Research Article

Structural insights into the histone H1-nucleosome complex

Bing-Rui Zhou, Hanqiao Feng, Hidenori Kato, Liang Dai, Yuedong Yang, Yaoqi Zhou, and Yawen Bai
PNAS November 26, 2013 110 (48) 19390-19395; https://doi.org/10.1073/pnas.1314905110

  1. Edited by S. Walter Englander, The University of Pennsylvania, Philadelphia, PA, and approved October 22, 2013 (received for review August 6, 2013)

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Significance

Linker H1 histones control the accessibility of linker DNA between two neighbor nucleosomes to DNA-binding proteins and regulate chromatin folding. We investigated the structure of the H1–nucleosome complex through a combination of multidimensional nuclear magnetic resonance spectroscopy, site-directed mutagenesis-isothermal-titration calorimetry and computational design/modeling. The results lead to a unique structural model for the globular domain of H1 in complex with the nucleosome that contains residue-level information and have implications for the dynamics of chromatin in vivo. In addition, our approach will be useful for testing the hypothesis that the globular domain of H1 variants might have distinct binding geometries within the nucleosome, and thereby contribute to the heterogeneity of chromatin structure.

Abstract

Linker H1 histones facilitate formation of higher-order chromatin structures and play important roles in various cell functions. Despite several decades of effort, the structural basis of how H1 interacts with the nucleosome remains elusive. Here, we investigated Drosophila H1 in complex with the nucleosome, using solution nuclear magnetic resonance spectroscopy and other biophysical methods. We found that the globular domain of H1 bridges the nucleosome core and one 10-base pair linker DNA asymmetrically, with its α3 helix facing the nucleosomal DNA near the dyad axis. Two short regions in the C-terminal tail of H1 and the C-terminal tail of one of the two H2A histones are also involved in the formation of the H1–nucleosome complex. Our results lead to a residue-specific structural model for the globular domain of the Drosophila H1 in complex with the nucleosome, which is different from all previous experiment-based models and has implications for chromatin dynamics in vivo.

Eukaryotic genomic DNA is packaged into chromatin through association with positively charged histones to form the nucleosome, the structural unit of chromatin (1⇓–3). The nucleosome core consists of an octamer of histones with two copies of H2A, H2B, H3, and H4, around which ∼146 bp of DNA winds in ∼1.65 left-handed superhelical turns (4). At this level of the DNA packaging, chromatin resembles a beads-on-a-string structure, with the nucleosome core as the beads and the linker DNA between them as the strings (5). At the next level of DNA packaging, H1 histones bind to the linker DNA and the nucleosome to further condense the chromatin structure (6, 7). H1-mediated chromatin condensation plays important roles in cellular functions such as mitotic chromosome architecture and segregation (8), muscle differentiation (9), and regulation of gene expression (10, 11).

Linker H1 histones typically are ∼200 amino acid residues in length, with a short N-terminal region, followed by a ∼70–80-amino acid structured globular domain (gH1) and a ∼100-amino acid unstructured C-terminal domain that is highly enriched in Lys residues. H1 stabilizes the nucleosome and facilitates folding of nucleosome arrays into higher-order structures (12⇓⇓–15). gH1 alone confers the same protection from micrococcal nuclease digestion to the nucleosome as the full-length H1 does (16). The N-terminal region of H1 is not important for nucleosome binding (16, 17), whereas the C terminus is required for H1 binding to chromatin in vivo (18, 19) and for the formation of a stem structure of linker DNA in vitro (17, 20, 21).

The globular domain structures of avian H5 (22) and budding yeast Hho1 (23), which are both H1 homologs, have been determined at atomic resolution and show similar structures. In addition, numerous studies have indicated that gH1/gH5 binds around the dyad region of the nucleosome (14, 24), leading to many conflicting structural models for how the globular domain of H1/H5 binds to the nucleosome (SI Appendix, Fig. S1) (24⇓–26). These models are divided into two major classes, symmetric and asymmetric, on the basis of the location of gH1/gH5 in the nucleosome. In the symmetric class, gH1/gH5 binds to the nucleosomal DNA at the dyad and interacts with both linker DNAs (16, 17, 27, 28). In the asymmetric class, gH1/gH5 binds to the nucleosomal DNA in the vicinity of the dyad axis and to 10 bp (27, 29⇓⇓–32) or 20 bp (19, 29, 33, 34) of one linker DNA, or is located inside the DNA gyres, where it interacts with histone H2A (35). In addition, Zhou and colleagues also characterized the orientation of gH5 in the gH5-nucleosome complex (29). The use of nonuniquely positioned nucleosomes and indirect methods may have contributed to the differences in these models (SI Appendix, Fig. S1).

Multidimensional nuclear magnetic resonance (NMR), and in particular methyl-based NMR, provides a direct approach to the structural characterization of macromolecular complexes (36, 37). We have previously assigned chemical shifts of the methyl groups of the side chains of residues Ile, Leu, and Val in the core histones (38) and the backbone amides in the disordered histone tails (39), which provide the fingerprints for investigating the interactions between H1 and the nucleosome. Here, we used NMR, along with several other methods, to determine the location and orientation of the globular domain of a stable mutant of Drosophila H1 on a well-positioned nucleosome.

Results

Histone Regions Involved in the Formation of the H1–Nucleosome Complex.

To identify the nucleosome-binding regions of H1, we first used a gel shift assay to examine the binding of several H1 fragments, which contain gH1 and various lengths of C-terminal regions, to the nucleosome centered on 167 bp DNA with the Widom “601” sequence (40), which can uniquely position the nucleosome. To be consistent with later NMR experiments, we used a stable mutant of H1 in which four residues in the hydrophobic core of gH1 are replaced with the corresponding residues in gH5 (22) (Fig. 1 A and B and SI Appendix, Fig. S2). This stable H1 mutant is necessary to allow NMR experiments to be conducted at the high temperature (35 °C) required to observe the methyl groups in the H1-nucleosome complex (38). In this study, H1 refers to this stable mutant unless specified otherwise. We found that all fragments of H1 shift the position of the nucleosome in the agarose gel (Fig. 1C). In particular, the fragment containing residues 37–211 shifted the nucleosome as efficiently as H137–256. Using isothermal titration calorimetry (ITC), we further showed that H11–256, H137–211, and the wild type full-length H11–256 have similar binding affinities for the nucleosome (SI Appendix, Fig. S3 and Table S1). Therefore, to avoid signal overlaps in NMR spectra (SI Appendix, Fig. S4A), we chose H137–211 for subsequent NMR studies. Using 15N/13C-labeled H137–211, we assigned the observable residues of H137–211 in complex with the nucleosome (SI Appendix, Fig. S4B). The unobserved residues of H137–211, which are presumably folded into the nucleosome, include gH1 (45–118), residues 119ASAKKEK125 immediately following the globular domain, and the Lys-rich region 164KKPKAKKAVAT174 in the middle of the C-terminal tail (Fig. 1 D and E). We further demonstrated that H137–211 and the wild-type full-length H11–256 used the same regions to bind to the nucleosome (SI Appendix, Fig. S4C). Similar results were observed when the nucleosome with 208 bp DNA was used (SI Appendix, Fig. S4D), indicating that H1 binds within the 10 bp of the linker DNAs that enter/exit the nucleosome core particle.

Fig. 1.
Fig. 1.

The globular domain and two discrete regions of linker histone H1 bind to the nucleosome. (A) Sequence of H1, highlighting the globular domain (cyan), the quadruple mutations (green and bold), and the C-terminal regions (red) that are involved in the binding of the nucleosome, and secondary structures. (B) The structural model of the globular domain, which is modeled using the B chain of the gH5 globular domain structure as the template (22). The mutated residues are shown as green sticks. The balls indicate the positions for spin labeling, except that positions 44 and 119 outside the gH1 are indicated with residues 45 and 118 in gH1, respectively. (C) Gel shift assay results for different fragments of H1. The nucleosome contains 167 bp DNA centered with the 601 sequence. (D) Overlay of 1H-15N HSQC spectra of H1 in free form (blue) and in complex with the nucleosome (black). In this experiment, the ratio of H137–211 to the nucleosome in the complex is ∼0.7. The disappearance of peaks indicates that many residues in H137–211 become folded on binding of the nucleosome. (E) Deviation of Cα chemical shifts of observable residues of H1 in complex with the nucleosome from random coil values.

To determine whether the histone core of the nucleosome interacts with H137–211, we compared the methyl spectra of the nucleosome histones in the absence or presence of H137–211. We found that H137–211 did not perturb the methyl spectra (SI Appendix, Fig. S5A), indicating that H137–211 and the histone core are not in direct contact. We then tested whether the C-terminal tails of H2A and the N-terminal tails of H3 interact with H137–211, as they are close to the linker DNA (Fig. 2A). The peak intensities/volumes of the C-terminal residues (119–122) of the 15N-labeled H2A decreased by about half in the 1H-15N heteronuclear single quantum correlation (HSQC) spectra on binding of H137–211 or the wild-type full-length H11–256, with little changes in their chemical shifts (Fig. 2 B–E), whereas those in 15N-labeled H3 and the N-terminal tails of 15N-labeled H2A remain unchanged in both peak intensities and chemical shifts (Fig. 2 B–E and SI Appendix, Fig. S5 B and C). Similar results were observed for H137–211 in complex with the nucleosome array, which has a longer linker DNA (SI Appendix, Fig. S6). These results indicate that one of the two disordered H2A C-terminal tails in the nucleosome and two short regions in the disordered C-terminal tails of the wild-type full-length H11–256 became folded upon binding of H1 to the nucleosome.

Fig. 2.
Fig. 2.

Asymmetric binding of H137–211 to the nucleosome. (A) Nucleosome structure, highlighting residues H2A T119 and H3 K37. The DNA structure is taken from the tetra-nucleosome structure with 10 bp DNA at both entry and exit regions (41): H2A is in orange, H3 is in blue, and H4 is in green. (B) Overlay of 1H-15N HSQC spectra of the nucleosome with 15N-labeled H2A in the absence (black) or the presence (orange) of mutant H137–211. (C) Same as in B except the wild-type full-length H11–256 (magenta) was used instead of mutant H137–211. Note that only strong peaks in the spectra are shown (SI Appendix, Fig. S7). In this experiment, the ratio of H137–211 to the nucleosome is 1.0. The dashed lines indicate that the volumes of the peaks in the boxes are integrated together. (D) Ratios of peak volumes of T119, K121, and the region that include K118, E120, and K122 of the H2A tails in the nucleosome in complex with mutant H137–211 to those in the free nucleosome. (E) Same as in D except that the wild-type full-length H11–256 was used instead of mutant H137–211. The asterisks in B–E indicate that the peaks in the dashed box were integrated together when the peak volumes were measured.

Determination of the Orientation of gH1 on the Nucleosome by PRE.

We next performed paramagnetic relaxation enhancement (PRE) experiments to determine the orientation of gH1 in the H137–211-nucleosome complex. In the PRE experiments, residues immediately outside the two ends of gH1 (P44 and A119) and on the surface (L60, A83, and K109) were chosen to minimize the perturbation to the gH1 structure (Fig. 1B). These residues were each individually mutated to Cys and linked to the paramagnetic compound [(S-(2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol-3-yl) methyl methanethiosulfonate) (MTSL) or ((S-methanethiosulfonylcysteaminyl)ethylenediamine-N,N,N′,N′-tetraacetic acid) (MTS-EDTA-Mn2+)] through a disulfide bond. Binding of paramagnetic spin-labeled H137–211 reduced the peak intensities of the observable backbone amide 1H-15N signals in the nucleosome in a manner dependent on distance from the paramagnetic center (42). The peak intensities of residues in the C-terminal tails of H2A were only weakly affected by MTSL but were strongly affected by MTS-EDTA-Mn2+ (Fig. 3 A–D and SI Appendix, Fig. S7), consistent with the anticipated PRE effects from the two types of spin labels (42). The intensity changes in the observable residue T119 in H2A, which is near the folded region of H2A, indicate it is closer to H137–211 residues A83 and K109 than to H137–211 residues L60, A119, and P44 (Fig. 3 A–D, Fig. 1B, and Fig. 2A). These results indicate that in the H137–211–nucleosome complex, the α3 helix of gH1, which is within the region 83–109, is closer to the nucleosome core than the gH1 N- and C-terminal border residues (P44 and A119) and loop 1 residue L60 (Fig. 1B).

Fig. 3.
Fig. 3.

Location and orientation of gH1 on the nucleosome determined by PRE effects. (A–D) PRE effects of spin labeling of H137–211 at positions 44, 60, 83, 109, and 119 on the amide protons of H2A C-terminal tail residues T119 and K122 and N-terminal R3 and K5 residues in the nucleosome. (E–G) Methyl-spectra of gH1 in complex with the nucleosome (red) and the nucleosomes with MTSL labels at H2A T119 (orange) and H3 K37 (blue), respectively. (H and I) Bar graphs showing PRE effects on the methyl groups of gH1 with MTSL label at H2A T119 (orange) and H3 K37 (blue), respectively.

Determination of the Location of gH1 on the Nucleosome by PRE.

We further used the methyl groups of Ile, Val, and Leu residues in gH1 as probes to determine the location of gH1 on the nucleosome. We first assigned most of the methyl groups in gH1 by combining three approaches (Fig. 3E and SI Appendix, Fig. S8): mutation of specific residues, comparison of the peak positions of the methyl groups of gH1 in the free form and in complex with the nucleosome, and spin-labeling effects on the methyl groups in gH1 in the absence or presence of the nucleosome. Using the assigned methyl groups, we examined the PRE effects on the methyl groups of gH1 with MTSL labeled at position T119 of H2A or at position K37 of H3 in the nucleosome (Fig. 2A). The residues at these positions are disordered in the nucleosome but are close to the nucleosome core, with relatively fixed locations. The spin labels have little effects on the chemical shifts of the methyl groups in gH1, indicating that they do not perturb the structure of the gH1–nucleosome complex (Fig. 3 E–G). The spin-label at position 119 of H2A has large PRE effects on the methyl groups of L97, V98, and V99 at the C-terminal region of the α3 helix of gH1, whereas the spin-label at position 37 of H3 strongly affected not only these methyl groups but also L60 in loop 1 and L103 and I104 in the C-terminal region of the β1 strand (Fig. 3 F–I).

Because there are two H3 K37 residues in the nucleosome structure that are separated by ∼65 Å (Fig. 2A), the observation that methyl groups in one region of gH1 are strongly affected by the MTSL labels at the H3 K37 sites further indicates that gH1 binds to the nucleosome asymmetrically. This is also consistent with the folding of only one of the two H2A C-terminal tails in the nucleosome on H137–211 binding (Fig. 2 B and C). Furthermore, as MTSL spin labels in gH1 have only small effects on the backbone amide NMR signals from the disordered H2A T119 (Fig. 3A), gH1 must be distant from this disordered H2A C-terminal tail in the H137–211–nucleosome complex. Therefore, the large PRE effects on some methyl groups of gH1 from MTSL labels at H2A T119 (Fig. 3H) indicate that gH1 is close to the folded H2A C-terminal tail in the H1–nucleosome complex.

Important H1 Residues for Nucleosome Binding.

We used site-directed mutagenesis and ITC to identify gH1 residues that are important for the binding of H137–211 to the nucleosome. Positively charged Lys residues on the surface of gH1 were each mutated to Ala, and the effects of the mutations on the dissociation equilibrium constant (KD) were measured by ITC (Fig. 4A and SI Appendix, Table S1). Mutation of each of the six residues (K58, K91, K95, K102, K107, and K116) showed larger effects (> a factor of 2.5) than others. These mutations are located on two distinct surfaces on nearly opposite sides of the gH1 structure (Fig. 4B). One surface includes K91 and K95 in the α3 helix. The other surface includes residues K58 at the C-terminal region of the α1 helix and residues K102, K107, and K116 in the two β strands. In addition, ITC experiments showed that gH1 binding to the nucleosome was ∼12 times weaker than H137–211 (SI Appendix, Fig. S9). These results are consistent with the NMR observation that residues 119ASAKKEK125 and 164KKPKAKKAVAT174 in the middle of the C-terminal tail of H11–256 are folded.

Fig. 4.
Fig. 4.

Effects of mutations in gH1 on the binding affinity of H137–211 to the nucleosome. (A) Effects of mutation of surface residues in gH1 on the binding affinity between H137–211 and the nucleosome. (B) Structural illustration of the distribution of the gH1 residues whose Ala mutations lead to a large decrease in binding affinity.

A Structural Model for the gH1–Nucleosome Complex.

Taking all the data together, our experimental results suggest that gH1 uses the two positively charged surfaces to bridge the nucleosome core and the linker DNA asymmetrically. GH1 is close to one of the two H2A C-terminal tails and one of the two H3 N-terminal tails, with its α3 helix facing the nucleosome core. Using these restraints and the HADDOCK program (43), we docked the gH1 onto the nucleosome by forcing the K91 and K95 residues in the α3 helix of gH1 to interact with the nucleosomal DNA near the dyad region and the K58, K102, K107, and K116 residues to interact with one of the nearby 10-bp linker DNAs (SI Appendix, Fig. S10). After initial rigid body docking and subsequent energy minimization that allows contacting residues to adopt different conformations, the calculated low-energy structures were clustered. We found that one of the clusters was most consistent with all of the experimental results (Fig. 5 A and B and SI Appendix, Fig. S10D), including the mutation effects on the binding affinity between H137–211 and the nucleosome (Fig. 5C), PRE effects on H2A T119 in the nucleosome by spin labels in gH1 (Fig. 5D), and PRE effects on the methyl groups of gH1 by spin labels at H2A T119 (Fig. 5E) and H3 K37 (Fig. 5F). In the model, although gH1 directly interacts with only one linker DNA, it is very close to the other linker DNA (Fig. 5B and SI Appendix, Fig. S10E). A slight shift of this linker DNA toward the nucleosome core could also lead to its interaction with gH1. Therefore, it is possible that gH1 may interact with both DNA linkers, one strongly and the other weakly (SI Appendix, Fig. S6A).

Fig. 5.
Fig. 5.

An asymmetrical structural model of the gH1-nucleosome complex. (A and B) Structural model of the gH1-nucleosome complex in surface and ribbon representations. The rectangular frame approximately shows the enlarged regions in C–F. (C) Illustration of the Lys residues that show a large increase in KD (> a factor of 2.5) when mutated to Ala. (D) Illustration of the disordered residue H2A_T119 and residues P44 (H45), L60, A83, K109, and A119 (S118) in gH1 in the gH1-nucleosome complex. (E) Illustration of the ordered residue H2A_T119 and the methyl groups in gH1. The methyl groups in red have an Ipara/Idia [Intensity (paramagnetic)/Intensity (diamagnetic)] of less than 0.3, whereas other methyl groups in cyan have larger values. (F) Illustration of one of the two H3_K37 residues and the methyl groups in gH1. The methyl groups in red have Ipara/Idia of less than 0.3, whereas methyl groups in cyan have larger values.

The C-Terminal Tail of Histone Variant H2A.Z Disfavors H1 Binding.

Earlier studies have shown that the nucleosome containing histone variant H2A.Z affects H1 binding (44), and the C-terminal tail of H2A.Z may play a role in the reduced binding affinity (45). The amino acid sequence of the very C-terminal region (119TEKKA123) of H2A is different from corresponding residues (123EETVQ127) of H2A.Z. To test the role of the C-terminal tail of H2A.Z in H1 binding, we measured the binding affinity of H137–211 to the nucleosome containing H2A with the last six residues replaced by the corresponding residues of H2A.Z. We found that the binding was reduced to an undetectable level by ITC (SI Appendix, Fig. S11), whereas deletion of the last five residues of H2A increased KD by a factor of ∼10. These results further confirm the earlier NMR observation that one of the C-terminal tails of H2A is involved in the formation of the H1–nucleosome complex.

Discussion

Our structural model of the gH1–nucleosome complex differs from all previous experiment-based models in either the location or orientation of gH1/H5 or the length of linker DNA involved. The observation that MTSL labels at H2A T119 or H3 K37 have large PRE effects on the methyl groups in only one region of gH1 clearly shows that the gH1 binds to the nucleosome asymmetrically. For a symmetric binding, large PRE effects on methyl groups of gH1 from spin labels at H3 K37 are not expected because they are separated by more than 20 Å (SI Appendix, Fig. S12A). It is interesting to note that Zhou and coworkers have also found that gH5 binds to the nucleosome complex asymmetrically, involving 10-bp linker DNA (SI Appendix, Fig. S1) (29). However, the orientation of gH5 in their model is opposite to that of gH1 in ours: The α3 helix of gH5 in their model binds to the linker DNA instead of the nucleosomal DNA, as in our model (SI Appendix, Fig. S12 B and C). One possibility for the differences among various models of the gH1/H5–nucleosome complex might be that there are multiple gH1/H5 binding modes for the nucleosome. Another possibility is that earlier models are derived from less-controlled systems, using indirect methods (SI Appendix, Fig. S1).

Among several computation-based models of the gH1/H5–nucleosome complex (30⇓–32, 46), two of them show similarities to our model in terms of both location and orientation of gH1 in the complex (30, 31). In addition, our model is consistent with several other experimental observations. For example, the two C-terminal tails of H2A in the Xenopus nucleosome cross-link to different positions of the DNA in the nucleosome on addition of H1 (47). Calf H1 is cross-linked to the C-terminal region of H2A (48). The location of gH1 in our model is very close to the disordered negatively charged C-terminal tail of HMGN2 in complex with the nucleosome, consistent with its role in inhibiting the binding of H1 to the nucleosome (38). Binding of H1 to the 10-bp linker DNA that neighbors the nucleosome core particle has been used to explain the preferential “out-of-phase” population of AT bases in the region (31, 49). The identification of two major discrete DNA-binding surfaces of gH1 in our mutation studies is in excellent agreement with earlier mutation studies of the histone H5 globular domain (50) and Mouse H10, using fluorescence recovery after photo-bleaching methods (19) (SI Appendix, Fig. S13). In addition, within the limit of the resolution, the tetra-nucleosome crystal structure can accommodate gH1 in our model (41) with only small clashes with the linker DNA that connects the neighboring nucleosomes (SI Appendix, Fig. S14), suggesting that a small structural rearrangement of the linker DNA could allow H1 to condense chromatin to the two-start zigzag nucleosome high-order structure (7, 51).

Our finding that two discrete regions in the C-terminal tail of H1, 119ASAKKEK125 and 164KKPKAKKAVAT174, are involved in nucleosome binding is consistent with the earlier experimental observations that the C-terminal tail of H1 plays an important role in nucleosome binding and chromatin structure condensation (18, 20, 21). In particular, our results support the conclusions that specific regions in the C-terminal tail of H1, rather than the distribution of positively charged residues, are responsible for interaction with linker DNA (17, 18, 52, 53). In the case of human H1.5, it has been shown that the seven residues following the globular domain, 121PKAKKAG127, are necessary for the formation of the linker DNA stem structure (17). Six of these seven residues are conserved in the 164KKPKAKKAVAT174 region of the C-terminal tail of Drosophila H1. Our results show that NMR provides a powerful tool for identifying specific nucleosome-binding regions in the C-terminal tail of H1.

It has been shown that the H2A.Z-containing nucleosome is typically present at regions flanking the nucleosome-free region (54), near the DNA double-strand break (55), and at the boundary of euchromatin and heterochromatin, which prevents spread of heterochromatin regions (56). In general, these chromatin regions are less condensed and more dynamic. Our finding that the very last several residues at the H2A.Z C-terminal tail inhibit the binding of H1 to the H2A.Z nucleosome provides a possible mechanistic explanation for the dynamic features of the H2A.Z-enriched chromatin regions.

Finally, our approach provides an experimental tool for testing the hypothesis that the globular domains of individual H1 variants might have distinct binding geometries within the nucleosome that contribute to the heterogeneity of chromatin structure (9, 19, 57).

Materials and Methods

Design of a Stable Mutant of gH1 and Preparation of Samples.

The gH1 structure model was built using homology modeling and the structure of gH5. Protein design programs were used to select the mutation (V53I, S56A, C81V, and A97L) that stabilizes gH1 structure. The Drosophila H1 gene was synthesized and inserted into the pET42b vector. Mutations were made using the QuikChange kit. Proteins were expressed in Escherichia coli and purified using chromatography. MTSL or MTS-EDTA was linked to the protein, with the target site mutated to Cys. Nucleosomes were reconstituted by stepwise salt dialysis in the absence of reducing agent, followed by HPLC to remove free DNA and immature nucleosomes (SI Appendix, Materials and Methods and SI Appendix, Fig. S15).

NMR, PRE, and ITC Experiments.

Isotope-labeled H1 mixed with the nucleosome, or vice versa. The spin-labeled H1 was mixed with nucleosomes containing 15N-labeled histones or vice versa. 1H-15N HSQC or 1H-13C heteronuclear multiple quantum correlation (HMQC) spectra were recorded. The NMR peak intensities or volumes were measured. ITC experiments were performed on a VP–ITC microcalorimeter (Microcal). The dissociation constant (KD) and the stoichiometry of binding (n) were determined by fitting the observed binding curves to the model with independent n and KD values (SI Appendix, Materials and Methods).

Docking Calculation.

We docked gH1 to the 167-bp DNA obtained from the tetra-nucleosome structure using HADDOCK program. Residues Lys91 and Lys95 in gH1 were forced to interact with the nucleosomal DNA near the dyad; the K58, K102, K107, and K106 residues were forced to interact with the nearby linker DNA. A cluster of structures that is most consistent with the experimental data were selected as the final model (SI Appendix, Materials and Methods).

Acknowledgments

We thank Dr. Jemima Barrowman for editing the manuscript. This work is supported by the intramural research program of National Cancer Institute and National Institutes of Health (NIH) (to Y.B.) and by the National Institute of General Medical Sciences of the NIH (R01GM085003, to Y.Z.).

Footnotes

  • ↵1Present address: Institute for Glycomics and School of Informatics and Communication Technology, Griffith University, Southport, QLD 4222, Australia.

  • ↵2To whom correspondence may be addressed. E-mail: yawen{at}helix.nih.gov.

  • Author contributions: B.-R.Z. and Y.B. designed research; B.-R.Z., H.F., L.D., Y.Y., and Y.Z. performed research; H.K. contributed new reagents/analytic tools; B.-R.Z., H.F., L.D., Y.Y., Y.Z., and Y.B. analyzed data; and B.-R.Z. and Y.B. wrote the paper.

  • The authors declare no conflict of interest.

  • This article is a PNAS Direct Submission.

  • This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1314905110/-/DCSupplemental.

References

  1. ↵
    1. Kornberg RD
    (1974) Chromatin structure: A repeating unit of histones and DNA. Science 184(4139):868–871.
    OpenUrlFREE Full Text
  2. ↵
    1. Kornberg RD,
    2. Thomas JO
    (1974) Chromatin structure; oligomers of the histones. Science 184(4139):865–868.
    OpenUrlFREE Full Text
  3. ↵
    1. Khorasanizadeh S
    (2004) The nucleosome: From genomic organization to genomic regulation. Cell 116(2):259–272.
    OpenUrlCrossRefPubMed
  4. ↵
    1. Luger K,
    2. Mäder AW,
    3. Richmond RK,
    4. Sargent DF,
    5. Richmond TJ
    (1997) Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389(6648):251–260.
    OpenUrlCrossRefPubMed
  5. ↵
    1. Felsenfeld G,
    2. Groudine M
    (2003) Controlling the double helix. Nature 421(6921):448–453.
    OpenUrlCrossRefPubMed
  6. ↵
    1. Thoma F,
    2. Koller T
    (1977) Influence of histone H1 on chromatin structure. Cell 12(1):101–107.
    OpenUrlCrossRefPubMed
  7. ↵
    1. Robinson PJ,
    2. Rhodes D
    (2006) Structure of the ‘30 nm’ chromatin fibre: A key role for the linker histone. Curr Opin Struct Biol 16(3):336–343.
    OpenUrlCrossRefPubMed
  8. ↵
    1. Maresca TJ,
    2. Freedman BS,
    3. Heald R
    (2005) Histone H1 is essential for mitotic chromosome architecture and segregation in Xenopus laevis egg extracts. J Cell Biol 169(6):859–869.
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Lee H,
    2. Habas R,
    3. Abate-Shen C
    (2004) MSX1 cooperates with histone H1b for inhibition of transcription and myogenesis. Science 304(5677):1675–1678.
    OpenUrlAbstract/FREE Full Text
  10. ↵
    1. Fan Y,
    2. et al.
    (2005) Histone H1 depletion in mammals alters global chromatin structure but causes specific changes in gene regulation. Cell 123(7):1199–1212.
    OpenUrlCrossRefPubMed
  11. ↵
    1. Shen X,
    2. Gorovsky MA
    (1996) Linker histone H1 regulates specific gene expression but not global transcription in vivo. Cell 86(3):475–483.
    OpenUrlCrossRefPubMed
  12. ↵
    1. Thoma F,
    2. Koller T,
    3. Klug A
    (1979) Involvement of histone H1 in the organization of the nucleosome and of the salt-dependent superstructures of chromatin. J Cell Biol 83(2 Pt 1):403–427.
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Singer DS,
    2. Singer MF
    (1976) Studies on the interaction of H1 histone with superhelical DNA: Characterization of the recognition and binding regions of H1 histones. Nucleic Acids Res 3(10):2531–2547.
    OpenUrlAbstract/FREE Full Text
  14. ↵
    1. Simpson RT
    (1978) Structure of the chromatosome, a chromatin particle containing 160 base pairs of DNA and all the histones. Biochemistry 17(25):5524–5531.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Noll M,
    2. Kornberg RD
    (1977) Action of micrococcal nuclease on chromatin and the location of histone H1. J Mol Biol 109(3):393–404.
    OpenUrlPubMed
  16. ↵
    1. Allan J,
    2. Hartman PG,
    3. Crane-Robinson C,
    4. Aviles FX
    (1980) The structure of histone H1 and its location in chromatin. Nature 288(5792):675–679.
    OpenUrlCrossRefPubMed
  17. ↵
    1. Syed SH,
    2. et al.
    (2010) Single-base resolution mapping of H1-nucleosome interactions and 3D organization of the nucleosome. Proc Natl Acad Sci USA 107(21):9620–9625.
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Hendzel MJ,
    2. Lever MA,
    3. Crawford E,
    4. Th’ng JP
    (2004) The C-terminal domain is the primary determinant of histone H1 binding to chromatin in vivo. J Biol Chem 279(19):20028–20034.
    OpenUrlAbstract/FREE Full Text
  19. ↵
    1. Brown DT,
    2. Izard T,
    3. Misteli T
    (2006) Mapping the interaction surface of linker histone H1(0) with the nucleosome of native chromatin in vivo. Nat Struct Mol Biol 13(3):250–255.
    OpenUrlCrossRefPubMed
  20. ↵
    1. Hamiche A,
    2. Schultz P,
    3. Ramakrishnan V,
    4. Oudet P,
    5. Prunell A
    (1996) Linker histone-dependent DNA structure in linear mononucleosomes. J Mol Biol 257(1):30–42.
    OpenUrlCrossRefPubMed
  21. ↵
    1. Bednar J,
    2. et al.
    (1998) Nucleosomes, linker DNA, and linker histone form a unique structural motif that directs the higher-order folding and compaction of chromatin. Proc Natl Acad Sci USA 95(24):14173–14178.
    OpenUrlAbstract/FREE Full Text
  22. ↵
    1. Ramakrishnan V,
    2. Finch JT,
    3. Graziano V,
    4. Lee PL,
    5. Sweet RM
    (1993) Crystal structure of globular domain of histone H5 and its implications for nucleosome binding. Nature 362(6417):219–223.
    OpenUrlCrossRefPubMed
  23. ↵
    1. Ali T,
    2. Coles P,
    3. Stevens TJ,
    4. Stott K,
    5. Thomas JO
    (2004) Two homologous domains of similar structure but different stability in the yeast linker histone, Hho1p. J Mol Biol 338(1):139–148.
    OpenUrlCrossRefPubMed
  24. ↵
    1. Thomas JO
    (1999) Histone H1: Location and role. Curr Opin Cell Biol 11(3):312–317.
    OpenUrlCrossRefPubMed
  25. ↵
    1. Vignali M,
    2. Workman JL
    (1998) Location and function of linker histones. Nat Struct Biol 5(12):1025–1028.
    OpenUrlCrossRefPubMed
  26. ↵
    1. Crane-Robinson C
    (1997) Where is the globular domain of linker histone located on the nucleosome? Trends Biochem Sci 22(3):75–77.
    OpenUrlCrossRefPubMed
  27. ↵
    1. Fan L,
    2. Roberts VA
    (2006) Complex of linker histone H5 with the nucleosome and its implications for chromatin packing. Proc Natl Acad Sci USA 103(22):8384–8389.
    OpenUrlAbstract/FREE Full Text
  28. ↵
    1. Meyer S,
    2. et al.
    (2011) From crystal and NMR structures, footprints and cryo-electron-micrographs to large and soft structures: Nanoscale modeling of the nucleosomal stem. Nucleic Acids Res 39(21):9139–9154.
    OpenUrlAbstract/FREE Full Text
  29. ↵
    1. Zhou YB,
    2. Gerchman SE,
    3. Ramakrishnan V,
    4. Travers A,
    5. Muyldermans S
    (1998) Position and orientation of the globular domain of linker histone H5 on the nucleosome. Nature 395(6700):402–405.
    OpenUrlCrossRefPubMed
  30. ↵
    1. Pachov GV,
    2. Gabdoulline RR,
    3. Wade RC
    (2011) On the structure and dynamics of the complex of the nucleosome and the linker histone. Nucleic Acids Res 39(12):5255–5263.
    OpenUrlAbstract/FREE Full Text
  31. ↵
    1. Cui F,
    2. Zhurkin VB
    (2009) Distinctive sequence patterns in metazoan and yeast nucleosomes: Implications for linker histone binding to AT-rich and methylated DNA. Nucleic Acids Res 37(9):2818–2829.
    OpenUrlAbstract/FREE Full Text
  32. ↵
    1. Bharath MM,
    2. Chandra NR,
    3. Rao MR
    (2003) Molecular modeling of the chromatosome particle. Nucleic Acids Res 31(14):4264–4274.
    OpenUrlAbstract/FREE Full Text
  33. ↵
    1. Wong J,
    2. Li Q,
    3. Levi BZ,
    4. Shi YB,
    5. Wolffe AP
    (1997) Structural and functional features of a specific nucleosome containing a recognition element for the thyroid hormone receptor. EMBO J 16(23):7130–7145.
    OpenUrlAbstract/FREE Full Text
  34. ↵
    1. An W,
    2. Leuba SH,
    3. van Holde K,
    4. Zlatanova J
    (1998) Linker histone protects linker DNA on only one side of the core particle and in a sequence-dependent manner. Proc Natl Acad Sci USA 95(7):3396–3401.
    OpenUrlAbstract/FREE Full Text
  35. ↵
    1. Pruss D,
    2. et al.
    (1996) An asymmetric model for the nucleosome: A binding site for linker histones inside the DNA gyres. Science 274(5287):614–617.
    OpenUrlAbstract/FREE Full Text
  36. ↵
    1. Bax A
    (2011) Triple resonance three-dimensional protein NMR: Before it became a black box. J Magn Reson 213(2):442–445.
    OpenUrlCrossRefPubMed
  37. ↵
    1. Tugarinov V,
    2. Hwang PM,
    3. Kay LE
    (2004) Nuclear magnetic resonance spectroscopy of high-molecular-weight proteins. Annu Rev Biochem 73:107–146.
    OpenUrlCrossRefPubMed
  38. ↵
    1. Kato H,
    2. et al.
    (2011) Architecture of the high mobility group nucleosomal protein 2-nucleosome complex as revealed by methyl-based NMR. Proc Natl Acad Sci USA 108(30):12283–12288.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    1. Zhou BR,
    2. et al.
    (2012) Histone H4 K16Q mutation, an acetylation mimic, causes structural disorder of its N-terminal basic patch in the nucleosome. J Mol Biol 421(1):30–37.
    OpenUrlCrossRefPubMed
  40. ↵
    1. Thåström A,
    2. Bingham LM,
    3. Widom J
    (2004) Nucleosomal locations of dominant DNA sequence motifs for histone-DNA interactions and nucleosome positioning. J Mol Biol 338(4):695–709.
    OpenUrlCrossRefPubMed
  41. ↵
    1. Clore GM,
    2. Tang C,
    3. Iwahara J
    (2007) Elucidating transient macromolecular interactions using paramagnetic relaxation enhancement. Curr Opin Struct Biol 17(5):603–616.
    OpenUrlCrossRefPubMed
  42. ↵
    1. Dominguez C,
    2. Boelens R,
    3. Bonvin AM
    (2003) HADDOCK: A protein-protein docking approach based on biochemical or biophysical information. J Am Chem Soc 125(7):1731–1737.
    OpenUrlCrossRefPubMed
  43. ↵
    1. Thakar A,
    2. et al.
    (2009) H2A.Z and H3.3 histone variants affect nucleosome structure: Biochemical and biophysical studies. Biochemistry 48(46):10852–10857.
    OpenUrlCrossRefPubMed
  44. ↵
    1. Vogler C,
    2. et al.
    (2010) Histone H2A C-terminus regulates chromatin dynamics, remodeling, and histone H1 binding. PLoS Genet 6(12):e1001234.
    OpenUrlCrossRefPubMed
  45. ↵
    1. Wong H,
    2. Victor JM,
    3. Mozziconacci J
    (2007) An all-atom model of the chromatin fiber containing linker histones reveals a versatile structure tuned by the nucleosomal repeat length. PLoS ONE 2(9):e877.
    OpenUrlCrossRefPubMed
  46. ↵
    1. Lee KM,
    2. Hayes JJ
    (1998) Linker DNA and H1-dependent reorganization of histone-DNA interactions within the nucleosome. Biochemistry 37(24):8622–8628.
    OpenUrlCrossRefPubMed
  47. ↵
    1. Boulikas T,
    2. Wiseman JM,
    3. Garrard WT
    (1980) Points of contact between histone H1 and the histone octamer. Proc Natl Acad Sci USA 77(1):127–131.
    OpenUrlAbstract/FREE Full Text
  48. ↵
    1. Travers AA,
    2. Muyldermans SV
    (1996) A DNA sequence for positioning chromatosomes. J Mol Biol 257(3):486–491.
    OpenUrlCrossRefPubMed
  49. ↵
    1. Goytisolo FA,
    2. et al.
    (1996) Identification of two DNA-binding sites on the globular domain of histone H5. EMBO J 15(13):3421–3429.
    OpenUrlPubMed
  50. ↵
    1. Schalch T,
    2. Duda S,
    3. Sargent DF,
    4. Richmond TJ
    (2005) X-ray structure of a tetranucleosome and its implications for the chromatin fibre. Nature 436(7047):138–141.
    OpenUrlCrossRefPubMed
  51. ↵
    1. Dorigo B,
    2. et al.
    (2004) Nucleosome arrays reveal the two-start organization of the chromatin fiber. Science 306(5701):1571–1573.
    OpenUrlAbstract/FREE Full Text
  52. ↵
    1. Lu X,
    2. Hansen JC
    (2004) Identification of specific functional subdomains within the linker histone H10 C-terminal domain. J Biol Chem 279(10):8701–8707.
    OpenUrlAbstract/FREE Full Text
  53. ↵
    1. Fang H,
    2. Clark DJ,
    3. Hayes JJ
    (2012) DNA and nucleosomes direct distinct folding of a linker histone H1 C-terminal domain. Nucleic Acids Res 40(4):1475–1484.
    OpenUrlAbstract/FREE Full Text
  54. ↵
    1. Cairns BR
    (2009) The logic of chromatin architecture and remodelling at promoters. Nature 461(7261):193–198.
    OpenUrlCrossRefPubMed
  55. ↵
    1. Morrison AJ,
    2. Shen X
    (2009) Chromatin remodelling beyond transcription: The INO80 and SWR1 complexes. Nat Rev Mol Cell Biol 10(6):373–384.
    OpenUrlCrossRefPubMed
  56. ↵
    1. Zofall M,
    2. et al.
    (2009) Histone H2A.Z cooperates with RNAi and heterochromatin factors to suppress antisense RNAs. Nature 461(7262):419–422.
    OpenUrlCrossRefPubMed
  57. ↵
    1. Alami R,
    2. et al.
    (2003) Mammalian linker-histone subtypes differentially affect gene expression in vivo. Proc Natl Acad Sci USA 100(10):5920–5925.
    OpenUrlAbstract/FREE Full Text
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Structural model of the H1-nucleosome complex
Bing-Rui Zhou, Hanqiao Feng, Hidenori Kato, Liang Dai, Yuedong Yang, Yaoqi Zhou, Yawen Bai
Proceedings of the National Academy of Sciences Nov 2013, 110 (48) 19390-19395; DOI: 10.1073/pnas.1314905110
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