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. Author manuscript; available in PMC: 2011 Sep 10.
Published in final edited form as: Mol Cell. 2010 Sep 10;39(5):711–723. doi: 10.1016/j.molcel.2010.08.012

The Chromodomains of the Chd1 Chromatin Remodeler Regulate DNA Access to the ATPase Motor

Glenn Hauk 1,*, Jeffrey N McKnight 1,*, Ilana M Nodelman 1, Gregory D Bowman 1,1
PMCID: PMC2950701  NIHMSID: NIHMS234390  PMID: 20832723

Summary

Chromatin remodelers are ATP-driven machines that assemble, slide, and remove nucleosomes from DNA, but how the ATPase motors of remodelers are regulated is poorly understood. Here we show that the double chromodomain unit of the Chd1 remodeler blocks DNA binding and activation of the ATPase motor in the absence of nucleosome substrates. The Chd1 crystal structure reveals that an acidic helix joining the chromodomains can pack against a DNA-binding surface of the ATPase motor. Disruption of the chromodomain-ATPase interface prevents discrimination between nucleosomes and naked DNA and reduces the reliance on the histone H4 tail for nucleosome sliding. We propose that the chromodomains allow Chd1 to distinguish between nucleosomes and naked DNA by physically gating access to the ATPase motor, and we hypothesize that related ATPase motors may employ a similar strategy to discriminate among DNA-containing substrates.

Introduction

Chromatin, the physical packaging of eukaryotic chromosomes, plays a central role in regulating patterns of gene silencing and expression across the genome. A major component of chromatin regulation relies upon the reduced accessibility of DNA that is wrapped up into nucleosomes, the basic units of chromatin packaging. Nucleosomes can be assembled, removed, and shifted along DNA, and these reorganizations of nucleosome structure, known as chromatin remodeling, alter DNA accessibility required for basic cellular processes such as DNA replication, recombination, repair, and gene transcription (Clapier and Cairns, 2009).

Chromatin remodeling is driven by Swi2/Snf2 ATPase motors, classified as superfamily II (SF2) helicase-like proteins, which consist of two RecA-like ATPase lobes (Singleton et al., 2007). To prevent wide-scale unwanted disruption of duplex DNA and protein-DNA complexes in the cell, helicase-like motors are often regulated by auxiliary domains (Singleton et al., 2007). Despite the considerable information gained from structure determination and analysis of independent remodeler domains, little is known regarding how the nucleosomal substrate is recognized or how the ATPase motors of chromatin remodelers are regulated. Chromatin remodeler ATPases translocate on nucleosomal DNA at an internal position approximately 20 basepairs from the dyad known as superhelical location 2 or SHL2 (Saha et al., 2005; Schwanbeck et al., 2004; Zofall et al., 2006). One nucleosomal element that has been shown to be important for Chd1 and Iswi-type remodelers is the N-terminus of histone H4, which extends from the nucleosome at SHL2 and is thought to be an allosteric activator (Clapier et al., 2001; Hamiche et al., 2001; Ferreira et al., 2007; Gangaraju et al., 2009). Footprinting studies with the Isw2 remodeler have demonstrated that protection of nucleosomal DNA at SHL2 is significantly reduced without the H4 tail, supporting the idea that the H4 tail stabilizes an active organization of the ATPase motor on the nucleosome (Dang et al., 2006).

Chd1 is a monomeric remodeler named for its three characteristic elements: an N-terminal pair of Chromodomains, a central Helicase-like ATPase motor, and a C-terminal DNA-binding domain (Delmas et al., 1993). Chd1 has been shown to assemble, slide, and space nucleosomes in vitro (Lusser et al., 2005; Stockdale et al., 2006), and to associate with several factors involved in gene transcription (Simic et al., 2003). Chd1 has also been found to interact with the histone chaperones Nap1 (Walfridsson et al., 2007) and HIRA (Konev et al., 2007), and play an important role in deposition of the histone H3 variants CENP-A, necessary for proper centromeric maintenance (Okada et al., 2009), and H3.3, a histone variant essential for both assembly of nucleosomes onto decondensing sperm DNA and for pronuclear fusion in Drosophila (Konev et al., 2007).

Here we present the crystal structure of the chromodomain-ATPase portion of the S. cerevisiae Chd1 remodeler, revealing an autoinhibited domain organization. The double chromodomains contact both lobes of the ATPase motor in a manner that blocks productive engagement of the ATPase motor with duplex DNA. We present evidence that the chromodomains can negatively regulate the ATPase motor, and allow for discrimination between nucleosomes and naked DNA. Based on the apparently simple and direct manner in which the chromodomains inhibit the ATPase motor, we expect that a similar strategy for sharpening substrate specificity is utilized by other remodeler-type ATPases.

Results

Structure Determination and Structural Overview

We crystallized a portion of the S. cerevisiae Chd1 remodeler encompassing both N-terminal chromodomains and the ATPase motor (residues 142 – 939; Figure 1). Diffraction from these crystals was notably anisotropic, with reflections extending beyond 3.2 Å in the best direction but only ~4.2 Å in the orthogonal directions. Using a two-wavelength MAD strategy with selenomethionine-labeled protein, we obtained electron density maps of excellent quality that revealed the backbone trace for a majority of the protein (see Supplemental Movie S1). Due to the anisotropic diffraction, these maps have a low-resolution quality that precludes the visualization of most individual side chains, and we consider the information content of these density maps to be equivalent to that of a 3.7 Å resolution structure (Table I).

Figure 1. Overview of the S. cerevisiae Chd1 remodeler structure.

Figure 1

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(A) Schematic of Chd1 domain organization and residue boundaries for the crystallization construct (S. cerevisiae numbering).

(B) Two views of the remodeler domain organization. The left view shows how the two ATPase lobes are flanked by the double chromodomains (yellow) and the extended C-terminal bridge (green). The right view emphasizes the meandering path of the C-terminal bridge from the second ATPase lobe back to the first ATPase lobe. This and all subsequent molecular images were made using PyMOL (http://www.pymol.org/).

(C) Alignment of the C-terminal bridge segment for several Chd1 and Iswi orthologs. This and other sequence alignments were produced using ClustalX (Larkin et al., 2007) and formatted with TEXshade (Beitz, 2000).

See also Figure S1 and Movie S2.

Table I.

X-ray Data Collection and Refinement Statistics

Data Collection
Spacegroup: P6122 a = b = 94.3 Å, c=450.1 Å 60.5% solvent 1 mol/ASU
Wavelengths 0.9792 Å (Se peak)  0.9611 Å (high remote)
Resolution ranges1 50.0 − 3.7 Å (3.70 – 3.83 Å)  50.0 − 3.1 Å (3.10 – 3.21 Å)
Completeness (%) 99.9 (99.8)  72.5 (5.1)
Redundancy 5.5 (5.5)  5.4 (4.3)
I/σ 20.9 (4.5)  19.0 (4.3)
Rsym (%) 6.9 (40.2)  8.0 (31.7)
Refinement Statistics
Resolution ranges 50.0 − 3.7 Å  50.0 – 3.1 Å
Unique reflections 12949  15938
Reflections in test set 688  841
R-work2 (%) 26.14  26.66
R-free3 (%) 31.83  33.56
Number of atoms 5745
r.m.s. deviations
Bond lengths (Å) 0.015
Bond angles (°) 1.648
Ramachandran statistics4
Most favored 75.2%
Additionally allowed 22.3%
Generously allowed 2.5%
Disallowed 0.0%
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1

The statistics for two resolution ranges are given for the peak wavelength from the same anisotropically diffracting crystal

2

R-work = Σ |F(obs) – F(calc)|Σ F(obs)

3

R-free was calculated as R-work using 5% of the data not included in refinement

4

Calculated using PROCHECK (Laskowski et al., 1993)

See also Movie S1

The placement of the chromodomains and two lobes of the ATPase motor were apparent from solvent-flattened MAD-phased electron density maps, and readily allowed docking of individual domains from previously solved crystal structures. The two chromodomains are organized as previously observed for human and yeast Chd1 chromodomains that were solved without the ATPase motor (Cα rms distances of 1.01 Å and 1.31 Å for the isolated S. cerevisiae and human Chd1 chromodomains, respectively (Flanagan et al., 2005; Flanagan et al., 2007)). For the ATPase motor, the core fold of each domain individually matches the corresponding Swi2/Snf2-type lobe of the two available Rad54 crystal structures (Cα rms distances of 1.2 Å to 1.5 Å (Dürr et al., 2005; Thoma et al., 2005)), though for each protein the two ATPase lobes are in distinct arrangements. The proper placement of protein domains and tracing of the backbone was confirmed by matching the experimentally determined selenium positions to the 21 methionine residues throughout the structure. Although the anisotropic nature of the data precluded a detailed analysis of side chain interactions, the unambiguous fit of secondary structural elements to the experimentally-derived electron density maps allowed us to confidently identify the sequence elements participating in domain-domain interactions.

The overall structure has a flattened, ring-like appearance, with the double chromodomain unit laying across the central cleft of the ATPase motor and contacting both ATPase lobes (Figure 1B). The two chromodomains are connected by two helices (S. cerevisiae residues 239 to 284) that protrude from the chromodomains with a characteristic wedge-shape. We will refer to these connecting helices as the chromo-wedge. The second helix of the chromo-wedge packs against a groove on the second ATPase lobe, and the second chromodomain is seated on the first ATPase lobe. The double chromodomains are tethered to the first ATPase lobe via a 35 residue linker that is well-ordered in the crystal and packs against the first ATPase lobe (Movie S2).

On the opposite side of the ATPase motor from the double chromodomains is a ~50 residue segment that extends C-terminally from the second ATPase lobe. This segment runs along one face of the second ATPase lobe and then crosses over to pack against the first ATPase lobe (Figure 1B). Interestingly, a similar organization has been described for the bacterial Swi2/Snf2 protein RapA (Shaw et al., 2008), where the segment immediately following the ATPase motor bridges the two lobes (Figure S1). This location provides a potential for sensing and influencing domain motions of the ATPase motor. The amino acid sequence of this C-terminal bridge segment for Chd1 is also conserved in Iswi orthologs (Figure 1C), suggesting that the analogous segment of Iswi-type remodelers could similarly pack against the two ATPase lobes.

The Chd1 ATPase Motor Is in an Inactive Conformation

In the Chd1 crystal structure, the two ATPase lobes are splayed relatively far apart and thus not properly organized for efficient ATP hydrolysis. An example of a closed, tightly packed SF2 ATPase that is believed to represent a catalytically competent state is given by the structure of Vasa (Sengoku et al., 2006). In Vasa, residues from the conserved helicase motif VI on the second ATPase lobe pack directly against the phosphates of the bound ATP analog AMP-PNP, coordinate the attacking water molecule, and provide “arginine fingers” to stabilize the transition state (Sengoku et al., 2006). In Chd1, the backbone Cα atoms for these corresponding arginines on motif VI are approximately 14 and 19 Å from the nucleotide phosphates (compared with Cα-phosphate distances of 7.7 Å and 8.7 Å for Vasa), and thus are too far to make direct contact and catalyze ATP hydrolysis (Figure 2). Transformation of the Chd1 ATPase motor to the tightly packed organization observed for Vasa would require a swiveling of the second ATPase lobe by 52° to close the ATPase cleft. In the Chd1 crystal structure, the position of the chromodomains against the ATPase motor appears to be incompatible with such a closure of the ATPase cleft. Chd1 was crystallized in the presence of the ATP analog ATPγS, and we believe that the strong density in the P-loop is likely a bound ATPγS molecule (Figure S2). Thus, while the organization observed in the crystal structure appears compatible with nucleotide binding, we conclude that this opened ATPase configuration would not support efficient ATP hydrolysis and represents a catalytically inactive state that may be stabilized in part through interactions with the double chromodomain unit.

Figure 2. Comparison with the SF2 helicase Vasa reveals that the ATPase cleft of Chd1 is opened and not properly organized for ATP hydrolysis.

Figure 2

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(A) Crystal structure of the Vasa RNA helicase (Sengoku et al., 2006); PDB code 2DB3). The ATP analog AMP-PNP in the ATPase cleft is shown as gray spheres. Helicase motif VI on the second ATPase lobe is colored green, with two residues considered to serve as arginine fingers, R579 and R582, shown as sticks and magenta spheres.

(B) The ATPase motor of Chd1 (this study; PDB code 3MWY). The coloring is similar to (A), with the ATP analog ATPγS shown as gray spheres, the region corresponding to helicase motif VI (residues 798–809) colored green, and the Cα positions of two arginine residues corresponding to the Vasa arginine fingers shown as magenta spheres. Swi2/Snf2-specific inserts on the second ATPase lobe are colored gray.

(C) A schematic diagram illustrating the more opened ATPase cleft of Chd1 compared to Vasa. Transformation of the Chd1 motor to match the tightly packed organization of Vasa would require a closure of the ATPase cleft by 52°. Unlike the closed, hydrolysis competent structure observed for Vasa, this opened cleft of Chd1 does not permit the two arginines of motif VI to directly contact the phosphate tail of the bound nucleotide.

See also Figure S2.

The Chromo-wedge Occludes a DNA Binding Surface of the ATPase Motor

The interface between the double chromodomain unit and the ATPase motor is electrostatically complementary, with a highly acidic character of the chromo-wedge matching a basic surface on the second ATPase lobe (Figure 3). On the chromo-wedge, the helix that contacts the ATPase motor maintains six to ten acidic residues in a 19 residue stretch among diverse Chd1 orthologs (Figure 3D), with the highest conservation of acidic positions within the first turn of the helix (residues 265 – 268, S. cerevisiae numbering). On the ATPase motor, the positively charged surface contacted by the chromo-wedge is conserved in basic character not only within the Chd1 subfamily, but also more broadly among more distantly related DNA translocases. This conservation stems from the common use of this basic patch as a nucleic acid binding surface in both SF1 and SF2 ATPases (Singleton et al., 2007). To illustrate where the second ATPase lobe is expected to bind to DNA, we structurally aligned Chd1 with three SF2 ATPase crystal structures solved in complex with nucleic acid substrates: the NS3 helicase of hepatitis C virus (Kim et al., 1998), the archaeal Hel308 helicase (Büttner et al., 2007), and the RNA helicase Vasa (Sengoku et al., 2006). Using only the core fold of the second ATPase lobe for the superposition, this structural alignment reveals a common placement of the nucleic acid strands on the Chd1 surface. Strikingly, the three nucleic acid strands all penetrate the acidic helix of the chromo-wedge (Figure 3E), suggesting that the crystallographically observed placement of the chromodomains would interfere with DNA binding.

Figure 3. An acidic helix on the double chromodomains contacts the ATPase motor at a predicted DNA-binding site.

Figure 3

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(A, B, and C) Electrostatic surface representations of the double chromodomains (A), the ATPase motor (B), and a close-up of the contact between the helical linker of the chromodomains and the second ATPase lobe, with the Cα atoms of acidic residues shown as red spheres (C). The electrostatic surface potentials were calculated using APBS (Baker et al., 2001), and shown in the range of ±5.0 kBT/e with the negative and positive electrostatic potentials shown as red and blue surfaces, respectively.

(D) Sequence alignment of the acidic helix of the chromo-wedge that contacts the second ATPase lobe in the crystal structure.

(E) A view of the predicted DNA binding surface on the second ATPase lobe. The structural cores of three other SF2 ATPases bound to their nucleic acid substrates (Vasa:RNA, PDB code 2DB3, magenta (Sengoku et al., 2006); Hel308:DNA, PDB code 2P6R, green (Büttner et al., 2007); NS3:DNA, PDB code 2F55, blue (Kim et al., 1998)) were superimposed on the second ATPase lobe of Chd1. Only the nucleic acid substrates are shown. The surface of the second ATPase lobe that is within 5 Å of the acidic chromodomain helix is shown as an orange footprint.

The Chromodomain-ATPase Interface is Required for Discrimination Between Nucleosomes and Naked DNA

The hydrolysis cycle for SF1 and SF2 ATPases is typically coupled to binding of nucleic acid substrates. We were therefore curious as to how disruptions of the chromodomain-ATPase interface may influence ATPase activity in the presence of DNA and nucleosome substrates. We introduced substitutions at the chromodomain-ATPase interface, both on the chromo-wedge (E265, D266, E268) and the second ATPase lobe (R722, R750, R751, R772) (Figure 4A). As positive and negative controls, we altered a pair of conserved surface-exposed residues on the first chromodomain (D243, N244) that do not make contact with the ATPase motor in the crystal structure, and introduced a Walker B substitution (D513N) expected to interfere with ATPase activation. Since full-length S. cerevisiae Chd1 expressed poorly in E. coli, we used a construct starting from residue 118 to the extreme C-terminus (residue 1468) as the wildtype standard against which all constructs were compared (hereafter referred to as Chd1ΔN; Figure 4B). These N-terminal 117 residues are poorly conserved in Chd1 orthologs, and in our hands, the Chd1ΔN protein displayed a level of mononucleosome sliding and nucleosome assembly comparable to that previously reported for S. cerevisiae and Drosophila Chd1 ((Lusser et al., 2005; Stockdale et al., 2006) and data not shown).

Figure 4. The wild-type chromodomain-ATPase interface is required for substrate discrimination.

Figure 4

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(A) Schematic of the S. cerevisiae crystal structure, highlighting residues targeted for mutagenesis.

(B) Schematic representations of S. cerevisiae constructs used for biochemical analysis.

(C and D) ATPase activities in the presence of buffer alone, naked DNA, or mononucleosome substrates, measured using an NADH-coupled assay (Kiianitsa et al., 2003). A 206 bp DNA fragment containing the core 601 nucleosome positioning sequence at one end was used for the DNA alone and mononucleosome substrates. All experiments were performed three or more times and shown as means with standard errors.

See also Figure S3.

Similar to previous observations of yeast Chd1 (Tran et al., 2000), Chd1ΔN was preferentially stimulated by nucleosomes, with a rate of only 20±7 ATP•min−1 in the presence of naked DNA compared with 218±29 ATP•min−1 in the presence of nucleosome substrates (Figure 4C). Substitutions on the first chromodomain far from the chromodomain-ATPase interface (D243A/N244A) had little impact on DNA- and nucleosome-stimulated ATPase activity, and the Walker B substitution (D513N), as expected, showed no ATPase stimulation in the presence of DNA or nucleosome substrates. In contrast, when single (E265K) and triple substitutions (E265A/D266A/E268A [called “AAA”]; and E265K/D266A/E268K [called “KAK”]) on the acidic linker were introduced, naked DNA was highly stimulatory, activating the ATPase motor to 166±19, 174±15, and 178±9 ATP•min−1, respectively (Figure 4C). Differences between Chd1ΔN[wildtype] and Chd1ΔN[KAK] were not due to changes in ATP binding affinity as the concentration of ATP in these assays (2.5 mM) was well above the Km values (Figure S3). The DNA-stimulated ATPase rates were between 67–83% of the ATPase activity achieved using nucleosome substrates, suggesting that disruption of the chromodomain-ATPase interface incurred a loss in substrate discrimination.

To determine if a similar loss of discrimination between DNA and nucleosomes would be observed in the absence of the chromodomains, we removed both chromodomains using a PreScission protease cleavage strategy (see Methods). Similar to E265K, AAA, and KAK substitutions at the chromodomain-ATPase interface, removal of both chromodomains allowed naked DNA to activate the ATPase motor to a similar extent as nucleosome substrates, supporting the hypothesis that the chromodomain-ATPase interface is required for substrate discrimination. Unlike the Chd1ΔN variants, however, Chd1-Δchromo hydrolyzed ATP at a rate of 696±51 ATP•min−1, approximately three times higher than nucleosome-stimulated Chd1ΔN (Figure 4C). This higher ATPase stimulation suggested that some regulation by the chromodomains was maintained even when substrate discrimination was diminished due to substitutions at the chromodomain-ATPase interface.

As an alternative means of disrupting the chromodomain-ATPase interface, charge-reversal substitutions (Arg→Asp) were independently introduced on the second ATPase lobe, opposite the acidic helix of the chromo-wedge. Except for a marginal increase in DNA-stimulated ATPase activity for R722D, discrimination between DNA and nucleosome substrates was largely maintained, but overall ATPase activity was significantly diminished for variants with R750D/R751D and R772D substitutions (Figure 4C). Since these residues lie on the basic patch of the ATPase motor, they likely participate in several aspects of ATPase stimulation, such as DNA binding and stabilization of a closed ATPase cleft, and therefore this assay did not allow us to evaluate the extent that these residues influence the chromodomain-ATPase interface (see Supplemental Discussion).

Our biochemical analysis indicated that the Chd1 chromodomains are necessary for preventing ATPase activation by naked DNA substrates. To determine whether the C-terminal DNA binding region (DBR) was also required for preventing ATP hydrolysis by DNA, we compared how the presence and absence of the DBR affected nucleosome- and DNA-stimulated ATPase activities. A C-terminally truncated Chd1 variant lacking the DBR and retaining both chromodomains (Chd1ΔN-ΔDBR, residues 118–939) failed to show significant DNA- or nucleosome-stimulated ATPase activity (Figure 4D). This lack of stimulation is consistent with the DBR being an important element for targeting the ATPase motor to nucleosomal substrates. Interestingly, removing the chromodomains and DBR allowed the isolated ATPase motor (Chd1-Δchromo/ΔDBR, residues 342–939) to be stimulated by both DNA and nucleosomes. Thus, while the DBR is necessary for robust ATPase activation by nucleosomes, the chromodomains alone appear to be sufficient for inhibiting the Chd1 ATPase motor.

The chromodomain-ATPase interface weakens the association of DNA with the ATPase motor

The low stimulation of the Chd1 ATPase by naked DNA compared with nucleosome substrates, together with our structural analysis, suggested that the chromodomains may directly block DNA binding to the ATPase motor. To test this prediction, we monitored the association of Chd1 variants with duplex DNA by EMSA (electrophoretic mobility shift assay). Since the DBR associates with DNA on its own (Stokes and Perry, 1995) and would mask interactions between the ATPase motor and DNA, we used the crystallization construct (Chd1142–939) containing only the double chromodomains and ATPase motor. For the Chd1 protein possessing the wild-type chromodomain-ATPase interface, we were unable to detect stable interactions with DNA using native PAGE (Figures 5 and S4). In contrast, substitutions on the acidic chromo-wedge improved associations with DNA, although the strength of binding varied among the different substitutions. For a 16-mer DNA duplex, Chd1142–939[KAK] shifted DNA to a single, more slowly migrating band that we interpret as a Chd1-DNA complex, whereas Chd1142–939[E265K] and Chd1142–939[AAA] failed to alter DNA migration (Figure 5 and data not shown). DNA binding by Chd1142–939[E265K] and Chd1142–939[AAA] was observed using a longer DNA duplex, suggesting that these variants possess a lower affinity for DNA compared with Chd1142–939[KAK] (Figure S4). We interpret the higher apparent affinity of chromo-wedge variants for DNA to indicate an increased accessibility of the ATPase motor, with the binding increases roughly correlating with the severity of amino acid changes at the chromodomain-ATPase interface. An increased accessibility to the ATPase motor is consistent with the higher DNA-stimulated ATPase activity observed with chromo-wedge variants (Figure 4C). Although the chromo-wedge variants differ in apparent DNA binding affinities in the context of the chromodomain-ATPase fragment (Figure 5), the presence of the DNA-binding domain, which is important for robust ATPase stimulation (Figure 4D), may mask the differences in DNA-stimulated ATPase activity among chromo-wedge variants.

Figure 5. Disruption of the chromodomain-ATPase interface enhances binding of DNA to the ATPase motor.

Figure 5

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Comparison of DNA binding abilities of wildtype and variant Chd1142–939 proteins (lacking the DNA-binding domain) with a FAM-labeled 16 bp DNA duplex using native PAGE. Protein concentrations were 1.7, 7, 28, 110 µM, and labeled DNA was 25 nM.

See also Figure S4.

The Chd1 Chromodomains Play Both Positive and Negative Roles in Nucleosome Sliding

The data above demonstrate that the Chd1 chromodomains restrict ATP hydrolysis activity of the ATPase motor. To determine the extent to which regulation by the chromodomains impacts chromatin remodeling activity, we monitored nucleosome sliding activities of Chd1 variants. Yeast Chd1 was previously shown to generate evenly spaced nucleosomal arrays (Lusser et al., 2005) and slide mononucleosomes to the center of short DNA fragments (Stockdale et al., 2006). We generated end-positioned mononucleosomes using unmodified recombinant yeast histones and fluorescently labeled DNA fragments containing the 601 nucleosome positioning sequence (Lowary and Widom, 1998), and monitored nucleosome positioning using native gel electrophoresis. Compared to wildtype Chd1ΔN, sliding activity was slightly but noticeably increased for the Chd1 variant with the KAK substitutions at the chromodomain-ATPase interface (Figure 6A). At remodeler concentrations above 1 nM, both wildtype and KAK Chd1ΔN shifted end-positioned nucleosomes to a single, centrally positioned species within 60 minutes. However, at lower (0.1 nM) concentrations, wildtype Chd1ΔN shifted the majority of nucleosomes to an off-center location, whereas Chd1ΔN [KAK] shifted a significant proportion of nucleosomes to the central location (Figure 6A, compare lanes 2 and 7; also Figure 6B, compare lanes 5 and 9).

Figure 6. The chromodomain-ATPase interface both positively and negatively influences nucleosome sliding activity.

Figure 6

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Mononucleosome sliding assays for Chd1 proteins with wildtype and mutated chromodomain-ATPase interfacial residues. Starting with end-positioned (0-601-60) mononucleosomes, this assay reports on nucleosome centering as an up-shift of the nucleosome bands.

(A) End-positioned mononucleosomes (12.5 nM) were incubated with Chd1ΔN and Chd1-Δchromo proteins for 60 minutes, or Chd1142–939 proteins for 180 minutes and resolved by native PAGE. Protein concentrations of Chd1ΔN and Chd1-Δchromo proteins were 0.1, 1, 10, and 100 nM, and concentrations of Chd1142–939 were 0.01, 0.1, 1, and 10 µM. Data are representative of experiments performed three or more times.

(B –E) Disruption of the chromodomain-ATPase interface partially relieves requirement for the histone H4 tail. Both wildtype (B) and H4Δtail (D) recombinant S. cerevisiae histone octamers assembled on differently labeled 0-601-60 DNA fragments (25 nM each) were mixed and incubated with 5 nM Chd1 proteins for the indicated times. Quantification of nucleosome sliding shown in (B) and (D) is shown in (C) and (E), respectively. The percent shifted was calculated as the loss in intensity of the bottom band (end positioned nucleosome) relative to all nucleosome bands in the lane. Data are representative of experiments performed four or more times.

(F and G) Chd1-Δchromo is sensitive to the absence of the H4 tail. Chd1-Δchromo protein (100 nM) was incubated with wildtype and H4Δtail nucleosomes as (B) and (D) above. Quantification of nucleosome sliding (G) was performed as (C) and (E) above. The 40 minute time points show averages of two measurements, and all other time points show the averages and standard deviations of three measurements.

A similar increase in activity was observed when the KAK substitution was introduced in the construct used for crystallization, Chd1142–939. As expected, removal of the DNA-binding region severely limited nucleosome sliding activity, and higher concentrations (10 µM) and longer incubation times (180 minutes) were required to observe nucleosome sliding (Figure 6A, lanes 19–20). The KAK substitution allowed Chd1142–939 to shift a greater proportion of nucleosomes away from the initial end-position at lower remodeler concentrations (compare lanes 19 and 25), consistent with the conclusion that the positioning of the chromo-wedge against the ATPase motor has an intrinsically inhibitory effect on nucleosome sliding ability.

Since deletion of the chromodomains markedly increased ATPase activity, we tested Chd1-Δchromo to see whether nucleosome sliding activity was correspondingly increased. In contrast to Chd1ΔN[KAK], which efficiently mobilized nucleosomes, deletion of the chromodomains impaired nucleosome sliding ability of Chd1-Δchromo, which required approximately 100-fold higher remodeler concentration (100 nM) to shift the majority of nucleosomes to a central position (lanes 12 – 15). These results indicate that whereas the chromodomain-ATPase interface antagonizes nucleosome sliding, the chromodomains also play an important positive role in promoting efficient nucleosome sliding.

Inhibition by the Chromodomains Opposes the Positive Influence of the Histone H4 Tail

The ability of Chd1 to distinguish between nucleosomes and naked DNA indicates that the remodeler can recognize and be activated by certain elements of the nucleosome. We therefore wondered whether disruption of the inhibitory chromodomain-ATPase interface might bypass the need for some nucleosomal elements that are important for remodeling. One nucleosomal element that has been shown to be required for efficient nucleosome sliding by Chd1 is the N-terminal tail of histone H4 (Ferreira et al., 2007), which has also been shown to influence sliding by Iswi-type remodelers (Clapier et al., 2001; Hamiche et al., 2001; Ferreira et al., 2007; Gangaraju et al., 2009).

To see if disruption of the chromodomain-ATPase interface could compensate for lack of the H4 tail, we monitored sliding of Cy5-and FAM-labeled nucleosomes with and without residues 2–19 of histone H4 (referred to as H4Δtail), respectively, in the same remodeling reaction. Similar to the previously reported properties for yeast Chd1 (Ferreira et al., 2007), wildtype Chd1ΔN was less effective at mobilizing H4Δtail compared with wildtype nucleosomes: less than 40% of H4Δtail nucleosomes were shifted after 30 minutes, compared to nearly 60% of wildtype nucleosomes shifted within the first minute (Figure 6B,C). In contrast, the Chd1ΔN variants E265K, AAA, and KAK were much less affected by the absence of the H4 tail, mobilizing more than 40% of H4Δtail nucleosomes within 5 minutes (Figure 6D,E). Thus, the amino acid substitutions at the chromodomain-ATPase interface reduced the negative impact of deleting the H4 N-terminus.

The partial compensation provided by disrupting the chromodomain-ATPase interface indicates that for wildtype Chd1, the H4 tail counteracts the negative regulation by the Chd1 chromodomains. To determine whether the primary role of the H4 tail is to directly relieve inhibition by the chromodomains, we tested whether Chd1-Δchromo could distinguish between wildtype and H4Δtail nucleosomes (Figure 6F,G). Although the sliding activity of Chd1-Δchromo was relatively slow for wildtype nucleosomes, sliding of H4Δtail nucleosomes was consistently slower, indicating that some region of Chd1 outside the chromodomains was positively affected by the presence of the H4 tail. These data indicate that, although disruption of the inhibitory chromodomain-ATPase interface can relieve some dependence on the H4 tail, the positive influence of the H4 tail on nucleosome sliding does not solely stem from interfering with chromodomain inhibition.

Discussion

The ATPase motor is the core element of chromatin remodelers responsible for shifting DNA past the histone core, but how other remodeler domains influence ATPase activity is poorly understood. The structural and biochemical analysis presented here demonstrates that the ATPase motor of the Chd1 remodeler is negatively regulated by the Chd1 chromodomains. In the Chd1 crystal structure, the double chromodomains interact with both ATPase lobes and appear to help stabilize the ATPase motor in an inactive conformation. An acidic helix in the linker joining the two chromodomains (the chromo-wedge) contacts a DNA-binding surface on the ATPase motor, and we demonstrate that this interaction interferes with DNA binding to the ATPase motor. For Chd1, naked DNA is not the preferred substrate for activating the ATPase motor (Tran et al., 2000), and we found approximately 10-fold higher ATPase activity from nucleosome substrates compared to DNA alone. This preference for nucleosomes over naked DNA was eliminated with a double chromodomain deletion and various substitutions at the chromodomain-ATPase interface, indicating that the chromodomains bias Chd1 towards nucleosome substrates by inhibiting DNA binding and blocking ATPase activation.

The Chd1 chromodomains Regulate the ATPase Motor Through Modular Allostery

Modular allostery describes a regulatory strategy whereby an enzymatic core can be inhibited by structurally independent domains or segments (Dueber et al., 2004; Pufall and Graves, 2002). The crystallographically observed packing for an acidic helix of the Chd1 chromodomains against a DNA binding surface of the ATPase motor suggests a steric occlusion that would be expected to interfere with DNA binding (Figure 3E). Consistent with this interpretation, we found that amino acid substitutions of conserved acidic residues at the chromodomain-ATPase interface promoted DNA binding and allowed DNA to serve as a potent activator of the ATPase motor (Figures 4 and 5). Another potential strategy for regulating the ATPase motor is to interfere with proper closure of the two ATPase lobes, a mechanism termed conformational modular allostery (Dueber et al., 2004). For Chd1, the ATPase cleft is in an opened conformation that is not properly organized for ATP hydrolysis (Figure 2). The interaction of the chromodomains with both ATPase lobes suggests that chromodomains would likely stabilize this open conformation, reducing the likelihood of ATPase closure and hydrolysis. Thus, regulation of the Chd1 ATPase motor appears to have elements of both steric and conformational modular allostery: steric occlusion directly interferes with an activator (DNA) that promotes closure of the ATPase cleft and hydrolysis, and stabilization of the ATPase lobes in an opened state helps maintain the motor in a conformation not properly organized for efficient ATP hydrolysis.

Implications for Regulation of Swi2/Snf2-type ATPase Motors

Our finding that removal of the chromodomains from Chd1 allows maximal activation by naked DNA supports the idea that the core Swi2/Snf2 ATPase motor is intrinsically activated by DNA alone. The finding that Chd1, like other Swi2/Snf2 ATPases, prefers a protein-DNA substrate over naked DNA is consistent with the idea of an inhibitory element responsible for substrate-specific stimulation. Similarly to Chd1, both Rad54 and CSB (Cockayne Syndrome complementation group B) have been shown to possess N-terminal segments that negatively regulate the ATPase motor. For Rad54, maximal ATPase activity requires Rad51 in addition to DNA (Sigurdsson et al., 2002). The requirement for Rad51-based stimulation relies on an N-terminal segment preceding the ATPase motor, and deletion of this N-terminal segment allows maximal ATPase activation in the presence of naked DNA (Alexiadis et al., 2004). For CSB, although a precise protein-DNA substrate has not yet been defined, the remodeler has been shown to specifically localize to chromatin in response to UV-induced DNA damage (Lake et al., 2010). A segment N-terminal to the CSB ATPase motor is required to prevent chromatin association in the absence of damage, and deletion of this N-terminal segment increases ATPase stimulation by naked DNA several fold (Lake et al., 2010). Another class of remodelers that is likely regulated by an inhibitory segment includes the Iswi-type remodelers. Like Chd1, Iswi remodelers are preferentially activated by nucleosome substrates over naked DNA (Tsukiyama et al., 1999), although further work is needed to identify the element(s) that allow for discrimination against naked DNA. Our finding that packing of an acidic helix against a basic DNA binding surface of the ATPase motor can interfere with activation of the ATPase motor by naked DNA suggests a general inhibitory strategy that may be utilized by other Swi2/Snf2 ATPases.

A General Model for Chromodomain-based Regulation of Chd1

We propose that regulation by the chromodomains yields at least two functionally relevant states of the ATPase motor, which we term “gated” and “ungated” (Figure 7). In a gated state, chromodomain interactions prevent activation of the ATPase motor by blocking stable binding to duplex DNA, whereas in an ungated state, the ATPase motor is available to clamp down on DNA and hydrolyze ATP. The gating of the ATPase motor by the chromodomains therefore increases the specificity of the remodeler, providing a means to discriminate between nucleosome and naked DNA substrates.

Figure 7. A model for the regulation of chromatin remodeling by the Chd1 chromodomains.

Figure 7

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In this model, the chromodomains can occupy an inhibitory (gated) position that prevents activation of the ATPase motor. Interaction with a nucleosome relieves this inhibition by stabilizing the chromodomains in an ungated state that allows the ATPase motor to achieve a closed, hydrolysis competent conformation. Subsequent ATP hydrolysis by the motor promotes nucleosome sliding.

Discrimination between nucleosomes and DNA requires that some element(s) of the nucleosome stabilize the ATPase motor in an ungated state. The H4 tail is necessary for efficient sliding by both Chd1 and Iswi remodelers (Ferreira et al., 2007) and has been shown to aid positioning of the Isw2 ATPase motor on nucleosomal DNA (Dang et al., 2006). For Chd1, we found that defects in sliding H4Δtail nucleosomes could be partially compensated by disrupting the chromodomain-ATPase interface (Figure 6B–E), suggesting that the H4 tail counteracts the inhibitory nature of the chromodomains. While we cannot exclude the possibility that the H4 tail directly interacts with the chromodomains, we favor a model where the H4 tail counteracts the chromodomains in an indirect manner. Sliding assays with Chd1-Δchromo showed that wildtype nucleosomes were better substrates than H4Δtail nucleosomes (Figure 6F,G), indicating that some region of Chd1 outside the chromodomains interact with the H4 tail. Potential H4-interacting regions include the ATPase motor and C-terminal bridge element of Chd1, which share homology with Iswi remodelers. A direct stabilization of the Chd1 ATPase motor at SHL2, as shown for Isw2 (Dang et al., 2006), could be incompatible with chromodomain gating and therefore would indirectly counteract the inhibitory action of the chromodomains.

In addition to allowing Chd1 to discriminate between DNA and nucleosome substrates, the chromodomains provide a potential regulatory switch for guiding the reaction either towards recycling or dissociation of the remodeler. Disruption of the chromodomain-ATPase interface increased the extent that low concentrations of remodeler could move nucleosomes to a more central position (Figure 6A,B). Interestingly, although deletion of the chromodomains lowered the overall activity of Chd1 (requiring higher protein concentrations), Chd1-Δchromo strongly favored shifting nucleosomes to the most central position (Figure 6A,F), consistent with an ability of the chromodomains to antagonize remodeler recycling. Nucleosome sliding by Iswi-type remodelers has recently been shown to be processive, where an initial ATP-dependent engagement with nucleosomes allows preferential sliding in the presence of competing substrates (Gangaraju et al., 2009). Interestingly, processive nucleosome sliding by Iswi requires the H4 tail, revealing a link between remodeler activation and re-engagement with the nucleosome substrate (Gangaraju et al., 2009).

Based on the conserved acidic character of the chromo-wedge, we expect that regulation of the ATPase motor by chromodomain gating will be a common feature of all Chd1 orthologs. Chromodomain gating provides an opportunity for external elements to influence the remodeling reaction, and we speculate that the inhibitory mechanism described here may be coupled to recognition of particular epigenetic modifications. Consistent with previous findings showing that human but not S. cerevisiae Chd1 binds to the H3K4me2,3 mark (Flanagan et al., 2005; Flanagan et al., 2007; Sims et al., 2005), yeast Chd1ΔN did not display higher ATPase activity in the presence of DNA and H3K4me3 peptides, nor was it able to discriminate between nucleosomes containing unmodified versus K4me3-analog histone H3 in sliding assays (data not shown). Future work will be needed to determine whether chromodomain binding to H3K4me3 by other Chd1 orthologs influences activation of the ATPase motor, and to clarify the molecular details of how inhibition by the chromodomains can be relieved.

EXPERIMENTAL PROCEDURES

Protein Expression and Purification

All S. cerevisiae Chd1 constructs were TOPO-cloned into pDEST17 vectors (Invitrogen) and modified to contain a PreScission Protease cleavage site before the start of the protein. The S. cerevisiae Chd1 construct used for crystallization (residues 142–939) was expressed in BL21(DE3)(star) cells, with addition of the RIL plasmid (Strategene) to aid expression and a Trigger Factor Chaperone plasmid for improved protein solubility (a kind gift of LiChung Ma and Guy Montelione). All other Chd1 variants were expressed in the presence of the Rosetta2 plasmid (Novagen). To obtain selenomethionine-derived protein, cells were grown in minimal media supplemented with 5mg/L methionine, 50 –100 mg/L of the other 19 natural amino acids, and 50mg/L L-selenomethionine. After induction and growth at 18°C for 4–18 hr, cells were lysed by sonication and lysozyme in 500 mM NaCl, 10% glycerol, and 30 mM Tris pH 7.9, and the lysate clarified by centrifugation. Chd1 proteins were purified by Ni-affinity chromatography, followed by cleavage of the His-tag using Prescission Protease, a second passage over a HisTrap column (GE Healthcare), and ion-exchange chromatography on a Source-Q or SP-FF (GE Healthcare).

To obtain Chd1 constructs lacking the N-terminal chromodomains, we introduced an 11 residue segment encoding the Prescission Protease cleavage site (SSGLEVLFQGP) immediately following the double chromodomains, between residues 341–342 (QuikChange kit, Stratagene). These constructs were purified as above, except that the Prescission Protease treatment occurred after the ion-exchange chromatography step, and the cleaved, ATPase-containing fragment was separated from the chromodomains and uncleaved protein by Ni-affinity and further ion exchange chromatography.

Crystallization and Structure Determination

Two related crystal forms grew in 15–20% PEG 3350, 400 mM K+/Na+ tartrate, 5% xylitol, 10 mM MgCl2 and 1 mM ATPγS. One form diffracted to 3.1–4.2 Å resolution and was used for structure determination. The other form diffracted to a maximum resolution of 5–6 Å. Crystals were propagated by streak-seeding, which enabled us to selectively grow the better diffracting form, and typically harvested within five days. Cryoprotection was achieved by stepwise transfer to a final buffer containing 25% PEG 3350, 18% xylitol, 225 mM K+/Na+ tartrate, 15 mM MgCl2 and 5 mM ATPγS, and crystals were flash cooled by plunging into a propane slurry.

A two-wavelength MAD dataset at the selenium peak and high remote was processed using HKL2000 (Otwinowski and Minor, 1997). Prior to data scaling, we generated a mask to exclude data outside of an ellipsoid with a major axis of 3.1 Å resolution and minor axes of 4.2 Å resolution (oriented with the major axis coincident with the long c-axis of the crystal). An initial heavy atom solution and electron density maps were produced using the SOLVE/RESOLVE package (Terwilliger and Berendzen, 1999), with further refinement of heavy atom parameters and density modification carried out using SHARP (La Fortelle and Bricogne, 1997) and DM (Collaborative Computational Project Number 4, 1994). The backbone traces of the previously solved Chd1 chromodomains (PDB codes 2B2W and 2H1E) and individual ATPase lobes of two Rad54 structures (PDB codes 1Z63 and 1Z3I) were manually docked into the electron density and rebuilt using O (Kleywegt and Jones, 1996). The final Chd1 model spans residues 175 – 922, with six loop segments omitted due to missing density: 191 – 198, 476 – 480, 565 – 573, 636 – 645, 677 – 680, and 842 – 857. The low resolution nature of the electron density made it difficult to manually build some backbone segments with proper stereochemistry, and we utilized the Rosetta program suite (http://www.pyrosetta.org/) to generate geometrically acceptable segments that matched electron density. Refinement was carried out using the PHENIX suite (Adams et al., 2002) and REFMAC (Murshudov et al., 1997). Parameters were refined for three TLS groups that corresponded to the three rigid bodies in the structure: the double chromodomains, ATPase lobe 1, and ATPase lobe 2. Due to the limited resolution of the data, the B-factors were not refined. The structure factors have been deposited in the PDB, and the accession code for the atomic coordinates and structure factors is 3MWY.

Nucleosome Reconstitution

Recombinant S. cerevisiae histones were purified from E. coli, and octamer was reconstituted as previously described ((Luger et al., 1999) and web-protocol from the Tsukiyama Laboratory, http://labs.fhcrc.org/tsukiyama/protocols.html). Using the gradient dialysis method, mononucleosomes were reconstituted with S. cerevisiae histone octamer and fluorescently labeled, PCR-amplified 206 base pair DNA fragments containing a terminal 601 positioning sequence (Lowary and Widom, 1998).

Nucleosome Sliding Assay

Nucleosome sliding was performed similarly to previously published methods (Längst et al., 1999), with indicated amounts of Chd1 remodeler and mononucleosomes at 25°C in sliding buffer (10 mM HEPES pH 7.8, 10 mM MgCl2, 0.1 mM EDTA, 10% glycerol, 5 mM ATP, and either 100 mM KCl (for Chd1ΔN and Chd1-Δchromo) or 50 mM KCl (for Chd1142–939). Reactions were stopped with 1 µg unlabeled competitor DNA (equivalent to the 206 basepair fragment used for nucleosome reconstitution). To reveal positions of histone octamers on DNA fragments, 5% native PAGE was used to separate mononucleosomes, with the fluorescently labeled DNA detected using a Typhoon 9410 variable mode imager (GE Healthcare).

DNA Binding Assay

DNA binding was carried out by incubating 25 nM FAM-labeled 16 bp DNA (Figure 5) or 25 nM Cy5-labeled 228 bp DNA (Figure S4) and indicated amounts of wildtype or chromo-wedge variant Chd1 proteins, all lacking the DNA binding domain (residues 142–939), for 90 min at room temperature in 10 µL reactions. The buffer used for DNA binding reactions was 10 mM Tris pH 7.8, 50 mM NaCl, 3 mM MgCl2, 1 mM DTT, 5% glycerol, and 0.5 mg/mL BSA. Bound and free DNA were resolved by electrophoresis on a 6% native acrylamide gel in 0.25× TBE at 4°C for 60 min at 100V. Fluorescent signal was detected using a Typhoon 9410 imager.

ATPase Assay

ATP hydrolysis was monitored using an NADH-coupled assay as previously described (Kiianitsa et al., 2003). Briefly, reactions (100 µL) contained sliding buffer with 2.5 mM ATP, 18–100 nM Chd1, 0.4 mg/mL NADH, 2.5 mM PEP, and 5 units of PK/LDH (Sigma). When present, mononucleosomes reconstituted on a 206 basepair DNA fragment, or the same 206 basepair DNA fragment alone, were added to a final concentration of 200 – 1000 nM as indicated. For measurements of Chd1ΔN proteins shown in Figure 4C, we determined that nucleosome concentrations at or above 200 nM were saturating (data not shown). Absorbance was measured every 25 seconds for 15 minutes using a microplate reader, with the change in A340 reporting on the rate of NADH oxidation.

Supplementary Material

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Acknowledgements

We thank C. Ralston, G. Hura, A. Saxena, and staff scientists of the Advanced Light Source (beamlines 8.2.2 & 12.3.1) the National Light Source (beamlines X25 & X29) for providing beam time and assistance. We are grateful to G. Hartzog for the yeast Chd1 gene, T. Tsukiyama for yeast histone expression plasmids, L. Ma and G. Montelione (Rutgers University) for chaperone co-expression plasmids, and J. Gray and S. Chaudhury for advice and support with the Rosetta modeling suite. We thank G. Hartzog and our colleagues in the Biology and Biophysics Departments for discussions and critical readings of the manuscript. This work is supported by NIH/NIGMS grant R01 GM084129.

Footnotes

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