-30 concentrators, and the main N1 position gel-purified on a native 6% PAGE using the Mini-Prep Cell as described for the DNA purification. NURF—The four subunits of NURF (NURF301, ISWI, p55, and p38) were co-expressed in Sf9 cell and purified by pull-down of the FLAGtagged 301 subunit by anti-FLAG antibody beads (Sigma, M2) as described (24). For the footprint experiments the complex was left on the beads, whereas the complex was eluted by 1 mM 3⫻ FLAG peptide (Sigma) for all other experiments. Protein concentration was determined on a SYPRO ruby-stained SDS-PAGE using BSA (Pierce, catalog number 23209) as a reference. Gels were scanned (Typhoon 9410, Amersham Biosciences) and quantified using the ImageQuant software (Amersham Biosciences). Nucleosome Remodeling Assays—Remodeling assays were essentially performed as described (21). For the time course experiment 300 fmol of end-labeled nucleosome were incubated in 10 ␮l of sliding buffer (10 mM Tris/HCl, pH 7.6, 50 mM NaCl, 3 mM MgCl2, and 1 mM ␤-mer-

captoethanol) with 825 fmol of NURF, and the reaction was started by addition of ATP to a concentration of 1 mM. At the indicated time points 1 ␮l of the reaction was removed and put into 5 ␮l of stop solution (12% sucrose, 8 mM ␥-S-ATP, 80 ng/␮l poly(dA-dT)䡠poly(dA-dT)). The different reactions were resolved on a native 6% PAGE in 0.5⫻ TBE and fluorescence-scanned using the Typhoon 9410 (488 nm excitation and 526 short path wavelength). Remodeling activity of the anti-FLAG beads bound NURF was tested by incubating 2 ␮l of beads containing ⬃5 pmol of NURF with 0.3, 0.9, or 3 pmol of nucleosomes in presence (⫹) or absence (⫺) of 1 mM ATP for 30 min at 37 °C in 10 ␮l of sliding buffer. The concentrations of nucleosomes and remodeling enzyme used in other experiments are indicated in the respective method or figure legend. Mapping of Nucleosome Positions by Hydroxyl Radical Footprinting—To generate the different nucleosome positions, 8.7 pmol of the purified nucleosome (N1 position) were remodeled by 2.6 pmol of NURF for 1 h at 37 °C in 20 ␮l of sliding buffer containing 1 mM ATP. The reaction was stopped by addition of 2 ␮g of poly(dA-dT)䡠poly(dA-dT) and buffer exchanged to MoMK-50 buffer (10 mM MOPS, 2 mM MgCl2, 50 mM KCl, and 100 ng/␮l BSA (New England Biolabs, pH 7.2)) by adding 500 ␮l of the buffer, concentrating with a Microcon YM-30, and repeating the procedure an additional time. The volume was adjusted to 50 ␮l, and the remodeled nucleosomes were digested with hydroxyl radicals (35) by placing 0.5 ␮l of the following solutions to the inner wall of the reaction tube: (i) 20 mM (NH4)2Fe(II)SO4)2 and 40 mM EDTA, (ii) 100 mM sodium ascorbate, and (iii) 0.3% (v/v) H2O2. The reaction was initiated by spinning down the solutions, incubated 1 min at room temperature, and stopped by addition of 5 ␮l of 100 mM thiourea, 0.5 ␮l of 500 mM EDTA, and 7 ␮l of 60% sucrose. The reaction products were resolved on a native 6% PAGE in 0.5⫻ TBE, the region containing the nucleosomes was blotted onto a DEAE membrane (NA45, Schleicher and Schuell), and bands N1–N3 were cut, eluted by 2 M ammonium acetate, and ethanol-precipitated. The DNA was dissolved in sequencing loading buffer (80% deionized formamide, 1 mM EDTA, pH 8.0, and 2% 5⫻ ClearBand loading buffer (MTR Scientific, LLC) as a loading dye). By scanning the fluorescence of a portion of each reaction approximately the same amounts were loaded onto an 8% sequencing gel containing 7 M urea using a buffer electrolyte-gradient system (36) on an S2 sequencing system (Invitrogen). Gels were scanned within the glass plates (non-fluorescent, The Gel Co., model number GGO20L) with a Typhoon 9410 (Amersham Biosciences) using the following settings for excitation and emission wavelength: 488/526 nm short path for detection of the bottom strand and 532/555 ⫾ 15 nm for the top strand. Standards were either generated using the Thermo Sequenase Dye Primer Manual Cycle Sequencing Kit (United States Biochemical Corp.) or with the ABI Prism dRhodamine Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems). In the latter case the bands appearing in the 532/555 nm channel represent a G⫹A marker. NURF Footprint on Nucleosomal DNA—To capture the ternary complexes of NURF-octamer/DNA, the nucleosomes were pulled down as outlined in Fig. 2. Three ␮l of anti-FLAG beads containing ⬃7.5 pmol of NURF were incubated with 2 ␮g/␮l BSA (New England Biolabs) for 2 h at 4 °C in MoMK-80 buffer (same as MoMK-50, but with 80 mM KCl) on a rotator and washed three times with 100 ␮l of MoMK-80. Beads were resuspended in 50 ␮l of MoMK-80, and 5.4 pmol of fluorescently endlabeled purified N1 nucleosome was added and incubated at room temperature for 10 min. Under the described conditions ⬃50% of the nucleosomes are bound by NURF. Both the bound and unbound fractions were digested with hydroxyl radicals within one reaction tube as described under “Mapping of the Nucleosomes by Hydroxyl Radical Footprinting.” The reactions were stopped by addition of 5 ␮l of thiourea and 0.5 ␮l of ETDA, and bound and free fractions were separated using the beads. The bound fraction was eluted from the beads by incubation with 2 M ammonium acetate for 1 h on a rotator at room temperature, and all fractions were ethanol-precipitated using 15 ␮g of linear acrylamide per sample as a carrier. Samples were applied to a sequencing gel and scanned as described above. The quantitative analysis of the footprints was essentially carried out as described previously (37, 38) with minor changes using ImageQuant 5.2 (Amersham Biosciences). Briefly, fluorescence intensities along the lanes were scanned (lines, 21-pixel width) and imported into the ALIGN software (gift from Dr. T. Heyduk, St. Louis, MO; available on request) to correct distortions of the gel. Aligned intensity plots were imported to EXCEL (Microsoft), the background was subtracted using intensity scans of the inter-lanes, and gel-loading efficiencies and extent of cleavage were normalized. Using the standards the relative mobilities of the bands were transformed into base pair positions. For the difference plots areas under the peaks representing one base pair were integrated and positions bound by


39935

FIG. 1. NURF slides the nucleosome in 10-bp steps. A and B, kinetics of the nucleosome sliding. 300 fmol of positioned nucleosome (N1) were incubated for different reaction times as indicated with 825 fmol of NURF and 1 mM ATP at 24 °C and analyzed on a native PAGE, and fluorescence was scanned. Nucleosome positions are represented by gray ovals on the left side of the gel. The asterisk denotes a minor position that likely represents further movement from the N3 position upon prolonged incubation. B, quantification and plot of band intensities of A. C and D, mapping of nucleosome positions by hydroxyl radical footprinting. 8.7 pmol of the purified positioned 194-bp nucleosome (N1) labeled with different Bodipy dyes at the top and at the bottom strand were remodeled by 2.6 pmol of NURF for 1 h at 37 °C. Remodeled nucleosomes were subjected to hydroxyl radical digestion and reaction products separated on a native PAGE. Equal amounts of DNA of the N1–N3 positions (see A) were loaded onto a sequencing gel along with a hydroxyl radical digestion of naked DNA and marker lanes. C, footprint of nucleosomes on the top strand. The deduced locations are indicated at the right side of the lane as black ovals. D, footprint of nucleosomes on the bottom strand as in C. The deduced pseudo-dyads were mapped to 121 bp for N1, 111 bp for N2, and 101 bp for N3 as indicated by the diamonds in the enlargements of C and D. NURF and free were compared using the relative cutting frequency, which is defined by (Ibound ⫺ Ifree)/Ifree, where Ibound is the integrated intensity at one base pair of the NURF-bound fraction and Ifree is the intensity at that base in the free fraction. The relative cutting frequencies at every single position of seven independent experiments were averaged, and regions of significant protection were calculated according to a Student’s t test with a significance level of p ⫽ 0.05. Missing Nucleoside and DNA Nick Interference on Sliding—For the missing nucleoside sliding interference assay 16.6 pmol of fluorescently end-labeled purified nucleosomes (N1) were digested with hydroxyl radicals in 100 ␮l of MoMK-50 buffer as described above but with 1 ␮l of each hydroxyl radical generating solution. The reaction was stopped by addition of 5 ␮l of glycerol and the buffer exchanged to sliding buffer in two rounds of dilution/concentration using the Microcon YM-30 concentrators (Millipore). Three-fourths of the reaction was subjected to a remodeling reaction with 2.6 pmol of NURF for 30 min at 37 °C in a 30-␮l reaction volume containing 1 mM ATP. The reaction was stopped by addition of 2 ␮g of poly(dA-dT)䡠poly(dA-dT) and loaded along with sucrose loading buffer onto a native 6% PAGE in 0.5⫻ TBE. One-fourth of the digestion was applied to the gel as “unremodeled” fraction. The DNA of the N1–N3 positions was isolated by electroblotting as described under “Mapping of the Nucleosomes by Hydroxyl Radical Footprinting,” and equal amounts of DNA analyzed were applied on a sequencing gel. For the DNA nick sliding interference assay, 40 ␮g of 194-bp DNA was partially digested in 1.5 ml of reaction buffer (50 mM NaCl, 25 mM HEPES, 10 mM MgCl2, 1 mM dithiothreitol, 0.1 mM EDTA, 0.1 mg/ml BSA, pH 7.6) with 2.3 units of DNase I (Worthington) at room temperature for 80 s. Nucleosomes were assembled and purified as described above. 14.4 pmol of nucleosomes were remodeled by 2.6 pmol of NURF in sliding buffer, and interference was analyzed as described for the missing nucleoside experiment. Projection of Results onto Nucleosome Structure—Regions of the DNA that were shown to be significantly protected in the footprint or interfered in the missing nucleoside interference assay were projected onto the 147-bp 1.9-Å nucleosome (PDB code: 1kx5) (39). To visualize

the entry site of the DNA a linker of 18 bp was manually fused to the nucleosome core structure. Therefore, a second PDB file for the 18-bp DNA was generated by the web-based program “model.it” (hydra.icgeb.trieste.it/⬃kristian/dna/) (40) using the option “straight B-DNA.” Both PDB files were joined in the program DeepView/Swiss-PDB Viewer (41) (us.expasy.org/spdbv/) and bases renumbered according to our 194-bp DNA fragment. Fig. 5 was generated by Protein Explorer (42) (proteinexplorer.org). A movie is available in the supplementary data. The fused PDB file and a script to load the structure as shown is available for Protein Explorer and RasMol on request. RESULTS

NURF Moves a Positioned Nucleosome within Seconds—To define the kinetics of nucleosome mobilization catalyzed by NURF in vitro, we analyzed the first detectable movement of a mononucleosome positioned at one end of a 194-bp DNA fragment. A fluorescently end-labeled 194-bp DNA that contains a strong nucleosome positioning sequence (33) abutting the right end of the fragment was reconstituted into mononucleosomes with purified histone octamers and incubated with NURF and ATP. Analysis of the repositioned nucleosome by native polyacrylamide gel electrophoresis, in which migration of an endpositioned nucleosome is fast relative to centrally positioned nucleosomes (43, 44), shows that the action of NURF moves the terminally positioned nucleosome (N1) to two major locations toward the center of the fragment (N2 and N3) (Fig. 1A). The time course indicates that the N2 position is detectable as early as 3 s after addition of ATP. The amount of the N2 species increases with time, peaking at 200 s of incubation and decreasing thereafter. Hence, the movement from the N1 to N2 position represents the first, apparent translational step taken

39936


by the mobilized nucleosome. The N3 nucleosome reaches a maximum level at a later time (500 s) concomitant with depletion of the N2 nucleosome, indicating that the first step to the N2 position is followed by a second step from the N2 to the N3 position (Fig. 1, A and B). Nucleosome Mobilization in 10-bp Steps—To measure the length of the first and second steps of nucleosome movement, we mapped the positions of the N1, N2, and N3 nucleosomes by hydroxyl radical footprinting. The nucleosomal DNA was 5⬘ end-labeled at the top and bottom strands with different fluorescent dyes (Bodipy 530/550, top; Bodipy 493/503, bottom). (For simplicity, we number the bases from left to right for both DNA strands). DNA cleavage at every 10 bases by hydroxyl radicals reflects the periodic exposure and protection of the DNA backbone to solvent when DNA is wrapped over the histone octamer in the nucleosome core particle; this periodic cleavage is reduced or absent in free linker DNA (45, 46). Thus, the cleavage pattern of the top strand shows that one border of the N1 nucleosome is located at ⬃50 bases from the left end of the DNA fragment (Fig. 1C). Given that the edge of a nucleosome is not always precisely defined by the hydroxyl radical cleavage pattern, we localized the position of the nucleosome pseudo-dyad, and thus the position of the nucleosome, by the increased helical periodicity of the DNA adjacent to the dyad (11–12 bp/turn), in comparison with the periodicity of peripheral regions (10 bp/turn) (45, 46). Accordingly, consecutive cleavage maxima spanning 22–23 bases were found to be centered at bases 122 (N1), 112 (N2), and 102 (N3) on the top strand (Fig. 1C) and bases 120 (N1), 110 (N2), and 100 (N3) on the bottom strand (Fig. 1D), mapping the nucleosome dyad to bases 121, 111, and 101 for N1, N2, and N3 nucleosomes, respectively (Fig. 1, C and D, diamonds). Thus, the linker length of the initial N1 nucleosome is 47 bp. The positional difference between N1, N2, and N3 nucleosomes, 10 bp in each case, suggests the step size of the first two movements induced on the end-positioned nucleosome by NURF. The direction of nucleosome movement is toward the linker DNA where NURF is bound (see below). Interaction of NURF with Nucleosomal DNA—To determine how NURF makes initial contact prior to nucleosome sliding, we analyzed NURF binding to the 194-bp N1 nucleosome with the use of hydroxyl radical footprinting (35, 47). Initial attempts were unsuccessful because of a requirement for glycerol, a potent scavenger of hydroxyl radicals, to stabilize NURF activity in solution. As an alternative, we immobilized NURF on agarose beads via the FLAG epitope tag of the NURF301 subunit, removed the glycerol by washing, and immediately incubated the NURF beads with the N1 nucleosome substrate in the absence of ATP (Fig. 2A). Reaction conditions were adjusted such that approximately half of the nucleosomes were bound to NURF to avoid oversaturation (Fig. 2B). We confirmed activity of the immobilized NURF by inducing sliding of the N1 nucleosome to the N2 and N3 positions under otherwise standard conditions (Fig. 2C). Hydroxyl radical footprinting of free and NURF-bound N1 nucleosome in the same incubation, followed by centrifugal separation and side-by-side analysis of cleavage patterns, allows direct identification of changes in the footprint caused by NURF binding (Fig. 2A). The hydroxyl radical cleavage patterns of the free (F) and NURF-bound (B) N1 nucleosomes show that there are substantial regions on both strands of nucleosomal DNA that are protected by NURF (Fig. 3A, blue bars; Fig. 3B, red bars). The NURF-dependent protection is highly reproducible in multiple experiments (but is less obvious than the periodic protection provided by the histone octamer). Intensity plots for both strands are given after subtraction of background, normaliza-

FIG. 2. Immobilized NURF for footprinting. A, Strategy. Fluorescently end-labeled positioned nucleosomes are incubated with NURF complex that is bound to the anti-FLAG agarose beads (␣-FLAG beads) via the FLAG-tagged NURF301 subunit. Conditions were chosen so that ⬃50% of the nucleosomes were bound and the rest was in solution (Free). Subsequent partial digestion with hydroxyl radicals generates single-strand breaks in the DNA backbone according to the solvent accessibility of the site, and bound and free fraction were analyzed on a sequencing gel. To test the activity of the anti-FLAG beads immobilized NURF a binding test (B) and a remodeling assay (C) were performed. B, three ␮l of anti-FLAG beads containing ⬃7.5 pmol of NURF were used to pull down fluorescently labeled N1 positioned nucleosome (5.4 pmol). Equal portions of bound and free nucleosomes were loaded onto a native PAGE and fluorescence was scanned. ⬃57% of the nucleosomes was bound by the immobilized NURF in this case. C, anti-FLAG beads containing ⬃5 pmol of NURF were incubated with 0.3, 0.9, or 3 pmol of nucleosomes in the presence (⫹) or absence (⫺) of ATP for 30 min at 37 °C. Approximately the same amounts of nucleosomes were separated on a native PAGE, and fluorescence was scanned.

tion, and peak alignment (Fig. 3, C and D). In addition to the protected regions, two hypersensitive sites induced by NURF binding are apparent only for the bottom strand at bases 105 and 135 (Fig. 3, B and D, asterisks). A summary of the relative cutting frequency per base pair where enhancement of and protection from cleavage is represented by positive and negative values, respectively, shows two regions on both DNA strands of the positioned nucleosome with significant protection against the hydroxyl radical probe (Fig. 3E, blue and red bars). The regions map to a long stretch of the linker DNA, up to and including the edge of the nucleosome core particle (top: bases 11–50; bottom: bases 30 –58), and the region on both sides of the nucleosome dyad (top: bases 99 –105, 107–109, 119 –128, and 131–136; bottom: bases 97–100, 107–111, 117– 123, 130 –132, and 181–182).


39937

FIG. 3. Mapping NURF binding sites on nucleosomal DNA. Anti-FLAG beads containing ⬃7.5 pmol of NURF were incubated for binding with 5.4 pmol of fluorescently end-labeled nucleosome (N1). Samples were partially digested using hydroxyl radicals, bound and free fractions were separated, and the DNA was recovered and separated on an 8% sequencing gel. Representative footprint gels of the top (A) and the bottom strand (B) are shown in duplicates (1st and 2nd). Markers show an A and T sequencing reaction (A; T) or a G⫹A reaction (G⫹A) and naked DNA digested with hydroxyl radicals. The position of the nucleosome is represented by the green ovals with the pseudo-dyad labeled by a diamond. The roughly protected areas for the top strand are shown as blue bars (A) or red bars for the bottom strand (B). The relative mobility of the aligned lanes was transformed into base pairs as outlined under “Experimental Procedures.” C and D, representative intensity versus base pair plots are shown for the top and the bottom strand, respectively. The green lines represent the free fractions for both strands. The bound fraction of the top (C) and the bottom strands (D) are shown by the blue and red line, respectively. The asterisk in B and D indicates two hypersensitive sites in the bottom strand. E, the peak areas of the free and bound fractions were integrated for every base pair and expressed as relative cutting frequency (see “Experimental Procedures”) so that negative values represent protection by NURF binding. The presented results for the top (blue line) and bottom strand (red line) are mean values from seven independent experiments. Significantly protected regions according to a Student’s t test (significance level: p ⫽ 0.05) are shown by blue bars (top strand) and red bars (bottom strand).

Local DNA Gaps and Nicks Interfere with Nucleosome Sliding—Cleavage by hydroxyl radicals nicks the DNA sugar-phosphate backbone and removes nucleosides, providing an ensemble of single-gapped molecules. In a classical “missing nucleoside assay” DNA bases that are important for binding to

a sequence-specific binding protein are revealed by the depletion of the corresponding cleaved fragments from the proteinbound and enrichment in the protein-free fractions (47). The missing nucleoside assay did not reveal any one base that was particularly important for NURF binding (data not shown), a

39938


not altogether surprising result, given that NURF mobilizes nucleosomes independent of the underlying sequence. We then used a variation of the assay to assess whether a missing nucleoside could interfere with the NURF-induced movement of the N1 nucleosome to N2 and N3 positions. The procedure involves hydroxyl radical cleavage of the N1 nucleosome before NURF-induced nucleosome sliding and native PAGE to separate the mobilized nucleosome products for analysis of DNA cleavage (Fig. 4A). Enrichment of specific fragments isolated from the N1 nucleosomes that remained in place after mobilization and depletion of the same fragments from the mobilized N2 or N3 species identifies DNA bases whose removal interferes with nucleosome sliding. As shown by sequencing gel analysis and intensity plots, fragments from only one region of the nucleosome showed substantial enrichment in N1 and depletion from N2 and N3 nucleosomes (Fig. 4). This region maps to base positions 96 –103 on both DNA strands, about two helical turns to the left of the nucleosome dyad (Fig. 4, B, C, F, and G), overlapping with the broader NURF footprint around the dyad (Fig. 3E). Additional, minor enrichments and depletions can be observed when comparing the cleavages from N1 and N2 or N3 nucleosomes. The interference on nucleosome movement by a gap in the DNA could be caused by loss of a NURF contact with the missing DNA base or by relaxation of structural stress on the DNA. To distinguish between these possibilities, we subjected the 194-bp DNA to partial DNase I cleavage, generating a nick along the DNA backbone without loss of the DNA base. Nicked DNA was assembled into the N1 nucleosome, followed by NURF-induced mobilization, native gel electrophoresis to separate N1, N2, and N3 nucleosomes, and fragment analysis on sequencing gels. Similar to the missing nucleoside results, we found that DNase I-cleaved fragments mapping to a region approximately two helical turns before nucleosome dyad (nucleotides 96 –102 on the top strand and 92 and 95–99 on the bottom strand) were enriched in the N1 nucleosome and depleted in N2 and N3 nucleosomes (Fig. 4, D, E, H, and I). Taken together, the results suggest that relaxation of structural stress caused by a nick or gap in this specific region of nucleosomal DNA (rather than loss of a specific NURF interaction) interferes with efficient nucleosome sliding.2 DISCUSSION

Knowledge of the contacts made by NURF on the nucleosome prior to the induction of nucleosome sliding is important to understanding the mechanism of ATP-dependent nucleosome mobilization. By hydroxyl radical footprinting, we identified two regions of nucleosomal DNA that are likely to be in close contact with NURF. One region maps to the linker DNA including the site of DNA entry into the nucleosome core particle (“linker-entry contacts”). In addition, a cluster of contacts map to the nucleosome dyad, extending asymmetrically 2.5 and 1.5 helical turns on the left and right sides, respectively (“neardyad contacts”). Projection of the hydroxyl radical footprints onto the 1.9-Å crystal structure of a 147-bp nucleosome (39) and manual superimposition of a stretch of linker DNA allows visualization of the sites contacted by NURF (colored in black) in the context of the spatial organization of the nucleosome (Fig. 5). The juxtaposition of “linker-entry” and “near-dyad” contacts in the threedimensional model reveals that NURF interacts primarily with the side of the nucleosome disc where the DNA enters or exits 2 In contrast to our findings with NURF the nucleosome sliding activity of the ISWI protein is reportedly increased when the nucleosomal DNA is nicked by DNase I in the linker region (50). This may be due to differences between the ISWI protein and the complete NURF complex involving additional interactions to the linker.

the histone octamer. The protection of linker DNA, as long as 40 nucleotides on the top strand and ⬃30 nucleotides on the bottom strand (the bottom strand footprint could not be further resolved for technical reasons) indicates that NURF encompasses all sides of the double helix for several helical turns in the linker-entry region. In contrast, the cluster of “dyad” contacts occur within the minor groove at sites where the DNA faces outward, with one exception (bases 107–109 of the top strand; see Fig. 5D). Our mapping of linker-entry contacts is consistent with previous studies that showed a requirement for significant amounts of linker DNA (at least ⬃40 bp) for the full binding activity of ISWI and maximal ATPase stimulation (48 –50). The cluster of hydroxyl radical footprints at and on both sides of the nucleosome dyad identifies a second major region to which NURF binds (Fig. 5, A–C, black backbone). Interestingly, contacts between ⫺1 and ⫺2.5 helical turns from the dyad indicate that NURF is spatially close to residues of the histone H4 tail as it emerges from the nucleosome core particle. Previous work has established that ISWI complexes like NURF and CHRAC/ACF require a unique interaction with the H4 tail, at residues 16KRHR19, for activation of the ISWI ATPase and catalysis of nucleosome movement (27–30) (Fig. 5, A–D, purple). Hence, the proximity of the histone H4 tail is consistent with a critical function for the second region of NURF contacts. The measurement of a 10-bp length for the first and second steps of NURF-induced nucleosome movement is of special interest. In models of nucleosome sliding, the ATP-driven propagation or diffusion of a local DNA twist or bulge over the histone octamer surface, initiating from the DNA entry or exit positions, causes the octamer to be relocated relative to the DNA sequence (51, 52). Evidence favoring the twist or bulge migration models has been discussed extensively in recent reviews (53, 54). A simple twist diffusion model in which a local change in the DNA twist equivalent to 1 bp (34°) would, after propagation over the histone octamer, yield a corresponding nucleosome movement of 1 bp. By contrast, bulge migration requires that DNA be lifted off at each turn where the minor groove makes close contact with the histone octamer, and thus the apparent step length of nucleosome movement is dictated by the helical period of DNA. Our finding of a 10-bp step is consistent with a bulge propagation mechanism for nucleosome sliding, although the 10 bp value could be due to the special properties of the strong nucleosome positioning sequence. Note that the 10-bp step length of nucleosome movement should not be confused with the step size of DNA translocation by the ISWI motor of NURF; the latter step remains to be determined and could be any value between 1 and 10. Recent studies have shown that ISWI and other members of the SWI/SNF family display a DNA translocating activity (48, 55). We speculate that NURF, tethered near the nucleosome dyad and a linker-entry site, is triggered by the action of proximal H4 tail residues to activate the ISWI ATPase, transducing chemical energy to processively translocate the ISWI subunit in the direction of the linker DNA to which NURF is bound. Because NURF is anchored on, and therefore stationary relative to the nucleosome, the linker DNA should be forced to move into the nucleosome until the first histone-DNA contact at the entry site (superhelical location: ⫺6.5) is lost. To allow propagation of the resulting DNA wave through the nucleosome dyad, this event should be coordinated with transient release of NURF-nucleosome contacts near the dyad, perhaps assisted by structural stress accumulating locally in the same region. After passage of the DNA wave, the subsequent reformation of NURF contacts around the nucleosome dyad should prevent backward diffusion, thereby ensuring continued move-


39939

FIG. 4. Local DNA gaps and nicks interfere with nucleosome sliding. To generate a population of nucleosomes that randomly contained missing nucleosides, N1 nucleosomes (16.6 pmol) were digested by hydroxyl radicals. This population was subjected to a remodeling reaction by NURF (2.6 pmol) for 30 min at 37 °C. A, the reaction products were separated on a native PAGE and the bands corresponding N1, N2, and N3 as well as the input fraction (without NURF remodeling) were cut out of the gel, and DNA was recovered. Equal amounts were run on an 8% sequencing gel. Missing nucleoside positions that interfered with the remodeling reaction are revealed as a more intense band in the immobile fraction (N1) and as a weaker band in the remodeled fractions (N2 and N3). Gels are shown in B for the top strand and in C for the bottom strand. Input (In) and marker lanes (A and T) run in parallel. Green ovals indicate the position of N1 with diamonds as the pseudo-dyad. D and E, DNA was pretreated with DNase I to generate random nicks and was assembled into N1 nucleosomes. N1 was subjected to a remodeling reaction by NURF as outlined in A–C, and DNA was analyzed on sequencing gels (D, top strand; E, bottom strand). Quantitative scans of B, C, D, and E are given in F, G, H, and I in plots of intensity versus bases for the pairs N1/N2 and N1/N3 for top and bottom strand, respectively. For clarity the N1/N2 base line is shifted 1.2 (F), 2.5 (G), 4 (H), or 6 units (I). Regions that interfere strongly with the remodeling reactions are shown by blue (top strand) and red bars (bottom strand).

39940


FIG. 5. Projection of footprints and interference results onto nucleosome structure. A–C, NURF footprint and the missing nucleoside interference were projected onto the crystal structure of the nucleosome (PDB code: 1kx5 (39)) and an additional 18 bp of linker DNA in a front (A), back (B), and on-the-dyad (C) views. The top and bottom strands are shown in blue and red, respectively. Regions that were significantly protected in the hydroxyl radical footprint are shown as a black backbone. Positions that interfered with the remodeling reaction are shown as space-filled atoms with gold colored bases. The pseudo-dyad on the DNA is shown in green, and the different core histones are shown in light colors as indicated. The residues Lys16 to Arg19 of histone H4 are highlighted in purple. The superhelical location relative to the pseudo-dyad is labeled in every major groove in A and B. For clarity only the first turn of DNA wrapped around the octamer is shown in A and B, except for the linker DNA in B (lightened). D, enlarged view from the front side (rotated 45° to the left compared with A), with the histones as a ribbon diagram to show the hidden protection site on the inside of the nucleosome (top strand base 107–109). A movie to view the molecule as shown here is available in the supplemental data.

ment of the wave and exit from the core particle. Our findings are in general agreement with a recent study of the topography of the yeast ISW2 complex bound to a positioned nucleosome (56). In that study, yeast ISW2 was shown to bind to a broad region of the linker and the adjacent DNA of the nucleosome entry site and also to a region near the nucleosome dyad. These main features of nucleosome binding for yeast ISW2 are similar to the binding we have observed for Drosophila NURF, although differences in the extent of linker and entry site binding, and in binding near the nucleosome dyad, may reflect distinctive properties unique to the fly and

yeast ISWI complexes, which share the ISWI ATPase as the only conserved component. Acknowledgments—We thank Dr. J. Widom (Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University) for providing the strong positioning sequence and Dr. T. Heyduk (St. Louis University School of Medicine) for the ALIGN program. We also thank Dr. T. J. Ha (University of Illinois) and the members of our laboratory for suggestions and stimulating discussions. REFERENCES 1. Luger, K., Mader, A. W., Richmond, R. K., Sargent, D. F., and Richmond, T. J. (1997) Nature 389, 251–260

Mechanism of Nucleosome Remodeling by NURF 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

14. 15. 16. 17. 18. 19.

20. 21. 22. 23. 24. 25. 26. 27. 28.

Richmond, T. J., and Davey, C. A. (2003) Nature 423, 145–150 Narlikar, G. J., Fan, H. Y., and Kingston, R. E. (2002) Cell 108, 475– 487 Fischle, W., Wang, Y., and Allis, C. D. (2003) Curr. Opin. Cell Biol. 15, 172–183 Martens, J. A., and Winston, F. (2003) Curr. Opin. Genet. Dev. 13, 136 –142 Becker, P. B., and Ho¨ rz, W. (2002) Annu. Rev. Biochem. 71, 247–273 Lusser, A., and Kadonaga, J. T. (2003) Bioessays 25, 1192–1200 Corona, D. F., and Tamkun, J. W. (2004) Biochim. Biophys. Acta 1677, 113–119 Tsukiyama, T., and Wu, C. (1995) Cell 83, 1011–1020 Varga-Weisz, P. D., Wilm, M., Bonte, E., Dumas, K., Mann, M., and Becker, P. B. (1997) Nature 388, 598 – 602 Ito, T., Bulger, M., Pazin, M. J., Kobayashi, R., and Kadonaga, J. T. (1997) Cell 90, 145–155 Mellor, J., and Morillon, A. (2004) Biochim. Biophys. Acta 1677, 100 –112 Poot, R. A., Dellaire, G., Hulsmann, B. B., Grimaldi, M. A., Corona, D. F., Becker, P. B., Bickmore, W. A., and Varga-Weisz, P. D. (2000) EMBO J. 19, 3377–3387 Bochar, D. A., Savard, J., Wang, W., Lafleur, D. W., Moore, P., Coˆ te´ , J., and Shiekhattar, R. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 1038 –1043 Guschin, D., Geiman, T. M., Kikyo, N., Tremethick, D. J., Wolffe, A. P., and Wade, P. A. (2000) J. Biol. Chem. 275, 35248 –35255 Tsukiyama, T., Daniel, C., Tamkun, J., and Wu, C. (1995) Cell 83, 1021–1026 Hochheimer, A., Zhou, S., Zheng, S., Holmes, M. C., and Tjian, R. (2002) Nature 420, 439 – 445 Badenhorst, P., Voas, M., Rebay, I., and Wu, C. (2002) Genes Dev. 16, 3186 –3198 Deuring, R., Fanti, L., Armstrong, J. A., Sarte, M., Papoulas, O., Prestel, M., Daubresse, G., Verardo, M., Moseley, S. L., Berloco, M., Tsukiyama, T., Wu, C., Pimpinelli, S., and Tamkun, J. W. (2000) Mol. Cell 5, 355–365 Fyodorov, D. V., Blower, M. D., Karpen, G. H., and Kadonaga, J. T. (2004) Genes Dev. 18, 170 –183 Hamiche, A., Sandaltzopoulos, R., Gdula, D. A., and Wu, C. (1999) Cell 97, 833– 842 La¨ ngst, G., Bonte, E. J., Corona, D. F., and Becker, P. B. (1999) Cell 97, 843– 852 Corona, D. F., La¨ ngst, G., Clapier, C. R., Bonte, E. J., Ferrari, S., Tamkun, J. W., and Becker, P. B. (1999) Mol. Cell 3, 239 –245 Xiao, H., Sandaltzopoulos, R., Wang, H. M., Hamiche, A., Ranallo, R., Lee, K. M., Fu, D., and Wu, C. (2001) Mol. Cell 8, 531–543 Eberharter, A., Ferrari, S., Langst, G., Straub, T., Imhof, A., Varga-Weisz, P., Wilm, M., and Becker, P. B. (2001) EMBO J. 20, 3781–3788 Kukimoto, I., Elderkin, S., Grimaldi, M., Oelgeschlager, T., and Varga-Weisz, P. D. (2004) Mol. Cell 13, 265–277 Hamiche, A., Kang, J. G., Dennis, C., Xiao, H., and Wu, C. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 14316 –14321 Clapier, C. R., La¨ ngst, G., Corona, D. F., Becker, P. B., and Nightingale, K. P.

39941

(2001) Mol. Cell. Biol. 21, 875– 883 29. Clapier, C. R., Nightingale, K. P., and Becker, P. B. (2002) Nucleic Acids Res. 30, 649 – 655 30. Georgel, P. T., Tsukiyama, T., and Wu, C. (1997) EMBO J. 16, 4717– 4726 31. Martı´nez-Balba´s, M. A., Tsukiyama, T., Gdula, D., and Wu, C. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 132–137 32. Gdula, D. A., Sandaltzopoulos, R., Tsukiyama, T., Ossipow, V., and Wu, C. (1998) Genes Dev. 12, 3206 –3216 33. Lowary, P. T., and Widom, J. (1998) J. Mol. Biol. 276, 19 – 42 34. Luger, K., Rechsteiner, T. J., Flaus, A. J., Waye, M. M., and Richmond, T. J. (1997) J. Mol. Biol. 272, 301–311 35. Tullius, T. D., and Dombroski, B. A. (1986) Proc. Natl. Acad. Sci. U. S. A. 83, 5469 –5473 36. Sheen, J. Y., and Seed, B. (1988) BioTechniques 6, 942–944 37. Schwanbeck, R., Manfioletti, G., and Wisniewski, J. R. (2000) J. Biol. Chem. 275, 1793–1801 38. Borrmann, L., Schwanbeck, R., Heyduk, T., Seebeck, B., Rogalla, P., Bullerdiek, J., and Wisniewski, J. R. (2003) Nucleic Acids Res. 31, 6841– 6851 39. Davey, C. A., Sargent, D. F., Luger, K., Maeder, A. W., and Richmond, T. J. (2002) J. Mol. Biol. 319, 1097–1113 40. Munteanu, M. G., Vlahovicek, K., Parthasarathy, S., Simon, I., and Pongor, S. (1998) Trends Biochem. Sci 23, 341–347 41. Guex, N., and Peitsch, M. C. (1997) Electrophoresis 18, 2714 –2723 42. Martz, E. (2002) Trends Biochem. Sci. 27, 107–109 43. Duband-Goulet, I., Carot, V., Ulyanov, A. V., Douc-Rasy, S., and Prunell, A. (1992) J. Mol. Biol. 224, 981–1001 44. Meersseman, G., Pennings, S., and Bradbury, E. M. (1992) EMBO J. 11, 2951–2959 45. Hayes, J. J., Tullius, T. D., and Wolffe, A. P. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 7405–7409 46. Roberts, M. S., Fragoso, G., and Hager, G. L. (1995) Biochemistry 34, 12470 –12480 47. Dixon, W. J., Hayes, J. J., Levin, J. R., Weidner, M. F., Dombroski, B. A., and Tullius, T. D. (1991) Methods Enzymol. 208, 380 – 413 48. Whitehouse, I., Stockdale, C., Flaus, A., Szczelkun, M. D., and Owen-Hughes, T. (2003) Mol. Cell. Biol. 23, 1935–1945 49. Brehm, A., La¨ngst, G., Kehle, J., Clapier, C. R., Imhof, A., Eberharter, A., Mu¨ller, J., and Becker, P. B. (2000) EMBO J. 19, 4332– 4341 50. La¨ngst, G., and Becker, P. B. (2001) Mol. Cell 8, 1085–1092 51. Flaus, A., and Owen-Hughes, T. (2001) Curr. Opin. Genet. Dev. 11, 148 –154 52. La¨ngst, G., and Becker, P. B. (2001) J. Cell Sci. 114, 2561–2568 53. Flaus, A., and Owen-Hughes, T. (2003) Biopolymers 68, 563–578 54. La¨ngst, G., and Becker, P. B. (2004) Biochim. Biophys. Acta 1677, 58 – 63 55. Saha, A., Wittmeyer, J., and Cairns, B. R. (2002) Genes Dev. 16, 2120 –2134 56. Kagalwala, M. N., Glaus, B. J., Dang, W., Zofall, M., and Bartholomew, B. (2004) EMBO J. 23, 2092–2104