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  2. Structural Biology and Molecular Biophysics
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Nucleosome-CHD4 chromatin remodeler structure maps human disease mutations

  1. Lucas Farnung  Is a corresponding author
  2. Moritz Ochmann
  3. Patrick Cramer  Is a corresponding author
  1. Max Planck Institute for Biophysical Chemistry, Department of Molecular Biology, Germany
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Cite this article as: eLife 2020;9:e56178 doi: 10.7554/eLife.56178

Abstract

Chromatin remodeling plays important roles in gene regulation during development, differentiation and in disease. The chromatin remodeling enzyme CHD4 is a component of the NuRD and ChAHP complexes that are involved in gene repression. Here, we report the cryo-electron microscopy (cryo-EM) structure of Homo sapiens CHD4 engaged with a nucleosome core particle in the presence of the non-hydrolysable ATP analogue AMP-PNP at an overall resolution of 3.1 Å. The ATPase motor of CHD4 binds and distorts nucleosomal DNA at superhelical location (SHL) +2, supporting the ‘twist defect’ model of chromatin remodeling. CHD4 does not induce unwrapping of terminal DNA, in contrast to its homologue Chd1, which functions in gene activation. Our structure also maps CHD4 mutations that are associated with human cancer or the intellectual disability disorder Sifrim-Hitz-Weiss syndrome.

Introduction

In the nucleus of eukaryotic cells, DNA is compacted into chromatin. The fundamental building block of chromatin is the nucleosome, a complex of ~146 base pairs (bp) of DNA wrapped around an octamer of histone proteins. The degree of chromatin compaction influences DNA replication, transcription, and repair. Maintenance of the appropriate chromatin state requires ATP-dependent chromatin-remodeling enzymes. These ‘chromatin remodelers’ are divided into four families called CHD, SWI/SNF, ISWI, and INO80 (Clapier et al., 2017). All chromatin remodelers contain a conserved ATPase core that hydrolyses ATP to alter contacts between nucleosomal DNA and the histone octamer and to facilitate nucleosome assembly, sliding, ejection, or histone exchange.

Members of the CHD (‘chromodomain helicase DNA-binding’) family of chromatin remodelers all contain a central SNF2-like ATPase motor domain and a double chromodomain in their N-terminal region. The double chromodomain binds modified histones (Sims et al., 2005) and interacts with nucleosomal DNA to regulate ATPase activity (Nodelman et al., 2017). Recent structures of the yeast remodeler Chd1 in complex with a nucleosome uncovered the architecture of one subfamily of CHD remodelers (subfamily I) and its interactions with the nucleosome (Farnung et al., 2017; Sundaramoorthy et al., 2018). A unique feature of these structures is that Chd1 binding induces unwrapping of terminal DNA from the histone octamer surface at superhelical location (SHL) −6 and −7 (Farnung et al., 2017; Sundaramoorthy et al., 2018). However, the resolution of these studies was limited, such that atomic details were not resolved.

The human CHD family member CHD4 (Woodage et al., 1997) shows nucleosome spacing activity (Silva et al., 2016). CHD4 is also known as Mi-2 in Drosophila melanogaster (Kehle et al., 1998). CHD4, CHD3, and CHD5 form CHD subfamily II, which differs in domain architecture from subfamily I. CHD3, CHD4, and CHD5 contain two N-terminal plant homeodomain (PHD) zinc finger domains (Schindler et al., 1993), a DNA-interacting double chromodomain, and the ATPase motor. CHD4 contains an additional high mobility group (HMG) box-like domain in its N-terminal region (Silva et al., 2016) and two additional domains of unknown function that are located in the C-terminal region.

CHD4 is implicated in the repression of lineage-specific genes during differentiation (Liang et al., 2017) and is required for the establishment and maintenance of more compacted chromatin structures (Bornelöv et al., 2018). CHD4 mutations have a high incidence in some carcinomas (Kandoth et al., 2013) and in thyroid and ovarian cancers (Längst and Manelyte, 2015). Mutations in CHD4 have also been implicated in intellectual disability syndromes (Sifrim et al., 2016; Weiss et al., 2016).

CHD4 is a subunit of the multi-subunit Nucleosome Remodeling Deacetylase (NuRD) complex (Tong et al., 1998; Xue et al., 1998; Zhang et al., 1998). NuRD also contains the deacetylase HDAC1/2 and accessory subunits that serve regulatory and scaffolding roles. NuRD is implicated in gene silencing, but also gene activation (Gnanapragasam et al., 2011). It is essential for cell cycle progression (Polo et al., 2010), DNA damage response (Larsen et al., 2010; Smeenk et al., 2010), establishment of heterochromatin (Sims and Wade, 2011), and differentiation (Bornelöv et al., 2018; Burgold et al., 2019). In addition, CHD4 is part of the heterotrimeric ChAHP complex that is also involved in gene repression (Ostapcuk et al., 2018).

Thus far, structural studies of CHD4 have been limited to individual domains (Kwan et al., 2003; Mansfield et al., 2011). Here, we report the cryo-electron microscopy (cryo-EM) structure of human CHD4 bound to a nucleosome at an overall resolution of 3.1 Å. CHD4 engages the nucleosome at SHL +2 and induces a conformational change in DNA at this location in the presence of the ATP analogue adenylyl imidodiphosphate (AMP-PNP). Structural comparisons show that CHD4, in contrast to Chd1, does not induce unwrapping of terminal DNA, and this is also observed in biochemical assays. Maintenance of the integrity of the nucleosome in the presence of CHD4 is consistent with the role of CHD4 in gene repression, and in heterochromatin formation and maintenance. Finally, the detailed nucleosome-CHD4 structure enables mapping of known human disease mutations (Kovač et al., 2018; Sifrim et al., 2016; Weiss et al., 2016) and indicates how these may perturb enzyme function.

Results

Nucleosome-CHD4 complex structure

To investigate how the human chromatin remodeller CHD4 engages a nucleosome, we determined the structure of H. sapiens CHD4 bound to a Xenopus laevis nucleosome core particle in the presence of the ATP analogue AMP-PNP. We recombinantly expressed and purified full-length CHD4 and reconstituted a complex of CHD4 with a pre-assembled nucleosome core particle. The nucleosome comprised 145 base pairs (bp) of DNA, corresponding to the Widom 601 sequence (Lowary and Widom, 1998) with additional 4 and 30 bp of extranucleosomal DNA on the entry and exit side of the nucleosome, respectively. The nucleosome-CHD4 complex was purified by size exclusion chromatography (Figure 1—figure supplement 1).

To determine the structure of the nucleosome-CHD4 complex, we collected single particle cryo-EM data on a Titan Krios (FEI) microscope equipped with a K2 direct electron detector (Gatan) (Materials and methods). We obtained a cryo-EM reconstruction of the nucleosome-CHD4 complex at an overall resolution of 3.1 Å (FSC 0.143 criterion) (Figure 1—figure supplements 24, Video 1). The nucleosome was resolved at a resolution of 3.0–4.5 Å, whereas CHD4 was resolved at 3.1–5.0 Å, depending on the protein region. The register of the DNA was unambiguously determined based on distinct densities for purine and pyrimidine nucleotides around the dyad (Figure 1—figure supplement 3h). Well-defined density was also obtained for AMP-PNP and a coordinated magnesium ion in the CHD4 active site (Figure 1—figure supplement 3i). The model was locally adjusted and real-space refined, leading to very good stereochemistry (Materials and methods) (Table 1).

Table 1
Cryo-EM data collection, refinement and validation statistics.
Nucleosome-CHD4 complex
(EMD-10058)
(PDB 6RYR)
Nucleosome-CHD42 complex
(EMDB-10059)
(PDB 6RYU)
Data collection and processing
Magnification130,000130,000
Voltage (kV)300300
Electron exposure (e–/Å2)43–4543–45
Defocus range (μm)0.25–40.25–4
Pixel size (Å)1.051.05
Symmetry imposedC1C1
Initial particle images (no.)650,599650,599
Final particle images (no.)89,62340,233
Map resolution (Å)
FSC threshold
3.1
0.143
4.0
0.143
Map resolution range (Å)3.0–53.7–8.3
Refinement
Initial models used (PDB code)3LZ0, 5O9G, 2L75, 4O9I, 6Q3M3LZ0, 5O9G, 2L75, 4O9I, 6Q3M
Map sharpening B factor (Å2)−36−86
Model composition
Non-hydrogen atoms
Protein residues
Nucleotides
Ligands

17,834
1463
298
4

23,598
2180
298
8
B factors (Å2)
Protein
Nucleotide
Ligand

45.28
71.82
60.10

95.29
112.27
125.7
R.m.s. deviations
Bond lengths (Å)
Bond angles (°)

0.003
0.638

0.005
1.028
Validation
MolProbity score
Clashscore
Poor rotamers (%)

1.54
5.69
0.08

1.92
6.52
1.64
Ramachandran plot
Favored (%)
Allowed (%)
Disallowed (%)

96.50
3.50
0.0

94.16
5.84
0.0
Video 1
Cryo-EM density and structure of the nucleosome-CHD4 complex.

CHD4 architecture

The CHD4 ATPase motor binds the nucleosome at SHL +2 (Figure 1, Figure 1—figure supplement 4f). Binding at this location has also been observed for the chromatin remodelers Chd1 (Farnung et al., 2017; Sundaramoorthy et al., 2018), Snf2 (Liu et al., 2017), and Swr1 (Willhoft et al., 2018). The ATPase motor is in a closed, post-translocated state with AMP-PNP bound in the active site. The same state and a similar conformation was observed for Chd1 when bound to ADP·BeF3 (Farnung et al., 2017; Sundaramoorthy et al., 2018; Sundaramoorthy et al., 2017). The double chromodomain is located at SHL +1 and contacts the nucleosomal DNA phosphate backbone via electrostatic interactions, in a fashion similar to that observed for S. cerevisiae Chd1 (Figure 1; Farnung et al., 2017; Nodelman et al., 2017).

Figure 1 with 4 supplements see all
Structure of the nucleosome-CHD4 complex.

(a) Schematic of domain architecture of CHD4. Domain borders are indicated. (b-d) Cartoon model viewed from the top (b), dyad (c), and side (d). Histones H2A, H2B, H3, H4, tracking strand, guide strand, CHD4 PHD finger 2, double chromodomain, ATPase lobe 1, and ATPase lobe 2 are colored in yellow, red, light blue, green, blue, cyan, pink, purple, orange, and forest green, respectively. Color code used throughout. The dyad axis is indicated as a black line or a black oval circle. Magnesium and zinc ions shown as pink and grey spheres, respectively. AMP-PNP shown in stick representation.

The PHD finger 2 of CHD4 is located near SHL +0.5 and the double chromodomain. This is consistent with NMR studies that predicted binding of this PHD finger close to the dyad axis and the H3 tail (Gatchalian et al., 2017). Additionally, we observe parts of the C-terminal bridge (Hauk et al., 2010), an amino acid segment that follows the ATPase lobes. Part of the C-terminal bridge docks against ATPase lobe 2 and extends toward the first ATPase lobe (Figure 1, Figure 1—figure supplement 3j). This region was not resolved in the nucleosome-Chd1 structures but was observed in a previously published crystal structure of auto-inhibited Chd1 (Hauk et al., 2010). Taken together, CHD4 and Chd1 share a core architecture that involves the ATPase motor and the double chromodomain but differ in their peripheral subfamily-specific protein features.

CHD4 does not detach exit side nucleosomal DNA

In contrast to the nucleosome-Chd1 structure (Farnung et al., 2017), we did not observe unwrapping of nucleosomal DNA from the histone octamer on the second DNA gyre at SHL −6 and −7 (Figure 2a). To test whether this structural difference can be recapitulated biochemically in solution, we used a Förster Resonance Energy Transfer (FRET) assay to monitor putative DNA unwrapping activity by these two chromatin remodellers. The DNA 5’ ends of the nucleosome were labelled with Cy3 or Cy5 (Figure 2b). Using the doubly labeled nucleosome, FRET efficiencies were measured in the absence and presence of S. cerevisiae Chd1 (residues 1–1247) or full-length H. sapiens CHD4, and in the presence of AMP-PNP or ADP·BeF3.

Comparison with nucleosome-Chd1 structure.

(a) CHD4 (left) does not possess a DNA-binding region and does not detach DNA from the second gyre. Chd1 (right) detaches DNA from SHL −7 to −5, stabilizes the detached DNA via its DNA-binding region, and introduces a ~ 60° bend with respect to the canonical DNA position observed in the nucleosome-CHD4 structure. (b) Schematic of experimental FRET setup. (c) Fluorescence emission spectra produced after excitation at 510 nm of Cy3/Cy5 labeled nucleosome in the presence of S. cerevisiae Chd1 (residues 1–1247) or H. sapiens CHD4 and AMP-PNP or ADP·BeF3 show unwrapping of nucleosomal DNA by Chd1 but not by CHD4.

In these assays, Chd1 showed an increase in fluorescence emission of the donor and a reduction in the acceptor emission (Figure 2c). This indicated that the distance between the two DNA ends of the nucleosome increased upon Chd1 addition, and was consistent with the structurally observed DNA unwrapping of terminal DNA. In contrast, fluorescence emissions measured for the CHD4 sample did not differ from the nucleosome controls (Figure 2c), showing that CHD4 was unable to unwrap nucleosomal DNA both in the presence of AMP-PNP or ADP·BeF3.

The major difference in DNA unwrapping between these two remodelers may be due to a lack of a DNA-binding region in CHD4, when compared to Chd1. Chd1 uses its DNA-binding region to interact extensively with terminal DNA on the exit side at SHL −7, and such contacts are absent in the nucleosome-CHD4 structure (Figure 2). It is likely that other CHD family members from subfamily II such as CHD3 and CHD5, which also lack a DNA-binding region, will also not induce unwrapping of terminal DNA.

CHD4-DNA interactions

The high resolution of our nucleosome-CHD4 structure enables a detailed description of the interactions of the ATPase motor with nucleosomal DNA. CHD4 contacts the phosphate backbone of the tracking and guide strands via electrostatic interactions that are mediated by lysine and arginine residues (Figure 3). These interactions with the DNA phosphate backbone are formed by residues in the canonical ATPase motifs Ia, Ic, II, IV, IVa, V, and Va and by residues present in non-canonical motifs (e.g. Lys810) (Figure 3, Figure 3—figure supplement 1).

Figure 3 with 1 supplement see all
CHD4-DNA interactions and DNA distortion.

(a) CHD4 interacts extensively with nucleosomal DNA around SHL +2. ATPase lobe 1 and lobe 2 of CHD4 are shown. Guide and tracking strands are indicated. ATPase motifs are shown as colored spheres and labelled. (b) Schematic depiction of DNA interactions of the double chromodomain, ATPase lobe 1 and lobe 2. (c) Asn1010, Trp1148 and Arg1227 insert into the minor groove between DNA tracking and guide strand. The two conformations of the Arg1127 side chain are shown. Nucleic acids are shown as cartoons with their respective surfaces. (d) Detailed cartoon representation of DNA distortion at SHL +2. Canonical nucleosome (PDB code 3LZ0, grey), AMP-PNP bound NCP-CHD4 structure (this study, blue and cyan), and ADP bound nucleosome-Snf2 structure (PDB code 5Z3O, red and yellow) are shown. Phosphate atoms shown as spheres.

We also observe that residues Asn1010, Arg1127, and Trp1148 insert into the DNA minor groove over a stretch of seven base pairs (Figure 3c). Asn1010 is not part of a canonical ATPase motif and inserts into the DNA minor groove around SHL +2.5. Arg1127 (motif V) is universally conserved in all CHD chromatin remodelers and inserts into the DNA minor groove at SHL +2. Our density is consistent with two alternative conformations of the Arg1127 side chain, with the guanidinium head group pointing either toward the tracking or the guide strand of DNA. Trp1148 is located in motif Va, inserts into the minor groove near the guide strand, and plays a critical role in coupling ATPase hydrolysis and DNA translocation (Liu et al., 2017). We further observe a contact between a positively charged loop in ATPase lobe 1 (residues 832–837) and the second DNA gyre at SHL −6. This loop is present in CHD3, CHD4, and CHD5, but not in Snf2 or ISWI remodelers (Figure 3—figure supplement 1).

CHD4 binding distorts DNA at SHL +2

Comparison of our structure with a high-resolution X-ray structure of the free nucleosome (Vasudevan et al., 2010) reveals a conformational change in the DNA where the ATPase motor engages its DNA substrate (SHL +2) (Figure 3d). The high resolution of the nucleosome-CHD4 structure shows that ~5 DNA base pairs between SHL +1.5 and SHL +2.5 are pulled away from the octamer surface by up to 3 Å. This distortion does not include the previously observed ‘bulging’ or a ‘twist defect’ that is characterized by a 1 bp local underwinding of the DNA duplex and observed when the ATPase motor adopts the open/apo or ADP-bound states (Li et al., 2019). In contrast, the DNA distortion observed in our AMP-PNP-bound state is an intermediate between the bulged and the canonical DNA conformation (Figure 3d). Such an AMP-PNP-bound intermediate DNA state was predicted based on biochemical experiments (Winger et al., 2018). This observation demonstrates that the extent of DNA distortion at SHL +2 depends on the functional state of the ATPase motor and is consistent with the proposed twist defect propagation model of chromatin remodeling (Winger et al., 2018).

CHD4 binds the histone H4 tail

As observed for S. cerevisiae Chd1 (Farnung et al., 2017), H. sapiens CHD4 contacts the histone H4 tail with its ATPase lobe 2. The H4 tail is located between ATPase lobe 2 and the nucleosomal DNA at SHL +1.5. The conformation of the H4 tail differs from that observed in structures of the free nucleosome where the tail makes inter-nucleosomal contacts with the ‘acidic patch’ of a neighboring nucleosome. It also differs from the H4 position observed in a higher order structure where the H4 tail extends over the DNA interface between two nucleosomes (Schalch et al., 2005). A loop in lobe 2 of the ATPase (CHD4 residues 1001–1006) replaces the H4 tail in this position, apparently inducing H4 positioning that allows ATPase lobe 2 binding (Figure 4a).

CHD4 contacts H3 and H4.

(a) ATPase lobe 2 interacts extensively with the H4 tail. (b) A loop in ATPase lobe 2 contacts H3 alpha helix 1 and neighboring residues. (c) The double chromodomain of CHD4 contacts the H3 N-terminal tail. H3 core is shown in blue, H3 tail density from the low-pass filtered final map (7 Å) in teal, and the double chromodomain in purple.

ATPase lobe 2 contains a highly acidic cavity formed by Asp1080, Glu1083, Asp1084, and Glu1087 (Figure 4a). This acidic cavity is conserved across all CHD family members. The basic side chain of the H4 histone tail residue Arg17 inserts into this acidic cavity (Figure 4a). Similar interactions with the H4 tail have also been reported for Snf2 and ISWI remodelers (Armache et al., 2019; Yan et al., 2019). The side chain of H4 Lys16 also points toward the acidic cavity and is positioned in close proximity to residues Asp1080 and Glu1083. Acetylation of H4 Lys16 is therefore predicted to weaken these charge-based interactions and to reduce the affinity of chromatin remodellers for the H4 tail. This was noted before (Yan et al., 2016) and is consistent with CHD4 activity in repressed regions that lack such H4 acetylation.

CHD4 interacts with histone H3

The ATPase lobe 2 also contacts the core of histone H3 (alpha helix 1, Gln76 and Arg83) via CHD4 residues Asn1004 and Leu1009, respectively (Figure 4b). This contact is critical for chromatin remodeling. Deletion of the homologous region in Chd1 leads to abolishment of chromatin remodeling activity (Sundaramoorthy et al., 2018). However, it remains unclear if these contacts are required for proper substrate recognition and positioning or whether they are also necessary to generate the force required for DNA translocation. Low-pass filtering of our map further shows the H3 N-terminal tail trajectory, which extends to the double chromodomain (Figure 4c). The contact between the H3 tail and the double chromodomain could target CHD4 to nucleosomes methylated at Lys27 of H3 (Kuzmichev et al., 2002), a classical mark for gene repression.

Two CHD4 molecules can engage with the nucleosome

During 3D classification of our cryo-EM dataset we observed a distinct class of particles that showed two CHD4 molecules bound to the same nucleosome (Figure 5, Figure 1—figure supplements 24, Video 2). Refinement of this class of particles yielded a reconstruction at an overall resolution of 4.0 Å (FSC 0.143 criterion) (Table 1). A model of this nucleosome-CHD42 complex was obtained by docking the refined nucleosome-CHD4 model into the density and then placing another CHD4 molecule into the additional density observed on the opposite side. The resulting nucleosome-CHD42 complex structure shows pseudo-twofold symmetry with CHD4 molecules bound at SHL +2 and SHL −2 (Figure 5). The second CHD4 molecule uses its double chromodomain and PHD finger 2 to contact nucleosomal DNA at SHL +1 and +0.5, respectively. Binding of the second CHD4 molecule also did not lead to unwrapping of terminal DNA.

The nucleosome can bind two copies of CHD4.

Cartoon model of the nucleosome-CHD42 structure viewed from the top (a), and dyad view (b).

Video 2
Cryo-EM density and structure of the nucleosome-CHD42 complex.

Binding of two chromatin remodellers to a single nucleosome was previously observed for S. cerevisiae Chd1 (Sundaramoorthy et al., 2018) and H. sapiens SNF2H (Armache et al., 2019). However, in contrast to the structure of the nucleosome-SNF2H2 complex, we do not observe a distortion in the histone octamer due to the presence of the chromatin remodellers. Binding of two remodeler molecules could allow for higher efficiency in positioning the nucleosome at a precise location but necessitates coordination of the remodellers. A possible mechanism for coordination could be that twist defects that are introduced by remodeler binding are propagated from the entry SHL 2 into the exit side SHL 2 (Brandani et al., 2018; Brandani and Takada, 2018). Presence of the twist defect at the second remodeler binding site could interfere with the translocation activity of the second remodeler (Sabantsev et al., 2019).

Cancer-related CHD4 mutations

Many studies have reported mutations in CHD4 that are related to human diseases, in particular cancer (Xia et al., 2017). Mutations involved in various cancer phenotypes have been observed in the PHD finger 2, the double chromodomain, and both lobes of the ATPase motor. To elucidate effects of such mutations on CHD4 activity, the Drosophila melanogaster CHD4 homologue Mi-2 has been used as a model protein for functional analysis (Kovač et al., 2018). CHD4 mutations have been found to fall in two categories. Whereas some mutations influence ATPase and DNA translocation activity (Arg1162, His1196, His1151 and Leu1215), other mutations seem to change protein stability (Leu912, and Cys464) or disrupt DNA binding (Val558 and Arg572).

To try and rationalize these findings, we mapped known CHD4 mutations on our high-resolution structure (Figure 6, Table 2). Selected sites of mutation are described below. Mutation of residue His1151 to arginine results in a significant reduction of ATPase activity and abolishes chromatin remodeling activity (Kovač et al., 2018). The close proximity of this residue to motif Va (CHD4 residues 1147–1150) makes it likely that the mutation disrupts motif Va function, leading to an uncoupling of the ATPase activity from chromatin remodeling. Similar findings were made for Snf2 where mutation of the tryptophan residue in motif Va resulted in an uncoupling phenotype (Liu et al., 2017). The most frequently mutated residue in endometrial cancer, arginine 1162, is located in the ATPase motif VI. It forms an ‘arginine finger’ that directly interacts with AMP-PNP in our structure. Consistent with this observation, mutation of Arg1162 to glutamine impairs ATP hydrolysis in biochemical assays (Kovač et al., 2018).

CHD4 mutations in cancer and Sifrim-Hitz-Weiss syndrome.

Missense mutations that occur in endometrial cancer (blue spheres) and Sifrim-Hitz-Weiss syndrome (yellow spheres) mapped onto the CHD4 structure. Residue numbering is indicated. Nucleosomal DNA at SHL +2 is shown in a semi-transparent cartoon representation.

Table 2
CHD4 mutations in cancer and Sifrim-Hitz-Weiss syndrome.
Mutated
Residue
LocationPredicted effect based on structureBiochemical observations
Cancer
Cys464TyrPHD finger 2Disruption of Zn2+ binding in PHD finger 2Reduction in ATPase activity (Kovač et al., 2018)
Val558PheDouble chromodomainReduced ATPase activity (Kovač et al., 2018)
Arg572GlnDouble chromodomainDisruption of contact with tracking strandReduced DNA binding affinity, Loss of full remodeling activity and ATPase activity (Kovač et al., 2018)
Leu912ValATPase lobe 2No prediction madeReduction of ATPase activity (Kovač et al., 2018)
His1151ArgATPase lobe 2In close proximity to motif Va, might disrupt contact of Trp1148Reduction of ATPase activity, abolishment of remodeling activity (Kovač et al., 2018)
Arg1162GlnATPase lobe 2, motif VILocated in ATPase motif VI (arginine finger), Disruption of interaction with ATPReduction of ATPase activity (Kovač et al., 2018)
His1196TyrATPase lobe 2Located in the C-terminal bridge region, Removes negative regulationSpeed of chromatin remodeling is increased and better nucleosome centering capability (Kovač et al., 2018)
Leu1215ATPase lobe 2/C-terminal bridgeNot located in modeled region
Sifrim-Hitz-Weiss syndrome (Sifrim et al., 2016; Weiss et al., 2016)
Cys467TyrPHD finger 2Disruption of Zn2+ binding in PHD finger 2
Ser851TyrATPase lobe 1
Gly1003AspATPase lobe 2Disruption of contact with H3
Arg1068HisATPase lobe 2Disruption of structural integrity of RecA fold
Arg1127GlnATPase lobe 2Disruption of contact with DNA minor groove, equivalent arginine residue in SMARCA4 is implicated in ‘Coffin Siris syndrome’
Trp1148LeuATPase lobe 2, motif VaDisruption of contact with guide strandUncoupling of ATPase activity and chromatin remodeling (Liu et al., 2017)
Arg1173LeuDestabilization
Val1608IleNot located in modeled region

Other disease-related CHD4 mutations

De novo missense mutations in CHD4 are also associated with an intellectual disability syndrome with distinctive dysmorphisms (Sifrim et al., 2016; Weiss et al., 2016). Mutations observed in patients with this syndrome are located in PHD finger 2 (Cys467Tyr) and predominantly in ATPase lobe 2 (Ser851Tyr, Gly1003Asp, Arg1068His, Arg1127Gln, Trp1148Leu, Arg1173Leu, and Val1608Ile). We mapped the sites of these mutations onto our structure (Figure 6) and attempted to predict the effects of the mutations as far as possible (Table 2).

The Cys467Tyr mutation disrupts coordination of a zinc ion in PHD finger 2. Gly1003 in ATPase lobe2 is located in a loop near H3 alpha helix 1. Deletion of this loop in Chd1 results in a loss of chromatin remodeling activity (Sundaramoorthy et al., 2018). Residue Arg1068 forms a hydrogen bond network with the side chain of Thr1137 and the main chain carbonyl groups of Phe1112 and Gln1119. The Arg1068Cys mutation disrupts this network and is predicted to impair the integrity of the ATPase fold. Mutation of Arg1127 disrupts its interactions with the DNA minor groove (Figure 3c). The equivalent arginine residue in SMARCA4, which is one of the catalytic subunits of the BAF complex, has been implicated in the rare genetic disorder Coffin-Siris syndrome (Tsurusaki et al., 2012). Trp1148, which is part of ATPase motif Va, contacts the guide strand in a fashion similar to Chd1 and Snf2 (Farnung et al., 2017; Liu et al., 2017; Figure 3c). Mutation of this residue uncouples ATP hydrolysis and chromatin remodelling (Liu et al., 2017). Arg1173 inserts into an acidic pocket formed by residues Glu971, Asp1147, and Asp1153. Mutation of the arginine residue to leucine is likely to destabilize ATPase lobe 2 folding.

Discussion

Here, we provide the 3.1 Å resolution cryo-EM structure of human CHD4 engaged with a nucleosome and the 4.0 Å resolution structure of a nucleosome-CHD42 complex that contains two molecules of CHD4. Our structure of the nucleosome-CHD4 complex reveals how a subfamily II CHD remodeler engages with its nucleosomal substrate. We observe a distortion of nucleosomal DNA at SHL +2 in the presence of AMP-PNP. Similar observations were previously made for the Snf2 chromatin remodeler (Li et al., 2019; Liu et al., 2017) in its apo and ADP-bound states.

Our high-resolution structure fills a gap in our understanding of the mechanism of chromatin remodeling by capturing an additional enzymatic state. The DNA distortion at SHL +2 that we observed in the AMP-PNP bound state differs from distortions observed previously in the apo and ADP-bound state that involved a twist distortion (Li et al., 2019; Winger et al., 2018). This is consistent with a proposed ‘twist defect’ mechanism for chromatin remodeling (Li et al., 2019; Sabantsev et al., 2019). In this model, binding of the ATPase motor at SHL ± 2 induces a twist defect in the DNA. Subsequent ATP binding, captured by AMP-PNP and ADP·BeF3 structures, then leads to closing of the ATPase motor and to propagation of the twist defect toward the dyad. It is possible that previous nucleosome-Chd1 structures with ADP·BeF3 (Farnung et al., 2017; Sundaramoorthy et al., 2018) contained the same DNA distortion but that the lower resolution prevented its observation. Finally, ATP hydrolysis would reset the remodeller and the enzymatic cycle can resume at the next DNA position.

A major difference between the subfamily I remodeller Chd1 and the subfamily II remodeller CHD4 is that Chd1 induces unwrapping of the terminal nucleosomal DNA, whereas CHD4 does not change the DNA trajectory between SHL −7 and −5. DNA unwrapping is observed for Chd1 in structures and in solution and is independent of which ATP or transition state analogue is bound to the motor domain, indicating it is achieved with the use of binding energy only. Our observations are consistent with a single-molecule FRET study (Zhong et al., 2019). This major difference in Chd1 and CHD4 molecular function is likely related to a striking difference in cellular function. Whereas Chd1 functions in euchromatic regions of the genome during active transcription (Skene et al., 2014), CHD4 plays a central role in the establishment and maintenance of repressive genome regions. Consistent with these findings, DNA unwrapping should be prevented in stable heterochromatic regions. It is possible that these differences in functionality were achieved during evolution by the addition of distinct auxiliary domains in different CHD subfamilies.

Our structure also maps causative disease mutations and helps to investigate how these can impair CHD4 function. Our structure suggests that various mutations may disrupt DNA binding, impede ATP hydrolysis, or uncouple ATP hydrolysis and DNA translocation. The structure thus suggests the effects of CHD4 mutations in cancer and intellectual disability syndromes on chromatin remodeling. It also helps in understanding disease phenotypes of other chromatin remodelers such as the BAF complex that shows a related domain architecture for its ATPase motor. Due to its high resolution, the structure may also guide drug discovery using chromatin remodelers as targets in the future.

Materials and methods

Preparation of CHD4

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H. sapiens CHD4 (Uniprot Accession code Q14839-1) was amplified from human cDNA using the following ligation-independent cloning (LIC) compatible primer pair (Forward primer: 5’-TAC TTC CAA TCC AAT GCA ATG GCG TCG GGC CTG-3’, reverse primer: 5’-TTA TCC ACT TCC AAT GTT ATT ACT GCT GCT GGG CTA CCT G-3’). The PCR product containing CHD4 was cloned into a modified pFastBac vector (a gift from S. Gradia, UC Berkeley, vector 438 C, Addgene: 55220) via LIC. The CHD4 construct contains an N-terminal 6xHis tag, followed by an MBP tag, a 10x Asn linker sequence, and a tobacco etch virus protease cleavage site. All sequences were verified by Sanger sequencing.

The CHD4 plasmid (500 ng) was electroporated into DH10EMBacY cells (Geneva Biotech) to generate a bacmid encoding full-length H. sapiens CHD4. Bacmids were subsequently selected and prepared from positive clones using blue/white selection and isopropanol precipitation. V0 and V1 virus production was performed as previously described (Farnung et al., 2017). Hi5 cells (600 ml) grown in ESF-921 media (Expression Systems) were infected with 200 μl of V1 virus for protein expression. The cells were grown for 72 hr at 27°C. Cells were harvested by centrifugation (238 g, 4°C, 30 min) and resuspended in lysis buffer (300 mM NaCl, 20 mM Na·HEPES pH 7.4, 10% (v/v) glycerol, 1 mM DTT, 30 mM imidazole pH 8.0, 0.284 μg ml−l leupeptin, 1.37 μg ml−1 pepstatin A, 0.17 mg ml−1 PMSF, 0.33 mg ml−1 benzamidine). The cell resuspension was frozen and stored at −80°C.

H. sapiens CHD4 was purified at 4°C. Frozen cell pellets were thawed and lysed by sonication. Lysates were cleared by two centrifugation steps (18,000 g, 4°C, 30 min and 235,000 g, 4°C, 60 min). The supernatant containing CHD4 was filtered using 0.8 μm syringe filters (Millipore). The filtered sample was applied onto a GE HisTrap HP 5 ml (GE Healthcare), pre-equilibrated in lysis buffer. After sample application, the column was washed with 10 CV lysis buffer, 5 CV high-salt buffer (1 M NaCl, 20 mM Na·HEPES pH 7.4, 10% (v/v) glycerol, 1 mM DTT, 30 mM imidazole pH 8.0, 0.284 μg ml−1 leupeptin, 1.37 μg ml−1 pepstatin A, 0.17 mg ml−1 PMSF, 0.33 mg ml−1 benzamidine), and 5 CV lysis buffer. The protein was eluted with a gradient of 0–100% elution buffer (300 mM NaCl, 20 mM Na·HEPES pH 7.4, 10% (v/v) glycerol, 1 mM DTT, 500 mM imidazole pH 8.0, 0.284 μg ml−1 leupeptin, 1.37 μg ml−1 pepstatin A, 0.17 mg ml−1 PMSF, 0.33 mg ml−1 benzamidine). Peak fractions were pooled and dialysed for 16 hr against 600 ml dialysis buffer (300 mM NaCl, 20 mM Na·HEPES pH 7.4, 10% (v/v) glycerol, 1 mM DTT, 30 mM imidazole) in the presence of 2 mg His6-TEV protease. The dialysed sample was applied to a GE HisTrap HP 5 ml. The flow-through containing CHD4 was concentrated using an Amicon Millipore 15 ml 50,000 MWCO centrifugal concentrator. The concentrated CHD4 sample was applied to a GE S200 16/600 pg size exclusion column, pre-equilibrated in gel filtration buffer (300 mM NaCl, 20 mM Na·HEPES pH 7.4, 10% (v/v) glycerol, 1 mM DTT). Peak fractions were concentrated to ~40 μM, aliquoted, flash frozen, and stored at −80°C. Typical yields of H. sapiens CHD4 from 1.2 L of Hi5 insect cell culture are 2–4 mg.

Preparation of CHD1

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S. cerevisiae Chd1 (residues 1–1247) used for FRET assays was cloned, expressed, and purified similarly to the previously described strategy for full-length Chd1 (Farnung et al., 2017).

Nucleosome preparation

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Xenopus laevis histones were expressed and purified as described (Dyer et al., 2003; Farnung et al., 2017). DNA fragments for nucleosome reconstitution were generated by PCR essentially as described (Farnung et al., 2018). A vector containing the Widom 601 sequence was used as a template for PCR. Super-helical locations are assigned based on previous publications (Farnung et al., 2018; Farnung et al., 2017; Kujirai et al., 2018; Sundaramoorthy et al., 2018), assuming potential direction of transcription from negative to positive SHLs. Large-scale PCR reactions were performed with two PCR primers (Structural studies: forward primer: CC TGT TAT TCC TAG TAA TCA ATC AGT GCC TAT CGA TGT ATA TAT CTG ACA CGT GCC T, reverse primer: CCC CAT CAG AAT CCC GGT GCC G; FRET assay: forward primer:/5Cy3/CAA TCA GTG CCT ATC GAT GTA TAT ATC TGA CAC GTG CCT, reverse primer:/5Cy5/CCC CAT CAG AAT CCC GGT GCC G) at a scale of 25 mL. The DNA construct used for structural studies was designed based on previously reported constructs used for the study of CHD remodelers. Nucleosome core particle reconstitution was performed using the salt-gradient dialysis method (Dyer et al., 2003). Quantification of the reconstituted nucleosome was achieved by measuring absorbance at 280 nm. Molar extinction coefficients were determined for protein and nucleic acid components and were summed to yield a molar extinction coefficient for the reconstituted extended nucleosome.

Reconstitution of nucleosome-CHD4 complex

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Reconstituted nucleosome core particles and CHD4 were mixed at a molar ratio of 1:2. AMP-PNP was added at a final concentration of 1 mM and the sample was incubated for 10 min on ice. After 10 min compensation buffer was added to a final buffer concentration of 30 mM NaCl, 3 mM MgCl2, 20 mM Na⋅HEPES pH 7.5, 4% (v/v) glycerol, 1 mM DTT. The sample was applied to a Superose 6 Increase 3.2/300 column equilibrated in gel filtration buffer (30 mM NaCl, 3 mM MgCl2, 20 mM Na⋅HEPES pH 7.5, 5% (v/v) glycerol, 1 mM DTT). The elution was fractionated in 50 µL fractions and peak fractions were analyzed by SDS-PAGE. Relevant fractions containing nucleosome core particle and CHD4 were selected and cross-linked with 0.1% (v/v) glutaraldehyde. The crosslinking reaction was performed for 10 min on ice and subsequently quenched for 10 min using a final concentration of 2 mM lysine and 8 mM aspartate. The sample was transferred to a Slide-A-Lyzer MINI Dialysis Unit 20,000 MWCO (Thermo Scientific), and dialysed for 4 hr against 600 ml dialysis buffer (30 mM NaCl, 3 mM MgCl2, 20 mM Na⋅HEPES pH 7.4, 20 mM Tris⋅HCl pH 7.5, 1 mM DTT). The sample was subsequently concentrated using a Vivaspin 500 ultrafiltration centrifugal concentrator (Sartorius) to a final concentration of ~200–300 µM.

Cryo-EM analysis and image processing

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The nucleosome-CHD4 sample was applied to R2/2 gold grids (Quantifoil). The grids were glow-discharged for 100 s before sample application of 2 μl on each side of the grid. The sample was subsequently blotted for 8.5 s (Blot force 5) and vitrified by plunging into liquid ethane with a Vitrobot Mark IV (FEI Company) operated at 4°C and 100% humidity. Cryo-EM data were acquired on a Titan Krios transmission electron microscope (FEI/Thermo) operated at 300 keV, equipped with a K2 summit direct detector (Gatan) and a GIF Quantum energy filter. Automated data acquisition was carried out using FEI EPU software at a nominal magnification of 130,000 × in nanoprobe EF-TEM mode. Image stacks of 40 frames were collected in counting mode over 10 s. The dose rate was ~4.3–4.5 e per Å2 per s for a total dose of ~43–45 e−2. A total of 3904 image stacks were collected.

Micrograph frames were stacked and processed. All micrographs were CTF estimated and motion corrected using Warp (Tegunov and Cramer, 2018). Particles were picked using an in-house trained instance of the neural network BoxNet2 of Warp, yielding 650,598 particle positions. Particles were extracted with a box size of 3002 pixel and normalized. Image processing was performed with RELION 3.0-beta 2 (Zivanov et al., 2018). Using a 30 Å low-pass filtered ab initio model generated in cryoSPARC from 1679 particles (Figure 1—figure supplement 2c), we performed one round of 3D classification of all particle images with image alignment. One class with defined density for the nucleosome-CHD4 complex was selected for a second round of classification. The second round of classification resulted in two classes with one copy of CHD4 bound to the nucleosome. The respective classes were selected and 3D refined. The refined nucleosome-CHD4 model was subsequently CTF refined and the beam tilt was estimated based on grouping of beam tilt classes according to their exposure positions. The CTF refined particles were submitted to one additional round of masked 3D classification without image alignment. The mask encompassed CHD4. The most occupied class from this classification was subsequently CTF-refined. The final particle reconstruction was obtained from a 3D refinement with a mask that encompasses the entire nucleosome-CHD4 complex.

The nucleosome-CHD4 reconstruction was obtained from 89,623 particles with an overall resolution of 3.1 Å (gold-standard Fourier shell correlation 0.143 criterion). The final map was sharpened with a B-factor of −36 Å2. To exclude that the reconstruction could be a mixture of particles with CHD4 bound to either SHL –2 or SHL +2, CHD4 signal was subtracted and prior angular and translational information for every particle was removed. The subtracted particles were then refined against a synthetic nucleosome core particle map lacking CHD4. As expected, the refinement resulted in a reconstruction where only density for the nucleosome core particle was observed. Subsequently, the particle subtraction was reverted and a 3D classification without image alignment against a single class was performed. This 3D classification employed the angular and translational information provided from the subtraction refinement. The resulting reconstruction showed clear density for CHD4 only at SHL +2, and not at SHL −2, giving a clear indication that the final nucleosome-CHD4 reconstruction contains CHD4 bound only at SHL +2 (Figure 1—figure supplement 4f). We cannot rule out, however, that our map is still to some extent a mix of CHD4 bound on either side of the nucleosome.

The second round of 3D classification yielded a class with a nucleosome-CHD42 complex. The particles were subsequently classified and refined. The resulting reconstruction with 40,233 particles had an overall resolution of 4.0 Å (gold-standard Fourier shell correlation 0.143 criterion). The final map was sharpened with a B-factor of −86 Å2. Local resolution estimates for both structures were determined using the built-in RELION tool.

Model building

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Crystal structures of the X. laevis nucleosome with the Widom 601 sequence (Vasudevan et al., 2010) (PDB code 3LZ0) and the double chromodomain of CHD4 (PDB code 4O9I) were placed into the density of the nucleosome-CHD4 complex as rigid bodies using UCSF Chimera. The protein sequence of the ATPase motor of CHD4 (residues 706–1196) was ‘one-to-one threaded’ using the ATPase motor of S. cerevisiae Chd1 (PDB code 5O9G) as a template by employing Phyre2 (Kelley et al., 2015). The threaded model was placed into the density as a rigid body using UCSF Chimera (Goddard et al., 2018). Additional density belonging to helical extensions and loops present in the ATPase motor region were modeled de novo. The modeled sequence range 1405–1416 is assigned tentatively based on a previously published Chd1 crystal structure (PDB code 3MWY).

The nucleosome structure, double chromodomain structure, and ATPase motor model were adjusted manually in COOT (version 0.9-pre) (Emsley et al., 2010). The structure of PHD finger 2 (Mansfield et al., 2011) was then manually placed into the remaining, weaker density next to the double chromodomain and rigid-body docked (Figure 1—figure supplement 3), assisted by PDB code 6Q3M. Additional structural elements such as the H4 tail, the C-terminal bridge and loop regions of CHD4 were built using COOT. AMP-PNP and a coordinated Mg2+ ion were placed into the corresponding density. AMP-PNP was derived from the monomer library in COOT. The high resolution of our reconstruction enabled us to model some DNA-interacting side chains in two alternative conformations. The complete model was real-space refined in PHENIX (Afonine et al., 2018) with global minimization, local rotamer fitting, morphing, and simulated annealing. To model the nucleosome-CHD42 complex, the CHD4 model was duplicated and the second copy was rigid body docked into the additional density using UCSF ChimeraX (Goddard et al., 2018). The resulting structure was real space refined in PHENIX with global minimization, local rotamer fitting, morphing, and simulated annealing.

Förster resonance energy transfer (FRET) assay

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100 nM of NCP with Cy3 and Cy5 5’-terminal DNA ends was incubated with 300 nM S. cerevisiae Chd1 (residues 1–1247) or full-length CHD4 and 1 mM ADP·BeF3 or 1 mM AMP-PNP at final reaction conditions of 50 mM NaCl, 3 mM MgCl2, 20 mM Na⋅HEPES pH 7.4, 0.1 mg/mL BSA, 10% (v/v) glycerol, 1 mM DTT. To increase FRET efficiency, we used a DNA construct that is shortened by 18 bp on the DNA exit side compared to the construct used for the structural studies. The sample was subsequently incubated for 30 min and transferred to 384-well plates. The reaction was then monitored using a fluorescence emission scan from 520 to 740 nm in a Tecan infinite m1000 pro plate reader with an excitation wavelength of 510 nm. All reactions were performed in triplicates in independent experiments. Emission spectra were normalized by total emissions. Averages of the triplicates and corresponding standard deviations are reported. The results were plotted using Matplotlib.

Figure generation

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Figures were generated using PyMol (version 2.2.2) and UCSF ChimeraX.

References

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Decision letter

  1. Geeta J Narlikar
    Reviewing Editor; University of California, San Francisco, United States
  2. Jessica K Tyler
    Senior Editor; Weill Cornell Medicine, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

The dearth of atomic resolution structures has been a major roadblock in understanding the mechanisms of chromatin remodelers. The high-quality EM structure and the associated EM data in this work is expected to be a great resource for the chromatin field to develop and test mechanistic models.

Decision letter after peer review:

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your work entitled "Nucleosome-CHD4 chromatin remodeller structure explains human disease mutations" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered for publication in eLife.

As you will see from the individual reviews attached below all the reviewers found the structure to be of high quality. However, in the discussion amongst the reviewers it was also agreed that the structure by itself does not sufficiently advance our understanding of remodeling mechanisms. Comparison with previous Chd1 structures forms a key component of the mechanistic conclusions made in this work. The reviewers thought that such direct comparisons were not appropriate primarily because of the different ATP states of the two structures. Additionally, the first Chd1 structure had other proteins in the mix that are not present in the context of CHD4. Comparing structures of CHD4 and Chd1 bound to nucleosomes in the presence of the same ATP analog would be ideal. However, the reviewers recognized that this may be difficult and take time. So, in their discussion they tried to come up with possible biochemical experiments (instead of another structure) that could be carried out to test the mechanistic predictions of your current CHD4-nucleosome structure. These are summarized below.

1) Directly compare terminal DNA unwrapping by CHD4 and Chd1 (positive control) in the presence of AMP-PNP and ADP-BeFx using a FRET assay.

2) Introduce gaps at SHL 2 and determine if deformation of the DNA at SHL 2 has a functional role.

3) Test the functional effects of the new contacts and compare to corresponding effects with Chd1.

If any of these biochemical experiments or any others that you choose to carry out provide new mechanistic insights, then we would welcome a resubmission.

Reviewer #1:

Recent high-resolution cryo-EM structures of ATP-dependent chromatin remodelers are shedding new light on the mechanisms of these complex enzymes. Here the authors use cryo-EM to study the structure of CHD4, a human CHD enzyme, bound to a nucleosome in the presence of AMP-PNP. They find that the ATPase domain binds at SHL 2 as predicted from studies of other CHD enzymes and observe a distortion of nucleosomal DNA at this location. However, in contrast to their earlier structure of S. cerevisiae Chd1, the authors do not see unpeeling of the terminal DNA that is proximal to the bound ATPase domain. They argue that this difference may arise because CHD4 does not have the SANT-SLIDE DNA binding domains present in Chd1. They also observe interactions with the H4 tail as predicted by previous studies. Finally, they map specific disease associated mutations map on to the ATPase and accessory domains of Chd4.

Overall, the cryo-EM data appears to be of very high quality. My main concern is that the findings do not substantially move the field forward compared to previous structural studies on chromatin remodelers. There are some new findings in this work that have the potential to be mechanistically significant, but these need additional tests. Below are some suggestions for raising the mechanistic impact.

1) The authors suggest that the absence of unpeeled DNA compared to Chd1 may arise because of the absence of a known DNA binding domain. This difference could in principle point to significant mechanistic differences between the two CHD proteins due to different ways of engaging the DNA. However, the differences could also arise due to the difference in nucleotide state between the two studies. The author's previous study used ADP-BeFx while this study used AMP-PNP. To test for mechanistic differences between Chd4 and Cdh1 the authors should compare the two structures in the same ATP state.

2) The presence of distorted DNA is interesting, but the extent of distortion appears much less than observed with Snf2. The authors should test the mechanistic significance of the distortion by classical approaches previously used with ISWI and SWI/SNF enzymes. These experiments used nucleosomes with nicks or gaps at SHL 2. If the authors find substantial reduction in nucleosome sliding but not much reduction in nucleosome binding using nucleosomes with gaps or nicks at SHL 2, then this will imply that distortion of the DNA at SHL 2 plays a functional role.

3) The authors should test the consequences of mutating the specific interactions identified in their structure on remodeling by CHD4. This includes interactions made with the H4 tail, the histone surfaces and DNA. Comparison of the magnitude of the effects of these mutations with the corresponding published effects observed with Snf2, Chd1 and ISWI enzymes will allow for a direct comparison of how different remodelers use interactions with the nucleosome to drive octamer sliding.

Reviewer #2:

ATP-dependent chromatin remodeling is one of the mechanisms to modulate nucleosome structure, using the energy from ATP hydrolysis and dedicated ATPases. These macromolecular machines are employed and influence a range of processes, such as transcription, replication or repair. To maintain or effectuate changes in chromatin, eukaryotic cells evolved four known major families of remodeling factors: SWI/SNF, ISWI, Ino80 and CHD, that have their unique properties.

Farnung et al. report a high-resolution structure of CHD4 bound to a nucleosome, solved using cryo-EM. In their article, they use a full-length human CHD4 construct and a nucleosome with additional extranucleosomal DNA (4 bp on the entry and 30 bp at the exit of the nucleosome) and reveal the mode of binding and interaction between them. This is the first structure of group II CHD ATP-dependent remodelers, following on the studies by Farnung et al., 2017 of group I remodeler, yeast CHD1.

This is a well-written, strictly structural paper that provides a view on an important chromatin remodeler that in the eukaryotic cells is a part of larger remodeling complexes, such as NuRD and ChAHP. The authors could have provided some FRET data to confirm their model. This is important, especially since they derive functional differences from structural comparison of two not-fully resolved remodelers from different organisms on different nucleosomal substrates, using different ATP analogues. However, after some modifications, this data and paper could be accepted, as it is an important and well-approached study.

1) Upon comparing the PDB 3LZ0 with 6RYR, authors modeled 4 base-pairs on the exit site, which leads to the following question:

a) The map EMD-10058_6RYR: If there is 4bp on the entry, and 30bp at the exit, upon filtering the density, why would there be such a large additional amount of density on the entry-side at the low threshold level? Could that be an indication of a mixed population in the final reconstruction, or a translocation?

2) The authors write: "Structural comparisons show that CHD4, in contrast to Chd1, does not induce unwrapping of terminal DNA." The follow up to this is: "In contrast to the nucleosome-Chd1 structure (Farnung et al., 2017), we did not observe unwrapping of nucleosomal DNA from the histone octamer on the second DNA gyre at SHL -6 and -7 (Figure 2)". However, upon closer examination, this DNA density, while not unwrapped, appears to exhibit a lower occupancy than the rest of the nucleosome. Is that an effect of incomplete classification, or a partial distortion by the remodeler?

3) Regarding CHD1 and CHD4 comparison:

a) If comparing it only to the CHD1 publication from 2017 by Farnung et al., the sample there was significantly different. It involved CHD1-FACT-Paf1C-nucleosome assembly, of which only CHD1-nucleosome was visible. The DNA substrate was also different, containing 63 base pairs of extranucleosomal DNA on the exit side vs 30 in this study. Sundaramoorthy et al., 2018 contained only CHD1-nucleosome, but also with a different nucleosome substrate. Could the extranucleosomal DNA be of significance in this study, and could the authors provide a 1-2 sentence explanation of their particular exit/entry side DNA construct rationale?

b) Authors are comparing a yeast CHD1 bound by ADP-BeFx3, with human CHD4 bound by AMP-PNP. However, there were reported published cases where a comparison of states using cryo-EM involving those two ATP-analogues yielded different results, or different distribution of states (e.g.: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5307450/). Could it be completely excluded here?

Reviewer #3:

This manuscript describes a high-resolution cryo-EM structure of CHD4 bound to a nucleosome, thus adding to a small but growing number of chromatin remodeling protein structures. The main strength of the work is the high-resolution structure of this disease-relevant protein and the ability to discuss previous work in a structural context. Comparisons are made to previously defined remodeler-nucleosome structures, which mainly point to commonalities with a few noted exceptions including a lack of exit-side DNA distortion. An intriguing minor deformation in DNA trajectory is observed in the presence of AMP-PNP, which is made possible by the high resolution. Other strengths include determination of the placement of double chromodomains and one of two PHD fingers (albeit to lower resolution) as well as determination of a CHD3/4/5-specific contact with nucleosomal DNA. CHD4 disease-specific mutations are mapped onto the structure to help explain their previously defined functional consequences.

The biggest limitation of the work is the lack of functional information. This manuscript is entirely structural modeling. Though an exceptionally high-resolution structure for a remodeler-nucleosome complex, many of the findings were previously predicted, which makes the insight from the structure more limited than other contemporary remodeler-nucleosome structures. Another limitation is that the structure lacks a significant portion of the CHD4 protein, though this is not mentioned explicitly in the text. Figure 1 displays the fragment that was modeled, but it would be worthwhile to discuss that regions of CHD4 that may be functional or contributing to the remodeling cycle are not resolved. Otherwise it would be important to show that the modeled structure that is being used to justify a remodeling mechanism is sufficient for CHD4 remodeling.

It would be useful to incorporate at least some functional data, particularly to demonstrate function of the newly defined contacts (e.g. loop 832-837, Asn1010, Arg1068, Arg1173) to help solidify structure-function relationships that are proposed.

The title contains "explains disease mutations" but most (perhaps all) of the functional consequences of these disease mutations were defined previously in Mi-2 (Kovac et al) or could have been predicted from highly-conserved ATPase regions. While the manuscript gives credit to the previous characterizations, I believe the title overstates the contribution of the structure. Even simply changing to "Nucleosome-CHD4 chromatin remodeller structure maps human disease mutations" would be more acceptable. In addition, making it transparent in the Abstract that the effects of these mutations were previously defined would be appreciated.

Similarly, the Discussion states that "our structure elucidates the mechanism of chromatin remodeling". More appropriately, the structure may add to or fill gaps in models of the chromatin remodeling mechanism.

The exit-side Discussion section relating heterochromatin to lack of exit-side DNA dynamics in the cryo-EM structure seems speculative and unsupported. It is simpler to state that the lack of unwrapping at the exit side may be due to lack of DNA binding domain. However, unwrapping may still be seen in the context of CHD4 complexes like NuRD and ChAHP. It may be too early to speculate about intermediate DNA unwrapping states, as this structure captures one part of the CHD4 remodeling cycle that is complex and may require some distortion of DNA on both sides for nucleosome repositioning to occur.

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for submitting your work entitled "Nucleosome-CHD4 chromatin remodeller structure maps human disease mutations" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Senior Editor. The reviewers have opted to remain anonymous.

Earlier we had said that if any of the biochemical experiments that we had listed or any others that you choose to carry out provide new mechanistic insights then we would welcome a resubmission. In this context, the reviewers appreciated the addition of biochemical data to the resubmission. However, after extensive discussion all reviewers felt that the added FRET data did not provide sufficient new mechanistic insight to merit publication as a Research Article. At the same time, they agreed that your high-quality EM structure and the EM data would be a valuable resource for the chromatin field to develop and test mechanistic models. We can therefore offer to consider your work under the Tools and Resources category of eLife.

If you would like for your paper to be published in the Tools and Resources category, we ask that you submit a revised version that addresses the remaining EM-related questions. The major EM related question is summarized below.

As pointed out by our EM expert reviewer it appears that the structure could be a mix of the SHL +2 and SHL -2 bound particles. In this situation, the way to clearly define the structure would be to mask out or delete computationally the density for CHD4 and align particles based on some other feature. Then, without again aligning the data, go back to the original particles and project back the resulting information. This could show if there are SHL +2 and SHL -2 bound particles – the final maps could be at lower resolutions, nevertheless, it would be more precise. Even if this analysis shows no mixture of particles, we feel the analysis is necessary for conclusive interpretation of the data.

Carrying out such an analysis would comprehensively address the following two prior questions asked by the reviewer.

1) Upon comparing the PDB 3LZ0 with 6RYR, authors modeled 4 base-pairs on the exit site, which leads to the following question: The map EMD-10058_6RYR: If there is 4bp on the entry, and 30bp at the exit, upon filtering the density, why would there be such a large additional amount of density on the entry-side at the low threshold level? Could that be an indication of a mixed population in the final reconstruction, or a translocation?

2) The authors write: "Structural comparisons show that CHD4, in contrast to Chd1, does not induce unwrapping of terminal DNA." The follow up to this is: "In contrast to the nucleosome-Chd1 structure (Farnung et al., 2017), we did not observe unwrapping of nucleosomal DNA from the histone octamer on the second DNA gyre at SHL -6 and -7 (Figure 2)". However, upon closer examination, this DNA density, while not unwrapped, appears to exhibit a lower occupancy than the rest of the nucleosome. Is that an effect of incomplete classification, or a partial distortion by the remodeler?

Reviewer #1:

The FRET data presented by the authors addresses part of my core concern about comparing DNA unpeeling by Chd4 vs. Chd1. However, the experiments are quite minimal. Beyond suggesting some unpeeling in one context and not in the other, they do not provide much new insight into the mechanistic differences between Chd1 and Chd4.

In addition, several controls are missing.

1) The authors need to mention describe how they ensured that the nucleosome was fully bound by Chd1 and Chd4 under the FRET experimental conditions. This is important in the case of Chd4, which shows no FRET change.

2) To rule out any environmental effects on the Cy dyes upon binding by Chd1, the authors should show a scan for direct excitation of Cy5. The absence of an effect with direct excitation of Cy5 would rule out an environmental effect on Cy5.

3) It is not clear what is the significance of the extent of FRET change. Some comparison to the FRET changes observed upon increasing salt would allow for calibrating the observed effect with respect to other contexts where DNA is unpeeled.

Reviewer #2:

This revised manuscript from Farnung et al. presents a high resolution cryo-EM structure of the nucleosome core particle in complex with human CHD4, a disease-relevant chromatin remodeling protein with known roles in transcriptional repression. The structure is a nice addition to our limited but growing number of chromatin remodeling proteins in complex with nucleosome substrates. The work defines a potential remodeling intermediate where nucleosomal DNA is distorted at SHL +2, and contrasts CHD4 with Chd1 by showing CHD4 does not unwrap terminal DNA. The structure provides a valuable resource to the chromatin community and it bridges the gap between known structure, known chromatin remodeler function, and known disease mutations.

The major drawback from the work is the limited insight into new chromatin remodeler biology. There is limited new knowledge gained regarding how chromatin remodeling proteins work, and the function of most of the disease mutations were predictable based on a significant number of previous publications describing ATPase and chromatin remodeler biochemistry. Even the described remodeling intermediate is speculative, since there are no comparable structures of CHD4 in other nucleotide-bound states, so the extent of disruption of SHL +2 as a function of nucleotide state is unclear. As mentioned in the previous review, minimal biochemical experiments testing some of the newly identified remodeler-nucleosome contact points would have been quite useful to address these concerns.

Reviewer #3:

ATP-dependent chromatin remodeling is one of the mechanisms to modulate nucleosome structure, using the energy from ATP hydrolysis and dedicated ATPases. These macromolecular machines are employed and influence a range of processes, such as transcription, replication or repair. To maintain or effectuate changes in chromatin, eukaryotic cells evolved four known major families of remodeling factors: SWI/SNF, ISWI, Ino80 and CHD, that have their unique properties.

Farnung et al. report a high-resolution structure of CHD4 bound to a nucleosome, solved using cryo-EM. In their article, they use a full-length human CHD4 construct and a nucleosome with additional extranucleosomal DNA (4 bp on the entry and 30 bp at the exit of the nucleosome) and reveal the mode of binding and interaction between them. This is the first structure of group II CHD ATP-dependent remodelers, following on the studies by Farnung et al., 2017 of group I remodeler, yeast CHD1.

This is a well-written, well-approached structural paper that provides a view on an important chromatin remodeler that in the eukaryotic cells is a part of larger remodeling complexes, such as NuRD and ChAHP.

This is the second time this paper has been reviewed by me; thus, upon reading the paper and the changes made by the authors, I took the liberty of resubmitting this review in an almost unchanged form. From the previous submission, it is clear that the authors took their time to address the most criticized part and performed FRET studies. This is convincing and removes the greatest hurdle in their paper.

However, I still want to request answers to the question I made in the past, that has not been addressed.

I could not see it from the classification, so, could authors provide an indication of how they made sure that their nucleosome orientation in the reconstruction was done properly (30/4bp)? Could it be that the single CHD4 could bind on either side (as in Armache et al., 2019, SNF2H), and thus the EM assignment would have to be extra careful?

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Nucleosome-CHD4 chromatin remodeller structure maps human disease mutations" for further consideration by eLife. Your revised article has been evaluated by Jessica Tyler (Senior Editor) and a Reviewing Editor.

The reviewers agree that the work will provide a very valuable resource to the chromatin community. However, our EM expert reviewer has some remaining concerns. Below are excerpted the two key issues that the reviewer raises:

Since the authors decided not to provide a deeper insight into the mixed dataset angle I asked about before, and the density on either side of the nucleosome seems to not clearly support the statement about the 4bp, I would want them to add a sentence about this in the text. A statement, such as, for example: "We cannot rule out that our map is still to some extent a mix of the CHD4 bound on either side of the nucleosome, as filtering of the map suggests the presence of more than 4 bp". As this is to be a resource paper, I also want to suggest the paper acceptance provided that the raw EM data is uploaded to EMPIAR as soon as possible.

All three reviewers have discussed these comments and agree that the following actions are needed prior to acceptance:

1) Addition of the following sentence in the text: "We cannot rule out that our map is still to some extent a mix of the CHD4 bound on either side of the nucleosome, as filtering of the map suggests the presence of more than 4 bp".

2) Uploading the raw EM data to EMPIAR.

https://doi.org/10.7554/eLife.56178.sa1

Author response

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

[…]

1) Directly compare terminal DNA unwrapping by CHD4 and Chd1 (positive control) in the presence of AMP-PNP and ADP-BeFx using a FRET assay.

2) Introduce gaps at SHL 2 and determine if deformation of the DNA at SHL 2 has a functional role.

3) Test the functional effects of the new contacts and compare to corresponding effects with Chd1.

If any of these biochemical experiments or any others that you choose to carry out provide new mechanistic insights, then we would welcome a resubmission.

We thank the reviewers for good suggestions. We now provide the key additional biochemical experiment that the reviewers suggested, the direct comparison of Chd1 and CHD4 in their ability to detach nucleosomal DNA, as monitored by FRET (Suggestion #1). We carried out these assays in the presence of ADP.BeF3 or AMP-PNP. We found that, as expected from the structural data and comparisons, Chd1 detaches/unwraps DNA in the presence of either ATP or transition state analogue, whereas CHD4 does not. This assay complements our structural comparison with Chd1 and indicates an important mechanistic difference between these two CHD proteins. The new data and figure clearly improved our manuscript by providing not only structural but also biochemical evidence for mechanistic differences between members of two CHD subfamilies. We think the resulting manuscript is acceptable for publication. Please also refer to the detailed responses with respect to individual concerns by reviewers below.

Reviewer #1:

[…]

1) The authors suggest that the absence of unpeeled DNA compared to Chd1 may arise because of the absence of a known DNA binding domain. This difference could in principle point to significant mechanistic differences between the two CHD proteins due to different ways of engaging the DNA. However, the differences could also arise due to the difference in nucleotide state between the two studies. The author's previous study used ADP-BeFx while this study used AMP-PNP. To test for mechanistic differences between Chd4 and Cdh1 the authors should compare the two structures in the same ATP state.

We thank the reviewer for their comment. As explained above, we have addressed this by a FRET assay that monitors DNA unwrapping. The FRET assay was performed with both remodellers in the presence of ADP.BeF3- or AMP-PNP. Indeed, Chd1 unwraps DNA in the presence of either ATP/transition state analogue, whereas CHD4 does not. These data are presented now along the structural comparisons of the Chd1- and CHD4-nucleosome complexes in Figure 2B, C. The FRET assay confirms a significant mechanistic difference between these two CHD proteins. With respect to new structures, we trust the reviewer understands this is beyond the scope of the current work, and this is also what we understood from the editorial decision.

2) The presence of distorted DNA is interesting, but the extent of distortion appears much less than observed with Snf2. The authors should test the mechanistic significance of the distortion by classical approaches previously used with ISWI and SWI/SNF enzymes. These experiments used nucleosomes with nicks or gaps at SHL 2. If the authors find substantial reduction in nucleosome sliding but not much reduction in nucleosome binding using nucleosomes with gaps or nicks at SHL 2, then this will imply that distortion of the DNA at SHL 2 plays a functional role.

See above, we did not use this approach but concentrated to the comparisons between CHD4 and Chd1. We trust the reviewer understands this is beyond the scope of the current work, and this is also what we understood from the editorial decision.

3) The authors should test the consequences of mutating the specific interactions identified in their structure on remodeling by CHD4. This includes interactions made with the H4 tail, the histone surfaces and DNA. Comparison of the magnitude of the effects of these mutations with the corresponding published effects observed with Snf2, Chd1 and ISWI enzymes will allow for a direct comparison of how different remodelers use interactions with the nucleosome to drive octamer sliding.

See above, we did not use this approach but concentrated to the comparisons between CHD4 and Chd1. We trust the reviewer understands this is beyond the scope of the current work, and this is also what we understood from the editorial decision.

Reviewer #2:

[…]

1) Upon comparing the PDB 3LZ0 with 6RYR, authors modeled 4 base-pairs on the exit site, which leads to the following question:

a) The map EMD-10058_6RYR: If there is 4bp on the entry, and 30bp at the exit, upon filtering the density, why would there be such a large additional amount of density on the entry-side at the low threshold level? Could that be an indication of a mixed population in the final reconstruction, or a translocation?

We thank the reviewer for their comment. Our analysis revealed additional density of extranucleosomal DNA on the entry and exit side. The entry side, however, only showed additional density for approximately 4 base pairs as can be expected from the used DNA sequence. We did not choose to model these additional 4 base pairs because of the poor signal/noise for this region. In contrast, we were able to observe less ambiguous density at the DNA exit side and accordingly modelled 4 clearly distinguishable base pairs. We would like to point out that a mixed population or translocation events are highly unlikely due to the fact that we were able to assign the DNA register unambiguously using the density around the nucleosomal dyad, which exhibits high resolution (Figure 3C).

2) The authors write: "Structural comparisons show that CHD4, in contrast to Chd1, does not induce unwrapping of terminal DNA." The followup to this is: "In contrast to the nucleosome-Chd1 structure (Farnung et al., 2017), we did not observe unwrapping of nucleosomal DNA from the histone octamer on the second DNA gyre at SHL -6 and -7 (Figure 2)". However, upon closer examination, this DNA density, while not unwrapped, appears to exhibit a lower occupancy than the rest of the nucleosome. Is that an effect of incomplete classification, or a partial distortion by the remodeler?

The lower occupancy is likely due to a higher flexibility at SHL -7 and for the extranucleosomal DNA. Nevertheless, the flexibility around SHL -7 is comparable with the flexibility observed for canonical NCPs (see also FRET assay in Figure 2). Therefore, it is unlikely to be due to a partial distortion by the remodeler.

3) Regarding CHD1 and CHD4 comparison:

a) If comparing it only to the CHD1 publication from 2017 by Farnung et al., the sample there was significantly different. It involved CHD1-FACT-Paf1C-nucleosome assembly, of which only CHD1-nucleosome was visible. The DNA substrate was also different, containing 63 base pairs of extranucleosomal DNA on the exit side vs 30 in this study. Sundaramoorthy et al., 2018 contained only CHD1-nucleosome, but also with a different nucleosome substrate. Could the extranucleosomal DNA be of significance in this study, and could the authors provide a 1-2 sentence explanation of their particular exit/entry side DNA construct rationale?

We thank the reviewer for pointing this out. As indicated in Figure 1—figure supplement 1, we used the canonical 145 bp Widom 601 with additional 4 bp on the entry side of the DNA and 30 bp of linker DNA. The full sequence is given in the Materials and methods section of the submitted manuscript. The construct was designed based on the published NCP-Chd1 structures that clearly indicated that a construct with 30 bp of extranucleosomal DNA is sufficient for binding by a DNA binding region (as present in Chd1). We added a corresponding statement on this to the manuscript Materials and methods section.

b) Authors are comparing a yeast CHD1 bound by ADP-BeFx3, with human CHD4 bound by AMP-PNP. However, there were reported published cases where a comparison of states using cryo-EM involving those two ATP-analogues yielded different results, or different distribution of states (e.g.: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5307450/). Could it be completely excluded here?

We have addressed this comment by a FRET assay to monitor DNA unwrapping by both chromatin remodellers in the presence of ADP.BeF3 or AMP-PNP. We found that Chd1 unwraps DNA in the presence of either ATP/transition state analogue, whereas CHD4 does not. Thus, the concern is excluded.

Reviewer #3:

[…]

The biggest limitation of the work is the lack of functional information. This manuscript is entirely structural modeling. Though an exceptionally high-resolution structure for a remodeler-nucleosome complex, many of the findings were previously predicted, which makes the insight from the structure more limited than other contemporary remodeler-nucleosome structures. Another limitation is that the structure lacks a significant portion of the CHD4 protein, though this is not mentioned explicitly in the text. Figure 1 displays the fragment that was modeled, but it would be worthwhile to discuss that regions of CHD4 that may be functional or contributing to the remodeling cycle are not resolved. Otherwise it would be important to show that the modeled structure that is being used to justify a remodeling mechanism is sufficient for CHD4 remodeling.

It would be useful to incorporate at least some functional data, particularly to demonstrate function of the newly defined contacts (e.g. loop 832-837, Asn1010, Arg1068, Arg1173) to help solidify structure-function relationships that are proposed.

We thank the reviewer for this suggestion. Note we have added the key experiment that demonstrates a key difference between CHD4 and Chd1 in their ability to detach/unwrap terminal DNA. Rather than adding details, this experiment and comparison provide new conceptual insights into the CHD family of remodelers. See above for details.

The title contains "explains disease mutations" but most (perhaps all) of the functional consequences of these disease mutations were defined previously in Mi-2 (Kovac et al) or could have been predicted from highly-conserved ATPase regions. While the manuscript gives credit to the previous characterizations I believe the title overstates the contribution of the structure. Even simply changing to "Nucleosome-CHD4 chromatin remodeller structure maps human disease mutations" would be more acceptable. In addition, making it transparent in the Abstract that the effects of these mutations were previously defined would be appreciated.

We thank the reviewer for their suggestion and have changed the title of the manuscript accordingly to “Nucleosome-CHD4 chromatin remodeller structure maps human disease mutations”.

Similarly, the Discussion states that "our structure elucidates the mechanism of chromatin remodeling". More appropriately, the structure may add to or fill gaps in models of the chromatin remodeling mechanism.

We have changed the Discussion accordingly.

The exit-side Discussion section relating heterochromatin to lack of exit-side DNA dynamics in the cryo-EM structure seems speculative and unsupported. It is simpler to state that the lack of unwrapping at the exit side may be due to lack of DNA binding domain. However, unwrapping may still be seen in the context of CHD4 complexes like NuRD and ChAHP. It may be too early to speculate about intermediate DNA unwrapping states, as this structure captures one part of the CHD4 remodeling cycle that is complex and may require some distortion of DNA on both sides for nucleosome repositioning to occur.

We agree and have made sure this is correctly understood. Note we added the key experiment showing DNA dynamics in solution with the use of FRET. See above for details.

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

If you would like for your paper to be published in the Tools and Resources category, we ask that you submit a revised version that addresses the remaining EM-related questions. The major EM related question is summarized below.

We agree to publish our manuscript in the Tools and Resources category.

We have addressed the remaining EM-related questions as detailed below.

As pointed out by our EM expert reviewer it appears that the structure could be a mix of the SHL +2 and SHL -2 bound particles. In this situation, the way to clearly define the structure would be to mask out or delete computationally the density for CHD4 and align particles based on some other feature. Then, without again aligning the data, go back to the original particles and project back the resulting information. This could show if there are SHL +2 and SHL -2 bound particles – the final maps could be at lower resolutions, nevertheless, it would be more precise. Even if this analysis shows no mixture of particles, we feel the analysis is necessary for conclusive interpretation of the data.

Carrying out such an analysis would comprehensively address the following two prior questions asked by the reviewer.

1) Upon comparing the PDB 3LZ0 with 6RYR, authors modeled 4 base-pairs on the exit site, which leads to the following question: The map EMD-10058_6RYR: If there is 4bp on the entry, and 30bp at the exit, upon filtering the density, why would there be such a large additional amount of density on the entry-side at the low threshold level? Could that be an indication of a mixed population in the final reconstruction, or a translocation?

2) The authors write: "Structural comparisons show that CHD4, in contrast to Chd1, does not induce unwrapping of terminal DNA." The follow up to this is: "In contrast to the nucleosome-Chd1 structure (Farnung et al., 2017), we did not observe unwrapping of nucleosomal DNA from the histone octamer on the second DNA gyre at SHL -6 and -7 (Figure 2)". However, upon closer examination, this DNA density, while not unwrapped, appears to exhibit a lower occupancy than the rest of the nucleosome. Is that an effect of incomplete classification, or a partial distortion by the remodeler?

Note that our structure is at high enough resolution to support the unique assignment of the DNA register as shown in Figure 1—figure supplement 3H (purine and pyrimidine bases are distinguished, thus the DNA is uniquely assigned).

We have nevertheless followed the additional suggestion of the reviewer and have performed an analysis to test for possible symmetry artifacts. In short, CHD4 signal was subtracted from the Coulomb potential map and a masked refinement on the remaining NCP was performed with a synthetic NCP map as the model map. The particles were reverted to their non-subtracted state while maintaining the translational and angular information from the NCP refinement. Subsequently, a classification without image alignment was performed that resulted in clear density for CHD4 on only one side of the NCP. This clearly argues against a mixture of states where CHD4 is bound at SHL -2 and SHL +2 in the final reconstruction.

We added a corresponding paragraph to the Materials and methods section and have added these results to Figure 1—figure supplement 4F. Together with our high-resolution map, this entirely removes the concern.

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

[…]

1) Addition of the following sentence in the text: "We cannot rule out that our map is still to some extent a mix of the CHD4 bound on either side of the nucleosome, as filtering of the map suggests the presence of more than 4 bp".

We have added the sentence "We cannot rule out, however, that our map is still to some extent a mix of CHD4 bound on either side of the nucleosome.” to the text.

2) Uploading the raw EM data to EMPIAR.

The raw EMPIAR data has been uploaded to the EMPIAR database with accession code EMPIAR-10411. This was already previously indicated in the last revision of the manuscript. The accession codes for PDB, EMDB, and EMPIAR datasets can be found in the data availability statement. All data sets will be made available upon publication.

https://doi.org/10.7554/eLife.56178.sa2

Article and author information

Author details

  1. Lucas Farnung

    Max Planck Institute for Biophysical Chemistry, Department of Molecular Biology, Göttingen, Germany
    Contribution
    Conceptualization, Formal analysis, Supervision, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    For correspondence
    Lucas.Farnung@mpibpc.mpg.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8200-2493
  2. Moritz Ochmann

    Max Planck Institute for Biophysical Chemistry, Department of Molecular Biology, Göttingen, Germany
    Contribution
    Investigation, Visualization, Writing - review and editing
    Competing interests
    No competing interests declared
  3. Patrick Cramer

    Max Planck Institute for Biophysical Chemistry, Department of Molecular Biology, Göttingen, Germany
    Contribution
    Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    patrick.cramer@mpibpc.mpg.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5454-7755

Funding

Deutsche Forschungsgemeinschaft (SFB1064)

  • Patrick Cramer

Deutsche Forschungsgemeinschaft (SFB860)

  • Patrick Cramer

European Research Council (693023)

  • Patrick Cramer

Volkswagen Foundation

  • Patrick Cramer

Deutsche Forschungsgemeinschaft (EXC 2067/1-390729940)

  • Patrick Cramer

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank past and present members of the Cramer laboratory. We thank C Oberthür for help with protein purification, U Neef for insect cell maintenance, A Sawicka for providing cDNA, and SM Vos for valuable input and critical reading of the manuscript. We thank C Dienemann and U Steuerwald for support with electron microscopy. PC was supported by the Deutsche Forschungsgemeinschaft (SFB1064, SFB860), the European Research Council Advanced Investigator Grant TRANSREGULON (grant agreement No. 693023), and the Volkswagen Foundation.

Senior Editor

  1. Jessica K Tyler, Weill Cornell Medicine, United States

Reviewing Editor

  1. Geeta J Narlikar, University of California, San Francisco, United States

Publication history

  1. Received: March 10, 2020
  2. Accepted: June 15, 2020
  3. Accepted Manuscript published: June 16, 2020 (version 1)
  4. Version of Record published: July 6, 2020 (version 2)

Copyright

© 2020, Farnung et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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