Nucleosomes are the unitary structures of chromosome folding, and their arrangements are intimately coupled to the regulation of genome activities. Conventionally, structural analyses using electron microscopy and X-ray crystallography have been used to study such spatial nucleosome arrangements. In contrast, recent improvements in the resolution of sequencing-based methods allowed investigation of nucleosome arrangements separately at each genomic locus, enabling exploration of gene-dependent regulation mechanisms. Here, we review recent studies on nucleosome folding in chromosomes from these two methodological perspectives: conventional structural analyses and DNA sequencing, and discuss their implications for future research.

Introduction

Conventional structural analyses using electron microscopy (EM) and X-ray crystallography provided the early foundations to our understanding of chromatin structure (Figure 1). An outstanding early finding was the observation of nucleosome particles in chromosomes with EM in 1974, revealing that genomic DNA is compacted in a beads-on-a-string manner in the cell [13]. Another key achievement was the determination of the atomic structure of the nucleosome with X-ray crystallography, elucidating how 146 bp of DNA are compacted into a superhelix around an octamer of histone proteins consisting of two dimers of histones H2A and H2B and a tetramer of H3 and H4 [4]. Furthermore, important findings have also been made with EM regarding how nucleosomes are folded in chromatin [5], and the folding manner of nucleosomes is currently actively debated, with the aim of understanding principles of genome packaging and regulation at the molecular level. More recently, a striking alternative strategy to reveal chromosome structures has emerged with methods based on next-generation sequencing (Figure 1). These methods, represented by high-throughput chromosome conformation capture (Hi-C) techniques, have the unique feature of being able to provide structural information at each genomic locus because their readout is tied to DNA sequencing [6,7]. Interestingly, recent developments on this technique allowed interrogation of structures at a resolution up to the nucleosome level [8,9], which importantly now permits arguments on nucleosome arrangements from a similar viewpoint to the prior imaging studies. In this review, we summarize recent work investigating nucleosome folding from these two perspectives: conventional structural analyses and the newer sequencing-based approaches, and the types of local chromatin folding models they predict. We then highlight the critical next steps towards genome-wide maps of chromatin folding, which will facilitate a deeper understanding of the structure–function relationships operating across genomes.

Timeline of studies on nucleosome folding structure.

Figure 1.
Timeline of studies on nucleosome folding structure.

Schematic timeline showing landmark discoveries in the chromatin structure field over the last four decades. Top panels show key structures observed in native chromatin, while bottom panels show structures from in vitro reconstituted chromatin. Coloured circles depict the methodology of structural analyses including electron microscopy (blue), X-ray crystallography (green), computational simulation (yellow), sequencing (red), and optical microscopy (purple). Landmarks include ‘Beads-on-a-string’ pattern [1], coining of the term ‘nucleosome’ [10], one-start solenoid models [5,11] (EMDB code, 1782), two-start helical ribbon models [12,13], two-start crossed-linker model [14], nucleosome structure [4] (PDB code: 1KX5), two-start zig-zag model [15], tetranucleosome folding unit [16] (PDB code: 1ZBB), interdigitated one-start solenoid model [17], polymer melt model [18], fractal globule model [7], heteromorphic model [19], two-start zig-zag model with tetranucleosomal unit [20] (EMDB code: 2602), nucleosome clutch [21], gene crumple [8], and heterogenous 5- to 24-nm fibre [22]. Images are adapted from refs [7,15,2224].

Figure 1.
Timeline of studies on nucleosome folding structure.

Schematic timeline showing landmark discoveries in the chromatin structure field over the last four decades. Top panels show key structures observed in native chromatin, while bottom panels show structures from in vitro reconstituted chromatin. Coloured circles depict the methodology of structural analyses including electron microscopy (blue), X-ray crystallography (green), computational simulation (yellow), sequencing (red), and optical microscopy (purple). Landmarks include ‘Beads-on-a-string’ pattern [1], coining of the term ‘nucleosome’ [10], one-start solenoid models [5,11] (EMDB code, 1782), two-start helical ribbon models [12,13], two-start crossed-linker model [14], nucleosome structure [4] (PDB code: 1KX5), two-start zig-zag model [15], tetranucleosome folding unit [16] (PDB code: 1ZBB), interdigitated one-start solenoid model [17], polymer melt model [18], fractal globule model [7], heteromorphic model [19], two-start zig-zag model with tetranucleosomal unit [20] (EMDB code: 2602), nucleosome clutch [21], gene crumple [8], and heterogenous 5- to 24-nm fibre [22]. Images are adapted from refs [7,15,2224].

Conventional structural analyses reveal basic nucleosome arrangements

Understanding basic structural rules governing nucleosome folding in chromatin has been one of the major topics in the field. At an early stage, this was often investigated with EM imaging of chromatin within or isolated from the cell. The imaging data commonly showed a fibre structure condensed with a diameter of ∼30 nm, but with typical nucleosome arrangements varying among studies depending on the different experimental conditions. This variability led to respective proposals of different nucleosome folding models, such as the one-start solenoid model [5,25], two-start helical ribbon model [12,13], and two-start crossed-linker model [14,26].

As one approach to tackle this variability, structural analyses were performed using in vitro reconstituted nucleosomal arrays, in which purified histone proteins are periodically bound to a defined DNA sequence via nucleosome positioning elements such as the popular 601 sequences [27]. Despite this simplified experimental system, the basic nucleosome folding structure remained controversial, with two independent EM measurements of reconstituted arrays proposing the solenoid [17] and zig-zag [15] model, respectively. Subsequent X-ray crystallography and EM analyses on reconstituted arrays revealed the existence of a tetranucleosome folding unit more consistent with the zig-zag model [16,20]. A single-molecule force spectroscopic analysis on reconstituted arrays using magnetic tweezers also supported a tetranucleosomal structure [28].

Recent technological developments suggested further different models of nucleosome folding in native chromatin. A cryo-EM analysis on cellular chromatin, which is expected to provide a close-to-native chromatin structure by being frozen, indicated that neither the 30-nm fibre structure nor a regular nucleosome folding pattern exists in cells [18]. Supporting this, small-angle X-ray scattering studies indicated an absence of 30-nm fibres in both mitotic phase [29] and interphase [30] chromatin. In addition, an electron spectroscopic imaging study suggested that both open and closed chromatin regions consist of fibres no wider than 10 nm [31], which is close to the size of the nucleosome particle itself. These findings raised doubts about the existence of 30-nm fibres in cells [32,33], and led to the proposal of the polymer melt model, in which nucleosome folding is highly disordered and interdigitated [32].

Meanwhile, a dependence of nucleosome folding structure on the type of genomic region has also been argued. A super-resolution optical imaging study using the STORM technique to image immunolabelled nucleosomes suggested that the chromatin fibre is formed by heterogeneous groups of nucleosomes, termed ‘clutches’ [21]. This study also suggested that heterochromatin regions consist of larger and denser clutches, while open chromatin regions are associated with smaller and less dense clutches. Another more recent live-cell super-resolution study showed that nucleosomes come together in compact domains with a peak diameter of ∼160 nm [34]. These domains contained cohesin proteins and deacetylated nucleosomes and existed throughout the cell cycle even during mitosis. Furthermore, a recent method based on EM with tomographic imaging, called ChromEMT, revealed that chromatin forms flexible, structurally heterogeneous chains with ∼5–24 nm in diameter [22], which are packed together at different densities but without an apparent higher-order structure in both interphase and mitosis. This is consistent with earlier results from a method combining Monte Carlo simulation with EM-assisted nucleosome interaction capture, termed EMANIC, that suggested a heteromorphic chromatin fibre folding [19,35], where the fibre formed either a zig-zag or solenoid conformation.

As the above findings demonstrate, there are currently large discrepancies regarding basic nucleosome folding structures. These discrepancies probably arise from the different sample conditions required for each measurement method, such as the necessity of in vitro reconstruction of chromatin with purified proteins for X-ray crystallography and cryo-EM, and the need for genomic fusions of fluorescent proteins for fluorescence imaging studies (Table 1). In addition, these discrepancies are expected to be caused by structural heterogeneity across each genomic locus resulting from the DNA sequence and epigenetic factors. This issue highlights the requirement for structural analyses of specific endogenous loci using sequencing-based methods (see below).

Table 1
Comparison of representative methods for nucleosome folding structure
Method Sample Preparation Chemical fixation Probing method Observed structure Reference 
EM Native chromatin (isolated) Air-drying, freeze-etching Yes, no Negative stain, shadowing One-start solenoid model Finch and Klug [5
Native chromatin (isolated) Air-drying Yes Negative stain Two-start helical ribbon model Worcel et al. [12
Native chromatin (isolated) Air-drying, freeze-drying Yes, no Negative stain, shadowing, non-stained Two-start helical ribbon model Woodcock et al. [13
Native chromatin (isolated) Freeze-drying Yes Negative stain, shadowing, non-stained Two-start crossed-linker model Williams et al. [14
Reconstituted nucleosome array Air-drying Yes Negative stain Two-start model Dorigo et al. [15
Reconstituted nucleosome array Air-drying Yes Negative stain Interdigitated one-start solenoid model Robinson et al. [17
EM and in silico Reconstituted nucleosome array Air-drying Yes Positive-staining, shadowing Heteromorphic chromatin fibres Grigoryev et al. [19
Cryo-EM Native chromatin (isolated) Cryo-fixation No – Stem conformation of linker DNA segments formed by linker histones Bednar et al. [36
Reconstituted array Cryo-fixation No – Interdigitated solenoid model Robinson et al. [17
Reconstituted array Cryo-fixation No – Two-start zig-zag model with tetranucleosomal unit Song et al. [20
Electron spectroscopic image (ESI) with tomography Native chromatin (in cell) Ethanol dehydration, freeze-substitution Yes – Open and closed chromatin domains consisting of 10 nm fibres Fussner et al. [31
EM tomography Native chromatin (isolated) Freeze-substitution Yes Osmium ammine-B stain Variable two-start helical ribbon model Horowitz et al. [37
Native chromatin (in cell) Ethanol dehydration Yes Osmium staining of electron-dense precipitate produced by photoconverted DNA-bound fluorescent dye Heterogeneous 5–24 nm chromatin fibre Ou et al. [22
Low-angle X-ray scattering (LAXS) Isolated nuclei and chromatin Solution No – Two-start crossed-linker model Williams et al. [14
Small-angle X-ray scattering (SAXS) Isolated nuclei and chromatin Solution No – Irregularly folded nucleosome fibres Nishino et al. [29
X-ray crystallography Reconstituted nucleosome Crystal No – Nucleosome structure with angstrom-level resolution Luger et al. [4
Reconstituted nucleosome array Crystal No – Tetranucleosome folding unit Schalch et al. [16
Super-resolution light microscopy Native chromatin (in cell) – Yes Fluorescent protein, fluorescent-tagged antibody Nucleosome clutches along chromatin fibre Ricci et al. [21
Sequencing Native chromatin (in cell) – Yes Ligation Genome-wide structure with tri- or tetranucleosome folding unit Hsieh et al. [8
Method Sample Preparation Chemical fixation Probing method Observed structure Reference 
EM Native chromatin (isolated) Air-drying, freeze-etching Yes, no Negative stain, shadowing One-start solenoid model Finch and Klug [5
Native chromatin (isolated) Air-drying Yes Negative stain Two-start helical ribbon model Worcel et al. [12
Native chromatin (isolated) Air-drying, freeze-drying Yes, no Negative stain, shadowing, non-stained Two-start helical ribbon model Woodcock et al. [13
Native chromatin (isolated) Freeze-drying Yes Negative stain, shadowing, non-stained Two-start crossed-linker model Williams et al. [14
Reconstituted nucleosome array Air-drying Yes Negative stain Two-start model Dorigo et al. [15
Reconstituted nucleosome array Air-drying Yes Negative stain Interdigitated one-start solenoid model Robinson et al. [17
EM and in silico Reconstituted nucleosome array Air-drying Yes Positive-staining, shadowing Heteromorphic chromatin fibres Grigoryev et al. [19
Cryo-EM Native chromatin (isolated) Cryo-fixation No – Stem conformation of linker DNA segments formed by linker histones Bednar et al. [36
Reconstituted array Cryo-fixation No – Interdigitated solenoid model Robinson et al. [17
Reconstituted array Cryo-fixation No – Two-start zig-zag model with tetranucleosomal unit Song et al. [20
Electron spectroscopic image (ESI) with tomography Native chromatin (in cell) Ethanol dehydration, freeze-substitution Yes – Open and closed chromatin domains consisting of 10 nm fibres Fussner et al. [31
EM tomography Native chromatin (isolated) Freeze-substitution Yes Osmium ammine-B stain Variable two-start helical ribbon model Horowitz et al. [37
Native chromatin (in cell) Ethanol dehydration Yes Osmium staining of electron-dense precipitate produced by photoconverted DNA-bound fluorescent dye Heterogeneous 5–24 nm chromatin fibre Ou et al. [22
Low-angle X-ray scattering (LAXS) Isolated nuclei and chromatin Solution No – Two-start crossed-linker model Williams et al. [14
Small-angle X-ray scattering (SAXS) Isolated nuclei and chromatin Solution No – Irregularly folded nucleosome fibres Nishino et al. [29
X-ray crystallography Reconstituted nucleosome Crystal No – Nucleosome structure with angstrom-level resolution Luger et al. [4
Reconstituted nucleosome array Crystal No – Tetranucleosome folding unit Schalch et al. [16
Super-resolution light microscopy Native chromatin (in cell) – Yes Fluorescent protein, fluorescent-tagged antibody Nucleosome clutches along chromatin fibre Ricci et al. [21
Sequencing Native chromatin (in cell) – Yes Ligation Genome-wide structure with tri- or tetranucleosome folding unit Hsieh et al. [8

Nucleosome folding determinants revealed by conventional structural analyses

In addition to the static nucleosome folding structure, its changeability with cellular environments has often been investigated, especially its dependence on ion concentrations. Earlier EM studies found that salt concentration is a key parameter for the formation of chromatin fibres [5,13,38,39]. In these studies, by using extracted chromosomes, monovalent (Na+ and K+) and divalent (Mg2+ and Ca2+) cations were identified as important factors for controlling overall chromosome folding. This is consistent with the fact that chromosome condensation occurs during mitosis when Mg2+ and Ca2+ are enriched in chromosomes [40,41]. Sedimentation velocity analyses of reconstituted nucleosome arrays showed cation-dependent compaction with monovalent and divalent salt, and polyamine [19,42,43]. Positively charged ions neutralize the repulsion between the negatively charged DNA, thereby enhancing in vitro chromatin compaction. Other studies revealed transitions in nucleosome folding status depending on magnesium concentration [17,42,4446]. Whereas lower magnesium concentrations (∼0.5 mM) caused moderate nucleosome folding, intermediate concentrations (0.5–2 mM) induced fully compacted folding into a 30-nm fibre. Higher concentrations (3–5 mM) enhanced inter-fibre interactions, facilitating oligomerization towards higher-order structures. These magnesium concentrations are close to physiological values measured in cells (0.5–1.2 mM free and 10–30 mM bound to protein) [47].

Another important factor influencing nucleosome folding is binding of linker histone (H1 or H5), which binds to the nucleosome at the DNA entry and exit points. In earlier EM studies, linker histones stabilized wrapping of DNA around the nucleosome [48] as well as a higher-order filament-like chromatin fibre [5,25]. Further EM-based structural analyses showed that linker histones induce the formation of a stem-like organization of the linker DNA as it enters and exits the nucleosome particle [36], and also that linker histones direct the compaction of chromatin fibres [49]. EM and biochemical studies proposed two models for binding of linker histones, either symmetrical or asymmetrical binding with respect to the dyad axis of the nucleosome. In the symmetrical model, the globular domain of the linker histone is located on the dyad axis and binds both linker DNAs [48,50], whereas in the asymmetric model, it is displaced from the dyad to contact a single linker [51,52]. Despite recent structural analyses, these different binding models are still controversial [20,5355]; however, an existence of two different binding modes was suggested to contribute to varying degrees of chromatin compaction [53].

Another essential factor affecting nucleosome folding states are the chemical modifications in histones at their N- and C-terminal tail regions, which occur post-translationally on each of the core histones, H2A, H2B, H3, and H4, in forms such as methylation and acetylation. Because these modifications are associated with cellular functions like transcriptional regulation, structural analyses in the presence of different modifications provide insights into coupling between chromatin structures and functions. The effects of these epigenetic modifications on nucleosome folding have often been investigated by EM imaging and sedimentation analysis using reconstituted nucleosome arrays containing histones with chemically modified tails or substituted amino acids. For instance, structural analyses revealed that acetylation of lysine 16 on histone H4 (H4K16ac) inhibits formation of compacted nucleosome fibres in reconstituted arrays [5658]. Other studies revealed that acetylation in H3 (H3K4ac, H3K9ac, H3K14ac, H3K18ac, H3K23ac, H3K27ac, and H3K56ac) also abrogates inter- and intra-nucleosome array interaction [59,60]. Furthermore, X-ray crystallographic analyses have indicated that interactions between negatively charged regions in H2A and H2B, called the acidic patch [4,61], and positively charged tail regions in H4 are a significant determinant of fibre compaction [57,58,62]. Supporting this, it has been shown that replacement of H2A with its variants H2A.Z [63] or H2A.Bbd [64], which enhance or reduce the negative charge of H2A, causes more or less compacted fibre folding, respectively.

Sequencing-based nucleosome folding structural analysis methods

Although the methods discussed above can provide spatial information of the components of the target molecules, they have not yet been applied to analyze nucleosome folding at specific endogenous loci, which would require identification of the DNA sequence. A classical assay applying DNA sequencing to structural analysis is DNase I footprinting to locate the positions of DNA-bound factors. Recently, the advent of next-generation sequencing has allowed expansion of these techniques to a genome-wide level, revealing tracts of structured chromatin formed by chromatin-associated proteins. One such assay for locus-specific analyses is the DNA adenine methyltransferase identification (DamID) assay, in which genomic regions spatially proximate to a protein of interest are analyzed by sequencing methylated genomes from cells with Dam genetically fused to the protein of interest [65,66]. Studies using this approach revealed higher-order structures such as lamina-associating domains (LADs) [67,68].

While the above techniques identify chromatin structuring along the genome co-ordinate, in order to analyze the three-dimensional structure of chromosomes, information about the spatial proximity between pairs of genomic loci is required. Dekker et al. [6] developed an assay, called chromosome conformation capture (3C), that is capable of obtaining such information. In 3C, physical proximity between a pair of genomic loci is inferred from ligation frequencies of free DNA ends in chemically fixed and fragmented chromatin. Initially, this assay was restricted to interrogating interactions between one pair of loci at a time; however, next-generation sequencing also allowed parallelization of 3C, giving rise to Hi-C technologies [7], which have since been widely applied to characterize chromatin architecture across the genome [69]. Studies using Hi-C revealed characteristic features in higher-order chromosome folding, such as a broad segregation of the genome into compartments A and B [7,70], and topologically associated domains (TADs) [71,72]. The separation distance between loci tends to be inversely related to their ligation frequency, which allows three-dimensional (3D) chromosome structures to be obtained via computational methods that derive optimal spatial positions for loci that satisfy their interaction frequencies (e.g. Figure 1, fractal globule [7]) [73,74].

Recently, the resolution of Hi-C has been improved to the nucleosome level [8,9] from the original ∼1 Mbp resolution [7]. In Hi-C methods, the resolution depends on the genomic cutting frequency and this dramatic increase in resolution was achieved by fragmentation of chromatin to the nucleosome level using micrococcal nuclease (MNase), which only digests DNA that is not tightly bound to histones. This method, called Micro-C, revealed nucleosome folding structures across the yeast chromosome [8] (Figure 2). The results suggested an existence of tri- or tetranucleosome zig-zag folding motifs within the fibre, although periodic folding repeats were rarely observed across the genome. The results also suggested the presence of self-associating domains within gene loci called ‘gene crumples’. Meanwhile, another recent method termed RICC-seq employs ionizing γ-radiation to cleave genomic DNA that are spatially proximal followed by sequencing of the products, which provides locus-specific information of the cleaved fragments [75]. Subsequent computational analysis suggested zig-zag and solenoid structures in heterochromatin and open chromatin regions, respectively.

Nucleosome-level chromatin structure analysis with epigenetic annotations using sequencing-based methods.

Figure 2.
Nucleosome-level chromatin structure analysis with epigenetic annotations using sequencing-based methods.

3D nucleosome folding structure (top) constructed by computational modelling using nucleosome interaction data (second top) obtained from Hi-C methods. (The case for yeast chromatin [8] is shown.) ChIP-seq analyses provide genome-wide epigenetic profiles, including localization of chromatin-binding proteins, distribution of histone modification and nucleosome occupancies. The modelled 3D nucleosome folding structure reflects an integrative epigenetic feature that can predict locus status (left).

Figure 2.
Nucleosome-level chromatin structure analysis with epigenetic annotations using sequencing-based methods.

3D nucleosome folding structure (top) constructed by computational modelling using nucleosome interaction data (second top) obtained from Hi-C methods. (The case for yeast chromatin [8] is shown.) ChIP-seq analyses provide genome-wide epigenetic profiles, including localization of chromatin-binding proteins, distribution of histone modification and nucleosome occupancies. The modelled 3D nucleosome folding structure reflects an integrative epigenetic feature that can predict locus status (left).

Integrating sequencing-based methods to predict locus-specific nucleosome folding structures

Compared with the conventional structural analysis methods, sequencing-based methods lack the resolution for atom-level descriptions of nucleosomes; however, they have the distinct advantage of providing nucleosome folding structures in a genome-wide fashion, thereby opening the door to comparisons between nucleosome folding structures and the distinct molecular environments at individual gene loci. This ability will promote arguments on the nature of basic nucleosome folding structures at endogenous loci, for example, by testing whether and how frequently nucleosomes are folded in a zig-zag/solenoid or condensed/relaxed manner. Furthermore, considering that a variety of molecular factors bind to every genomic locus distinctively, this locus-specific structural information will allow discrimination between different molecular mechanisms of nucleosome folding as well as elucidation of their physiological meaning. In particular, it will be crucial to know how nucleosome folding structures are correlated with attributes of genomic loci such as transcriptional status, promoter/gene body, heterochromatin/euchromatin, histone modifications, associating linker histones/transcription factors, and the DNA sequence itself.

Since current genome-wide analysis methods allow identification of molecular factors bound to each genomic locus at nucleosome resolution, it is straightforward to correlate sequencing-derived nucleosome folding structures with various molecular factors (Figure 2). Chromatin immunoprecipitation sequencing (ChIP-seq) is a well-established method to analyze such factors, including histone modifications and occupancies of chromatin-binding proteins across the genome. Similar to Hi-C, nucleosome resolution in ChIP-seq can be realized by fragmenting the genome to nucleosome or sub-nucleosome levels with enzymatic digestion. For example, one method, called X-ChIP-seq, used MNase [76], while another method, ChIP-exo, employed λ-exonuclease, which digests DNA that is unbound to any proteins, to determine binding regions of target proteins at single-base resolution [77].

Such finer levels of genome-wide ChIP-seq data add an additional layer to the nucleosome-resolved structural data, allowing analyses of correlated changes in structure and epigenetic modifications under different conditions, such as switching a gene on or off. Indeed, although at a higher length scale, this type of analysis revealed many relationships between chromatin structures and binding factors [7,70,78]. For example, it has been shown that binding locations of the CCCTC-binding factor (CTCF), which works as a structural insulator of the chromosome, coincide with TAD boundaries [70,79,80], whereas the protein YY1 mediates long-range chromatin loops between enhancers and promoters [81]. For another example, it was found that LADs are correlated with transcriptionally repressive histone marks including H3K9me2/3 and H3K27me3 [82,83]. It is becoming clear that higher-order transitions in chromatin architecture are associated with multiple cellular phenotypes, for example, switching of loci between A and B compartments during cellular reprogramming [84]. An outstanding question however, is to what degree structural transitions at the much shorter length scales of nucleosome folding in the chromatin fibre can also influence gene expression and phenotype.

Future perspectives: chromatin folding at the sub-nucleosomal level

Recent studies demonstrated the need for structural analyses of native chromatin at a finer resolution than the nucleosome level, or in other words at the ‘nucleosome orientation’ level, where nucleosome orientation can be defined by the nucleosomal dyad axis, which divides the nucleosome-bound DNA in half (Figure 3A). Firstly, several studies suggest ‘nucleosome asymmetry’, which results from differently modified histone tails in the two copies of core histones or replacement of one of the core histones with a variant [85,86]. Secondly, it has been shown that, at the promoter region, the nucleosome dyad has asymmetric DNA–histone interactions due to activity of the RSC chromatin remodelling complex [87]. Thirdly, several studies found ‘non-canonical nucleosomes’ deviating from the usual histone octamer structure [88]. Although most nucleosomes consist of histone octamers with left-handed DNA wrapping (canonical nucleosomes), non-canonical nucleosomes are composed from different copies of histones or histone variants with a different manner of DNA wrapping. Examples of non-canonical nucleosomes include: reversome with right-handed DNA wrapping [89], tetrasome with two copies of CENP-A and H4 but lacking a H2A/H2B dimer [90], and hemisome with right-handed DNA wrapping and only one copy of each histone [91].

Nucleosome orientation as a metric of sub-nucleosomal chromatin structure.

Figure 3.
Nucleosome orientation as a metric of sub-nucleosomal chromatin structure.

(A) A model of the chromatin fibre where individual nucleosomes assume distinctive orientations with respect to one another based on the direction of their dyad axis. (B) Factors that may influence nucleosome orientation. (i) ATP-dependent chromatin remodelling complexes (blue) may change the orientation of stretches of nucleosomes. (ii) Swapping of canonical with non-canonical nucleosomes may result in characteristic changes to nucleosome orientation. (iii) Asymmetric histone modifications could influence nucleosome orientation to regulate the juxtaposition of binding sites for bivalent chromatin-binding proteins (purple). (iv) Unstable or dynamic nucleosomes may assume defined orientations upon binding of factors such as linker histones (light green), facilitating downstream regulatory steps.

Figure 3.
Nucleosome orientation as a metric of sub-nucleosomal chromatin structure.

(A) A model of the chromatin fibre where individual nucleosomes assume distinctive orientations with respect to one another based on the direction of their dyad axis. (B) Factors that may influence nucleosome orientation. (i) ATP-dependent chromatin remodelling complexes (blue) may change the orientation of stretches of nucleosomes. (ii) Swapping of canonical with non-canonical nucleosomes may result in characteristic changes to nucleosome orientation. (iii) Asymmetric histone modifications could influence nucleosome orientation to regulate the juxtaposition of binding sites for bivalent chromatin-binding proteins (purple). (iv) Unstable or dynamic nucleosomes may assume defined orientations upon binding of factors such as linker histones (light green), facilitating downstream regulatory steps.

Further improvement of the resolution of sequencing-based structural analyses down to the nucleosome orientation level will allow discovery of molecular mechanisms underlying nucleosome folding that regulate genomic processes. Increased resolution could be achieved by developing Hi-C protocols capable of finer-grained analysis of the ligation frequency of the DNA wrapped around nucleosomes. We expect that nucleosome orientation information will unveil new correlations between molecular factors and genome functions. Previous studies suggest that this individual nucleosome orientation could be changed by several factors (Figure 3B) [87]. For example, nucleosome orientation may be modulated by: the activity of chromatin remodelling complexes, incorporation of non-canonical nucleosomes, asymmetric nucleosome modifications, or binding of proteins (Figure 3B, i–iv). Such orientation change may serve as a regulatory input or output controlling a genomic function, such as transcription. Thus, we expect that revealing chromatin structure at such ‘sub-nucleosome-level’ resolution will unveil further mechanistic details of molecular processes on the genome.

Conclusions

Over the past four decades, various structural analysis techniques have contributed to a growing understanding of genome structure spanning a hierarchy of length scales from individual nucleosomes to whole chromosomes. As opposed to other methods, DNA sequencing-based methods can uniquely provide locus-dependent structures. For instance, these methods can be used to screen for characteristic genomic loci of interest, following which studies based on imaging and biochemical analyses can visualize and confirm nucleosome folding structure of targeted genomic loci with a finer resolution and in living cells. Such combined studies will elucidate typical structural changes caused by binding of chromatin-associated molecules for different processes, such as gene activation. Furthermore, sequencing-based chromatin structure studies may facilitate future clinical developments. To date, increasing evidence suggests long-range chromatin misfolding in diseases [92]. For example, it was found that transactivation of an oncogene is promoted by disruption of normal chromatin structure, resulting in increased cell proliferation [93]. Correction of disease-associated alterations to local chromatin structure will provide a promising avenue for future therapeutic interventions.

Abbreviations

     
  • 3C

    chromosome conformation capture

  •  
  • ChIP-seq

    chromatin immunoprecipitation sequencing

  •  
  • EM

    electron microscopy

  •  
  • LAD

    lamina-associating domains

  •  
  • MNase

    micrococcal nuclease

  •  
  • TAD

    topologically associated domain

Author Contribution

All the authors wrote and edited the manuscript.

Funding

This work was supported by PRESTO, Japan Science and Technology Agency [JPMJPR15F7], by grants-in-aid for Young Scientists (A) [24687022], Challenging Exploratory Research [26650055] and Scientific Research on Innovative Areas [23115005], Japan Society for the Promotion of Science, and by grants from the Takeda Science Foundation and the Mochida Memorial Foundation for Medical and Pharmaceutical Research. D.G.P. acknowledges additional support from a RIKEN Foreign Postdoctoral Researcher fellowship.

Competing Interests

The Authors declare that there are no competing interests associated with the manuscript.

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