This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

Physical Nature of Chromatin in the Nucleus
Kazuhiro Maeshima,1,2 Shiori Iida,1,2 and Sachiko Tamura1
1

Genome Dynamics Laboratory, National Institute of Genetics, Mishima, Shizuoka 411-8540, Japan2

Department of Genetics, School of Life Science, Sokendai (Graduate University for Advanced Studies),
Mishima, Shizuoka 411-8540, Japan

Correspondence: kmaeshim@nig.ac.jp

Genomic information is encoded on long strands of DNA, which are folded into chromatin
and stored in a tiny nucleus. Nuclear chromatin is a negatively charged polymer composed ofDNA, histones, and various nonhistone proteins. Because of its highly charged nature, chromatin structure varies greatly depending on the surrounding environment (e.g., cations,molecular crowding, etc.). New technologies to capture chromatin in living cells havebeen developed over the past 10 years. Our view on chromatin organization has drasticallyshifted from a regular and static one to a more variable and dynamic one. Chromatin forms
numerous compact dynamic domains that act as functional units of the genome in higher
eukaryotic cells and locally appear liquid-like. By changing DNA accessibility, these
domains can govern various functions. Based on new evidences from versatile genomics
and advanced imaging studies, we discuss the physical nature of chromatin in the crowded
nuclear environment and how it is regulated.

DNA, GENOME, AND INFORMATIONSTORAGE IN THE NUCLEUS

H

ow much information can be stored in the
nucleus of a cell? Biological (or genetic)
information is inscribed in deoxyribonucleic
acid. DNA is a negatively charged double-helix
polymer, composed of four bases (guanine, G;
adenine, A; thymine, T; cytosine, C), which arelinked by a negatively charged phosphate backbone (Fig. 1A). A single human cell has ∼2 m
of DNA (0.33 nm × 3 × 109 bp × 2 sets of chromosomes), which is ∼2 × 105-fold longer than
the typical diameter of a nucleus (∼10 µm) (Fig.
1B). Given that the human genome has 3 × 109
bp composed of four bases, there are 4(3×10^9)

combinations of information in it, which isequivalent to 2(6×10^9), or 750 M (0.75 G) bytes.
This roughly corresponds to only a single compact disc (CD) (∼700 M bytes) and is muchsmaller than our current smartphones’ memorystorage.
However, the memory density of the humangenome in the nucleus can be calculated as∼7.5 × 1014 bytes/mm3, considering that the nuclear volume is roughly 1000 µm3. This memory
density is several orders of magnitude higher
than a CD (5.2 × 104 bytes/mm3), blu-ray disc(QL) (9.4 × 106 bytes/mm3), or even a ﬂash
memory (16 Gb, 6.3 × 108 bytes/mm3) (Church
et al. 2012). Including possible chemical modiﬁcations on DNA and histones, such as methyl-

Copyright © 2022 Cold Spring Harbor Laboratory Press; all rights reserved
Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675

169© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.

A

5c
O

P
O

O

– –

Thymine(T)

Adenine
(A) N

O–

N

–

O

O

O

O

O–

O

H

N HN
N

N

O

O
O

H2N N

H

O

O

H2N

N

P

P
H

O

–O

NH N
N

O

O
O
O

P

H
N

NH
O

N

NH2

Guanine
(G)

P

H O

N

N

H

O

H2N

O

O

O
–

–

O

O

N

N

O

–

–

P
N

O

O–
O

O
NH2

H

P

O
–

–

N
O

N
O
O

OH

HN

N
O

3c

O

NH2

O–
O

–

P

O–
O

N
O

O

Cytosine
(C)

O

–

O–

3c

5c

DNA (deoxyribonucleic acid)

B

–

DNA

–

–

–

–
2 nm

–

–

–

–

–

Histone+Nucleosome(10-nm fiber)

–

–
–

–

–
–

–
–

11 nm

Nucleus~10 PmFigure 1. DNA, histones, and the nucleosome. (A) Schematic of the structure of deoxyribonucleic acid (DNA).
DNA is a long double-helix polymer made from repeating units called nucleotides, that consists of four bases:
adenine (A), cytosine (C), guanine (G), and thymine (T), which are attached to the sugar/phosphate. Adenine
pairs with thymine and guanine pairs with cytosine by hydrogen bonds. The two strands of DNA run in opposite
directions to each other in an antiparallel manner (shown by arrows). Phosphate backbones are negatively
charged (marked with “-”). (Illustration and legend from Nozaki et al. 2018; reproduced with modiﬁcations,with permission, from the authors.) (B) Negatively charged DNA (top row) is wrapped around basic (or positive)
core histone octamers (yellow on second row) to form 10-nm ﬁber, and it is further organized in a cell nucleus.
Note that the ﬁber is still negatively charged.
170

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

Physical Nature of Chromatin

ation and acetylation, this calculation increased
dramatically to encompass an enormous memory size and density. Reading of information
requires energy and if one assumes that a 1 Gb
ﬂash memory needs roughly ∼1 J (1 W per
1 sec) of electric power and a similar level ofenergy was required in each cell of our body(∼40 × 1012 cells; Bianconi et al. 2013) to readits genetic information, the total energy required
would be ∼4 × 1013 J, which is far greater than
the total electric energy produced by all the nuclear power plants on earth (one plant can produce ∼1 × 109 J). In other words, the nuclear
genome works with amazing energy efﬁciency.
It is thus appropriate to call the nucleus atruly “amazing memory device” in terms of bothinformation storage and readout. Although our
above example used the human cell nucleus, thisremarkable property is shared widely across all
eukaryotic nuclei. To function correctly, genome DNA needs to be carefully and properlyorganized in the nucleus, where differences in
DNA folding can bring about dramatic changes
in information readout. In this article, we review
and discuss how genomic DNA is organized andbehaves in the nucleus.
THE NUCLEOSOME

The electric charge of its components is an essential part of understanding the organization of the
genomic information in the nucleus. Negatively
charged DNA is wrapped around a core histone
octamer (Fig. 1B), which consists of four positively charged histone proteins (H2A, H2B, H3, andH4), and forms a nucleosome (Figs. 1B and 2;
Kornberg 1974; Olins and Olins 1974; Woodcock
et al. 1976). The nucleosome structure has been
resolved to 1.9 Å (Fig. 2B; Davey et al. 2002), in
which 147 base pairs of DNA are wrapped in 1.7
left-handed superhelical turns around the histone octamer to generate a 225 kDa nucleosome
core structure. The core histone octamer contains
two copies of each of the four core histones,
which are highly conserved in both length andamino acid sequences among eukaryotic species(Fig. 2A; Luger et al. 1997). H2A/H2B andH3/H4 form stable dimers at physiological conditions. Two H3/H4 dimers also form a stable

tetramer under such conditions. Each core histone contains an ∼60 amino acid residue histonefold domain, which accounts for ∼70%–75% of
the mass of each protein (Fig. 2A) and provides ahighly structured and extensive dimerization interface between the H3/H4 and H2A/H2B pairs(Fig. 2B). Each histone also has an amino-terminal tail domain, which is an intrinsically disordered region (IDR) and extends to the exterior of
the nucleosome, while H2A contains an additional carboxy-terminal tail domain. Thus, a single nucleosome has 10 tails (Fig. 2C; Luger et al.
1997; Davey et al. 2002), which are highly positively charged due to a preponderance of lysines
and arginines. For example, the ∼35-residue H3tail and the ∼20-residue H4 tail contain 13 and 9
positively charged residues, respectively. Becauseof this positive charge abundance, the long, disordered tail domains contribute to the thermal
stability of the nucleosome core (Ausio et al.
1989), and play important roles in the interactions between nucleosomes to contribute higher-order organization of chromatin. Chromatin
is composed of nucleosomes and associated nonhistone proteins (Figs. 1B and 3; discussed later;
Olins and Olins 2003).
In most eukaryotic species, an additionalfamily of histone proteins bind to the nucleosome. Linker histones (also referred to as H1s)
are highly basic proteins but share no structural
homology with the core histones (Fig. 2D; Woodcock et al. 2006; Bednar et al. 2017). Interestingly,
two IDRs in the carboxy- and amino-terminal
regions of H1s connected by an ∼80-residue“globular” domain are important for chromatincondensation (Turner et al. 2018). Whereas it isassumed that H1s bind to the exterior of nucleosomes and stabilize the wrapping of DNA within
nucleosomes (Fig. 2D), H1s seem to be highly
mobile in the cell (Misteli et al. 2000).
Each nucleosome particle is connected bylinker DNA segments (∼20–80 bp; ∼6.6–27 nm)
that generate repetitive motifs of ∼200 bp and aredescribed as “beads on a string” or as a “10-nm
ﬁber” (Figs. 1B and 3A; Olins and Olins 2003).
This 10-nm ﬁber produces electrostatic repulsion
between adjacent regions because only about halfof the DNA negative charges derived from thephosphate backbone are neutralized by the basic

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

171

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.

A

NH2 17
12

89 92

26

COOH
118

H2A

98

37
26

H2B
67

131

35

H3

31

93

20

B

H4

C

H2A

*

*

H2B

H4

*

*

*

*

*
*

*

H2B

*

*
*

* * *

H3 *

*

*

*

H4

*

* H3

*

*
*

*

H2AC

*
*

*

*

D

H1
carboxy-terminal
domainH1globulardomainFigure 2. Structures of core histones and nucleosome. (A) Schematic of the core histone structures; histone-folddomains are enclosed by gray boxes, tail domains are denoted by dashed lines, and α-helices are shown bycolumns. The positions of residues ﬂanking the histone fold in each protein are shown. The approximate residue
at the end of the tail domain nearest the histone fold is also shown. (B) The nucleosome structure at 1.9 Å
resolution (Davey et al. 2002). Note that histone colors correspond to A. Amino acid residues of histone tails(intrinsically disordered regions [IDRs]) are shown as ball models. Basic residues (lysines and arginines) in the
tail domains are highlighted with a darker color. (C) Amino-terminal tail domains in B are extended away fromthe nucleosome core. H2A also has a carboxy-terminal tail. Basic residues (lysines and arginines) in the taildomains are colored in orange. An asterisk indicates sites of lysine acetylation. (Illustrations are based on data in
Wolffe and Hayes 1999 and Pepenella et al. 2014a.) (D) Structural model of a nucleosome with a linker histone
H1. (The model in D was created from PDB data in Bednar et al. 2017.)
172

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

Physical Nature of ChromatinA
Mg2+ / cations
In vitro

110-nm fiber (no salt)

2330-nm fiber (low salt)

Condensate/liquid droplet?

B
CationMolecular crowding

C

Chromosome
territory

Compartment A

Compartment B

10-nm fiber

DomainCohesin~200 nm

Nucleosome

Figure 3. Chromatin structure. (A) (Left) 10-nm ﬁbers form in no cation conditions (no salt). (Center) Threetypes of 30-nm ﬁbers form with a low concentration of cations/Mg2+: (1) a solenoid (one-start) model, (2) a twostart zigzag, and (3) a zigzag tetranucleosomal model (Song et al. 2014). (Right) Large chromatin condensateswith interdigitated 10-nm ﬁbers that form in more physiological cations or higher cations. Note, this condensate
lacks the 30-nm structures (Maeshima et al. 2016b). Recently, under a certain speciﬁc condition, the condensates
were shown to be liquid droplets formed by liquid–liquid phase separation (Gibson et al. 2019). (Panels from
Maeshima et al. 2020; reproduced with modiﬁcations, with permission, from the authors.) (B) Polymer melt
model. Under low cation and/or molecular crowding conditions, 10-nm ﬁbers could form 30-nm chromatinﬁbers via intraﬁber nucleosome associations. An increase in cation concentration and/or molecular crowding
results in interﬁber nucleosomal contacts that interfere with intraﬁber nucleosomal associations, leading to amelted polymer-like or “sea of nucleosomes” state (Dubochet et al. 1986; Eltsov et al. 2008; Maeshima et al. 2010).
In this state, tetranucleosome zigzag motifs are occasionally seen (highlighted in red). Note that we show a highly
simpliﬁed two-dimensional nucleosome model in these illustrations. (C) A simpliﬁed scheme for hierarchicalchromatin organization in the nucleus. The 10-nm ﬁber is compacted into chromatin domains (e.g., topologicallyassociating domain [TAD]/contact domain/loop domain) (Dixon et al. 2012; Nora et al. 2012; Sexton et al. 2012;
Dekker and Heard 2015), which interact over long distances to form chromatin compartments (LiebermanAiden et al. 2009). Compartments generally represent a transcriptionally active or open chromatin state (compartment A) and an inactive or closed chromatin state (compartment B). A single interphase chromosome isoccupied in a chromosome territory (highlighted as different colors) (Cremer and Cremer 2001).

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

173

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.

core histones (Fig. 3A; Maeshima et al. 2014). For
further folding of the 10-nm ﬁber, the remaining
negative charges have to be screened by otherfactors, such as linker histones, nonhistone proteins, cations, and other positively charged molecules. Chromatin, which consists of 10-nm ﬁbers and various nonhistone proteins, is overall
still a negatively charged polymer because thenegative charge of DNA is not completely neutralized by these positively charged molecules.
Therefore, chromatin can dynamically change
its local structure depending on the electrostaticstate of its environment (Fig. 3A; Cole 1967;
Earnshaw and Laemmli 1983; Hansen 2002;
Hudson et al. 2003; Takata et al. 2013). Suchchanges in local chromatin structure are criticalfor gene expression because they directly govern
access to the DNA and therefore impact how the
genome is scanned and read in the nucleus.
STRUCTURAL VARIATIONS OF CHROMATIN
IN VITRO: 10-NM FIBERS, 30-NM FIBERS,
AND CHROMATIN CONDENSATES

A crucial factor that inﬂuences local chromatin
structure is the ionic condition of the surrounding environment. For instance, in the presenceof linker histones and a low concentration of
cations (e.g., <∼1 mM Mg2+ or <∼50 mMNa+), the puriﬁed 10-nm ﬁber (Fig. 3A, left) isfolded into a zigzag or solenoidal ﬁber structurewith a diameter of 30 nm, which is called the 30nm chromatin ﬁber (30-nm ﬁber) (Fig. 3A, center; Finch and Klug 1976; Woodcock et al. 1984;
Schalch et al. 2005; Robinson et al. 2006; Songet al. 2014). Such low ionic conditions partially
screen the ﬁber’s electrostatic charge, leading tolocal contacts between nucleosomes in the ﬁber
and the formation of the folded 30-nm structure(Fig. 3A, center). Note, the concentration of cations required for this folding depends on the sizeand concentration of the ﬁber.
More cations are present at physiologicalconcentrations that result in further charge
screening. At these levels, electrostatic repulsion
between neighboring nucleosomes almost disappears and large chromatin condensates form
(Hansen 2002; Maeshima et al. 2016b; Gibson
et al. 2019; Strickfaden et al. 2020). Such conden174

sates are organized like a melted polymer or “seaof nucleosomes” and do not contain the 30-nm
structure (Fig. 3A,B, right; McDowall et al. 1986;
Eltsov et al. 2008; Maeshima et al. 2016a). Nucleosomes are free to contact long distal nucleosome
partners, leading to interdigitation of their ﬁbers(Fig. 3A,B, right; Zheng et al. 2005; Kan et al.
2009; Maeshima et al. 2010).
This structural variation scheme of chromatin (Fig. 3A,B) was demonstrated in vitro (Maeshima et al. 2016b) using a well-deﬁned chromatin model, the 12-mer nucleosome array (Hansen
2002). Previous biochemical studies using thenucleosome arrays can explain why such large
condensates lack the 30-nm ﬁber (Dorigo et al.
2003; Kan et al. 2009; Sinha and Shogren-Knaak2010). The long tail domain of histone H4 (Fig.
2B,C) mediates the formation of both the 30-nm
ﬁber and large condensates. Consequently, organization into chromatin condensates can preventthe formation of 30-nm ﬁbers by sequestering theH4 tail. It is also important to note that this interdigitated chromatin folding might contributeto forming large chromatin structures such as
mitotic chromosomes (Hansen et al. 2018).
The physical state of chromatin condensatesis intriguing and a subject of ongoing research.
Gibson et al. showed that nucleosome arrays withcations developed into large chromatin condensates ∼10 µm in size. The resultant large condensates behaved like liquid droplets (Fig. 3A, right;
Gibson et al. 2019), in which the nucleosomes
self-organized through a process termed liquid–
liquid phase separation (LLPS) (Hyman et al.
2014; Mitrea and Kriwacki 2016; Rippe 2021).
The droplets rapidly fused to each other andthe ﬂuorescence recovery after photobleaching(FRAP) showed that ﬂuorescently labeled nucleosome arrays freely diffused in the droplets (Gibson et al. 2019). However, Strickfaden et al.
(2020) revealed that the condensates formed
from similar nucleosome arrays show a rather
solid-like property and that the liquid chromatincondensates, as shown by Gibson et al. (2019),
were only observed under a highly speciﬁc set of
conditions (Strickfaden et al. 2020). The condensates thus seem to have solid-like features at a
micron scale. Nonetheless, we infer that the condensates might have weak intrinsic droplet prop-

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

Physical Nature of Chromatinerties at a nanoscale (100–200 nm) range, giventhat each nucleosome possesses 10 IDRs (Fig. 2B,
C; Luger et al. 1997; Pepenella et al. 2014b). IDRs
often mediate multivalent contacts for droplet
formation (Lin et al. 2015). Emerging evidence
suggests that functional compartmental droplets
are formed by LLPS in the cell (Hyman et al.
2014; Mitrea and Kriwacki 2016). Further investigation will be required to examine the intrinsic
droplet property in the condensates.
CHROMATIN ORGANIZATION IN THE
NUCLEUSIn 1976, the 30-nm ﬁber was discovered in vitro
and has since been believed to be the basic chromatin structure in eukaryotic cells (Fig. 3A, center; Finch and Klug 1976; Woodcock et al. 1984;
Schalch et al. 2005; Robinson et al. 2006; Songet al. 2014). However, during the past 10 years,
extensive data from various techniques, including
cryo-electron microscopy (EM) (McDowall et al.
1986; Eltsov et al. 2008; Chen et al. 2016; Cai et al.
2018), X-ray scattering (Joti et al. 2012; Nishino
et al. 2012), electron spectroscopic imaging (ESI)(Fussner et al. 2012; Visvanathan et al. 2013), EM
tomography (Ou et al. 2017), stochastic opticalreconstruction microscopy (STORM) (Ricci et al.
2015), chromosome conformation capture (3C)/
Hi-C (Dekker 2008; Hsieh et al. 2015; Sanborn
et al. 2015; Ohno et al. 2019), and computational modeling (Collepardo-Guevara and Schlick
2014; Bajpai et al. 2017), demonstrated that chromatin consists of rather irregular and variablenucleosome arrangements like “sea of nucleosomes” (Fig. 3A,B, right). In addition to the cation issue discussed above, many factors in native
chromatin, including irregular nucleosome spacing, histone modiﬁcations and variants, andnonhistone protein binding, would preclude theformation of regular 30-nm ﬁbers, especially inactively transcribing cells. Thus, the 30-nm ﬁber
is unlikely to be the basic structure or unit ofchromatin in vivo. Nonetheless, the formationof tetranucleosome motifs is highly possible(Baldi et al. 2020; Krietenstein and Rando2020), as reported by new genomics studies thatfound occasional zigzag nucleosome conﬁgurations in budding yeast cells (Hsieh et al. 2015;

Ohno et al. 2019) and in heterochromatin regionsof human cells (Risca et al. 2017).
At a higher level of organization, a large-scalechromatin structure with a diameter ∼200 nm
has long been observed by light and electron microscopic imaging in more complex eukaryotes(Fig. 3C; Hu et al. 2009; Nozaki et al. 2017; Olinsand Olins 2018; Xu et al. 2018), while yeast cells
seem to have predominately open chromatinarrangements with clusters of only a few nucleosomes (Hsieh et al. 2015; Chen et al. 2016; Ohno
et al. 2019). Recently, a combination of 3D superresolution and scanning EM revealed “chromatin
domains,” which were observed as irregularly
sized clusters of nucleosomes (∼200 nm) (Hoffman et al. 2020; Miron et al. 2020). Interestingly,
the chromatin domains in euchromatic regions
maintained compact chromatin organizations
with active epigenetic marks, but their sizes
were smaller than those of heterochromatin,
which is consistent with the previous reports(Maeshima et al. 2015; Nozaki et al. 2017). ESI,
which maps phosphorus and nitrogen atoms
with contrast and resolution sufﬁcient to visualize
10-nm ﬁbers (Hendzel et al. 1999), also exhibited
such chromatin domain structures (Strickfaden
et al. 2020).
3C/Hi-C methods can be used to generate a
ﬁne contact probability map of genome DNA(Dekker and Heard 2015; Dekker 2021), andthis technology has revealed that the higher eukaryotic genome is partitioned into chromatin
domains (Fig. 3C): topologically associating domains (TADs) at the scale of several hundreds ofkilobytes (Dixon et al. 2012; Nora et al. 2012;
Sexton et al. 2012) and smaller contact/loop domains (mean size of ∼185 kb) (Rao et al. 2014).
These structures were conﬁrmed visually bysuperresolution imaging (Bintu et al. 2018).
The loop domains seemed to be held together
by cohesin (Figs. 3C and 5C,E), in cooperation
with CCCTC-binding factor (CTCF) (Zuin et al.
2014; Rao et al. 2017; Wutz et al. 2017; Bintu et al.
2018; for more details, see Dekker 2021). These
domains are often clustered as two distinct,
megabase scale compartments A and B (Lieberman-Aiden et al. 2009), which likely represent
transcriptionally active chromatin and inactive
chromatin, respectively (Fig. 3C). Collectively,

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

175

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.

chromatin organization in the nucleus appears tohave a hierarchy (Fig. 3C): from 10-nm ﬁbers todomains, domains to compartments A and B,
and from these compartments to chromosomes
to form the “chromosome territory” of the nucleus (Cremer and Cremer 2001).
The large-scale arrangements/domains described above seem to be fundamental chromatin features in the nucleus (Misteli 2020).
Notably, the domains have a compact organization (Fig. 3C), irrespective of euchromatin orheterochromatin (Nozaki et al. 2017; Mironet al. 2020; Strickfaden et al. 2020). To form
these compact domains, local nucleosome–nucleosome interactions seem to be critical (Nozaki et al. 2017; Strickfaden et al. 2020). Cations
like magnesium ion (Mg2+) can facilitate theseinteractions, as shown in the formation of cation(Mg2+)-dependent chromatin condensates invitro, which requires nucleosome–nucleosome
interactions by histone tails (Fig. 2C; Maeshima
et al. 2016b; Gibson et al. 2019; Sanulli et al.
2019; Strickfaden et al. 2020).
Furthermore, it is very likely that non-nucleosomal proteins and RNAs, as well as cations,contribute to the formation of compact chromatin domains through macromolecular crowding
effects (Figs. 3B and 5E; Asakura and Oosawa
1954; Marenduzzo et al. 2006; Hancock 2007).
Imai et al. (2017) found that the total density ofheterochromatin (208 mg/mL) was only 1.53fold higher than that of the euchromatic regions(136 mg/mL), while the DNA density of heterochromatin was 5.5–7.5-fold higher (Imai et al.
2017). This ﬁnding suggests that non-nucleosomal materials (proteins, RNAs, etc.), which are
dominant in both heterochromatin and euchromatin (∼120 mg/mL), play an important role incompact organization of the domains in the nuclear crowded environments.
The organization of chromatin may serve tomodulate the accessibility of larger protein complexes to target sites (Maeshima et al. 2015;
Miron et al. 2020). The compact domains might
hinder the access of large protein complexes suchas transcription factors and replication initiation
complexes to the inner core of chromatin domains (Maeshima et al. 2015; Miron et al.
2020). Decompaction of such domains with his176

tone modiﬁcations or the action of other proteins
can increase accessibility to the complexes to turnon gene transcription (Toth et al. 2004; Bian andBelmont 2012; Kieffer-Kwon et al. 2017; Nozaki
et al. 2017; Miron et al. 2020; Strickfaden
et al. 2020). Interestingly, it has been suggested
that various RNAs may be involved in the regulation of these compact chromatin domains (Nozawa and Gilbert 2019). Nozawa et al. (2017)
showed that scaffold attachment factor A (SAFA)/heterogeneous nuclear ribonucleoprotein U(hnRNP U) decompacts transcriptionally active
chromatin domains by forming a ﬁlamentshaped oligomer with nuclear RNA (Nozawa
et al. 2017). How RNAs act on the domain formation remains unclear and is an area to be investigated further.
Whereas the compact organization of the
genome appears to be primarily designed forstorage efﬁciency, it seems to have additional
functional roles to protect genomic information.
The compact organization can generate a springlike restoring force that resists nuclear deformation by mechanical stress and plays an importantrole in maintaining genomic integrity. Nuclei
with condensed chromatin possess signiﬁcantelastic rigidity, while those with decondensed
chromatin are considerably softer (Shimamoto
et al. 2017; Stephens et al. 2017; Maeshima
et al. 2018b; Agbleke et al. 2020; also see Miroshnikova and Wickström 2021). Interestingly, compact chromatin seems more resistant to radiationand chemical damage than the decondensed
form, probably because condensed chromatin
has lower reactive radical generation and is lessprone to chemical attack (Takata et al. 2013). The
maintenance of genomic DNA integrity by greater compaction may have contributed to the evolution of eukaryotic organisms.
LOCAL MOTION OF CHROMATIN IN THE
NUCLEUSThe view of chromatin organization discussed
so far suggests that chromatin is less physicallyconstrained and more dynamic than expected in
the regular static structure model with helicalcoiling (Maeshima et al. 2010). In addition, given that each nucleosome possesses 10 IDRs (Fig.

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

Physical Nature of Chromatin

2C; Luger et al. 1997; Pepenella et al. 2014b), it is
tempting to consider that chromatin locally behaves like a liquid (Fig. 3A,B, right).
Live-cell imaging studies using a LacO/LacIGFP system (Fig. 4A; Robinett et al. 1996) or related systems (Germier et al. 2017; Tasan et al.
2018; Eykelenboom et al. 2019) have long revealed
dynamic movements of chromatin in numerous
cells such as yeast, nematodes, ﬂies, and mammals(Marshall et al. 1997; Heun et al. 2001; Chubb
et al. 2002; Levi et al. 2005; Meister et al. 2010;
Hajjoul et al. 2013; Arai et al. 2017). This system
can track the motion of bacterial operator (LacO)
repeats (<256 repeats) inserted into the genome ofliving cells towhich ﬂuorescent LacI proteins bind(Fig. 4A; Robinett et al. 1996). Analyses of FRAP
have also demonstrated turnover rates of histones
or chromatin-associated proteins at bleachingsites in living cells and provided indirect information on chromatin motion (Misteli et al. 2000;
Kimura and Cook 2001; Meshorer et al. 2006).
Furthermore, CRISPR-based genome editing
technology has allowed a targeted region of interest in genome chromatin to be speciﬁcally labeled(e.g., Tasan et al. 2018) to visualize motion of that
region (Fig. 4B; Chen et al. 2013; Gu et al. 2018;
Guo et al. 2019; Ma et al. 2019).
Moreover, genome-wide local chromatin
motion has been investigated using single-nucleosome imaging and tracking in living cells
through ﬂuorescently tagged histone proteins(Fig. 4C; Hihara et al. 2012; Nozaki et al. 2017;
Nagashima et al. 2019; Lerner et al. 2020; GómezGarcía et al. 2021). This powerful imaging modality can trace the movement of nucleosomes
throughout the nucleus at the single-molecule
level with high resolution. Similar global chromatin movements have been described using ﬂuorescently labeled bulk histones, or DNA-bindingdye, to analyze their displacement correlation(Fig. 4D; Zidovska et al. 2013; Shaban et al.
2020). Development of imaging techniques thatrecord local chromatin motion at a genome-wide
scale enabled visualization of heterogeneity of local chromatin motion in a whole nucleus as achromatin heat map (Fig. 4E,F; Nozaki et al.
2017; Nagashima et al. 2019) or chromatin dynamics maps (Zidovska et al. 2013; Lerner et al.
2020; Shaban et al. 2020). These maps show over-all distributions of heterochromatin and euchromatin. As expected, the local chromatin motion
at the nuclear periphery (heterochromatin) is lessmobile (Fig. 4F). Consistently, statistical analysis
of genome-wide single-nucleosome tracking data
revealed heterogeneity of movements, whichroughly categorized chromatin into two types:
slow chromatin that moves under structurally
constrained environments and fast chromatinthat moves with fewer constraints (Fig. 4G; Ashwin et al. 2019, 2020). Lerner et al. (2020) suggested a correlation between chromatin mobility
and transcription factor binding. Furthermore,statistical analysis of single-nucleosome tracking
data has unveiled that the organized nucleosomes
within chromatin are locally mobile and behave
like a liquid at the scale of less than hundreds ofnanometers (Ashwin et al. 2019, 2020). In otherwords, when considered at the scale of the chromatin domain, chromatin is very dynamic andexhibits a liquid-like diffusion rather than a crystal-like structure, which has a long-range order. A
physical basis of the liquid-like behavior of chromatin is provided by polymorphism of the 10-nm
ﬁber with the long histone tails (Fig. 2C), which
can turn into various structures, including extended, folded, interdigitated, bent, and looped
structures (Collepardo-Guevara and Schlick2014).
FACTORS THAT CONSTRAIN CHROMATIN
MOTION IN THE NUCLEUS

Whereas local chromatin behaves like a liquid(Ashwin et al. 2019, 2020), each chromosome
occupied in the territory appears to be relatively
stable, without mixing of chromosomes (Fig. 3C;
Strickfaden et al. 2020). Certainly, the meansquare displacement (MSD) analysis of local nucleosome motion in living human cells revealed
that the plots almost reached a plateau value after
3 sec (Fig. 5A,B), which is proportional to thesquare of the radius of constraint (Rc; P = 6/5 ×Rc2) (Dion and Gasser 2013). In addition, theestimated radius of the nucleosome motion constraint in living human RPE-1 cells is 141 ±19.2 nm (mean ± SD), which roughly corresponds to the size of the chromatin domain mentioned above (i.e., local chromatin is constrained

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

177

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.

A

LacO/Lacl-GFP

B

CRISPR imaging

O
Lac

Chromatin

Lacl-GFP

dCas9-EGFP guideRNA

D

C

E

F

Nuclear interior

Surface2
13Chromatin dynamics
Small

Large
40

G

80 (nm/ 50 msec)

30
25

Slow

Fast

1

2
M/M*

P(M)

20
15
10
5
0

0

3

Figure 4. Visualization of local chromatin motion. Schematics for (A) LacO/LacI-GFP, (B) CRISPR-based
chromatin labeling, (C) single nucleosome imaging and tracking, and (D) bulk chromatin labeling for displacement correlation analyses. A small number of nucleosomes are labeled with photoactivatable GFP or otherﬂuorescent tags for single nucleosome imaging in C while much more are labeled in D. (E) Example of chromatin
dynamics used to generate a heat map. Movements are measured over 50 msec. Small movements are shown inblue and large movements are shown in red. Heat maps allow the visualization of the heterogeneity of local
chromatin motion in the nucleus. (F ) Diagrams of a chromatin heat map where the nuclear interior (left) andnuclear periphery (right) of a living HeLa cell are shown. The boxed regions 1–3 show nucleoplasm, nuclear
periphery, and around nucleolus periphery, respectively. Note that regions 2 and 3 are presumably heterochromatin regions shown in dark blue. The map of the nuclear periphery (surface) appears more bluish, showing lessmobile chromatin in the heterochromatin-rich regions. (G) Mean square displacement (MSD) distribution of
local chromatin motion. The MSD at the ﬁrst minimum between the peaks is denoted by M . P (M, 0.5 sec) from
10 cell samples were scaled by M and shown, indicating fast and slow peaks. For details, see Ashwin et al. (2019).
(Illustrations and legend from Maeshima et al. 2020; reproduced with modiﬁcations, with permission, from theauthors.)

178

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

A

B

Rc = 141 ± 19.2 nm
0.030
0.025

MSD (Pm2)

100 nm

0.020
0.015

Control

0.010

FA-fixed

0.005
0.000
0.0

C

0.5

1.0 1.5 2.0
Time (sec)

Chromatin motion

2.5

3.0

Chromatin motion

TSA
CohesinCohesinCohesinKD

Global holding

Chromatin motion

D
Active RNAPII

RNAPII-KD
Transcription
inhibition

NucleusRNAPII-Ser5P (glue)
Transcription factors

Hub

RNAPIIE

Hub
Cohesin
DomainTranscriptional
machinery
DomainFigure 5. Local chromatin motion and chromatin domains in the nucleus. (A) Three representative trajectories ofsingle nucleosomes (Nagashima et al. 2019). (B) Mean square displacement (MSD) plots (±SD among cells) of
single nucleosomes in living (black) and formaldehyde-ﬁxed (red) RPE-1 cells over time (0.05 to 3 sec). For eachsample, n = 20 cells. (Data from Nagashima et al. 2019.) (C) (Left) Decondensed chromatin in cells treated withhistone deacetylase (HDAC) inhibitor trichostatin A (TSA) shows increased chromatin movements resulting
from weakened nucleosome–nucleosome interactions and subsequently less local chromatin constraining. (Center) Usual state of chromatin with reduced movements. The chromatin domain is organized by local nucleosome–nucleosome interactions and global holding by cohesin. (Right) Loss of cohesin leads to less constraining
and a resultant increase in chromatin motion. (D) (Left and center), Cluster/condensate of active RNA polymerase II (RNAPII) and transcription factors (blue spheres) can work as a transient hub (cyan sphere) to weakly
connect multiple chromatin domains and to globally constrain chromatin motion. (Right) RNAPII inhibition, or
its rapid depletion, releases chromatin constraints and increases chromatin motion. (E) A current model ofchromatin domains. Chromatin is locally very dynamic, but the local chromatin motion is constrained by various
physical or geometrical factors to keep each chromosome in the chromosome territories. (Diagrams in panels C
and D from Babokhov et al. 2020; reproduced with modiﬁcations, with permission, from the authors.)Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

179

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.

to a range of the chromatin domain size and does
not appear to travel long distances) (Fig. 3C; Nagashima et al. 2019). Furthermore, the DNA replication foci observed via pulse ﬂuorescent labeling revealed constrained euchromatin andheterochromatin behavior in living mouse cells(Strickfaden et al. 2020). These properties are notconsistent with liquid behavior and suggest highly constrained motion. If chromatin behaves likea liquid locally but does not move over long distances, what constrains the chromatin in a nuclear aqueous environment? How are chromosome
territories maintained? How can chromatin motion be adjusted depending on the physiologicalstate(s) of the cell?
In contrast to ATP-dependent long-range
movements of chromatin (see reviews by Soutoglou and Misteli 2007; Seeber et al. 2018), local
chromatin motion is likely to be isotropic andprimarily driven by thermal ﬂuctuation (Fig.
5A; Marshall et al. 1997; Chubb et al. 2002; Leviet al. 2005; Nozaki et al. 2017; Ashwin et al. 2019).
Variability in this motion can be created by constraints from several physical or geometrical factors (Fig. 4G; Ashwin et al. 2019).
Nucleosome Interactions

Nucleosome–nucleosome interactions are one of
the factors to constrain chromatin motion. Chromatin becomes more mobile when treated withthe histone deacetylase (HDAC) inhibitor trichostatin A (TSA) (Fig. 5C, left). This drug increases histone H3 and H4 tail acetylation (Fig.
2C) and decondenses chromatin (Fig. 5C, left),presumably by decreasing the number of nucleosome–nucleosome interactions (Gorisch et al.
2005; Ricci et al. 2015; Nozaki et al. 2017; Strickfaden et al. 2020). Such artiﬁcial decondensationof chromatin (i.e., extended ﬁbers or ﬁbers with
nucleosome-free regions) reduces constraints
that restrict thermal-ﬂuctuating motion and results in elevated local chromatin movement (Fig.
5C, left; Amitai et al. 2017; Nozaki et al. 2017;
Ashwin et al. 2019).
Nucleoplasmic Milieu

Another factor is the quantity of free cationsin the nucleus. An increase in cations, such as180

Mg2+, can condense chromatin and lower its
local motion, since nucleosomes have residual
negative charges and tend to repulse one another as discussed above (Fig. 3A,B). Consistently,
intracellular ATP reduction, which releases
Mg2+ from ATP-Mg (Maeshima et al. 2018a),
hypercondenses chromatin and decreases its
motion (Visvanathan et al. 2013; Nozaki et al.
2017; Maeshima et al. 2018a). These ﬁndings
offer a regulatory mechanism for the higher-order chromatin organization by the intracellular
Mg2+-ATP balance. Since intracellular ATP levels vary depending on cell types (Pecqueur et al.
2013; Qian et al. 2014; Kieffer-Kwon et al. 2017),
they may regulate cellular functions such as
differentiation via changing higher-order chromatin organization. Moreover, hypertonic treatment (∼570 mOsm) had similar effects (Albiez
et al. 2006; Nozaki et al. 2017; Strickfaden et al.
2020). This may be due to an increase in intracellular cations and molecular crowding following hypertonic treatment (Albiez et al. 2006).
Chromatin Proteins

Various chromatin proteins constrain the local
chromatin motion driven by thermal ﬂuctuation, contributing to the retention of each chromosome in its “chromosome territory” (Fig. 3C;
Cremer and Cremer 2001) as well as genome
functions. The cohesin complex is an important
protein complex that can capture chromatin ﬁbers with its ring structure to form loops and forsister chromatid cohesion (Nasmyth and Haering 2005; Morales and Losada 2018; Nishiyama2019). Loss of cohesin leads to fewer chromatinconstraints and thereby a drastic increase in local chromatin motion (Fig. 5C, right; Dion et al.
2013; Nozaki et al. 2017; Ashwin et al. 2019; Itohet al. 2021).
Transcription

Interestingly, the RNA transcriptional machinery has a constraining role for local chromatin
motion in the nucleus (Fig. 5D,E; Nagashima
et al. 2019; Babokhov et al. 2020; Itoh et al.
2021). Knockdown of active RNA polymerase
II (RNAPII) or other transcription regulatory

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

Physical Nature of Chromatin

factors, as well as RNAPII inhibitor treatments,
releases RNAPII from chromatin and raises local chromatin movements (Nagashima et al.
2019). Consistent with this ﬁnding, some speciﬁc genomic loci in human breast cancer andﬂy embryos became less dynamic when activelytranscribed (Germier et al. 2017; Chen et al.
2018). Furthermore, it was recently shown that
active RNA polymerase I (RNAPI) also forms
clusters (Andrews et al. 2018) and constrains
ribosomal DNA chromatin in the nucleoli (Ideet al. 2020). In this case, transcription inhibition
dissociated RNAPI from rDNA chromatin andmoved like a liquid within the newly formed
droplet in the nucleolus (Ide et al. 2020). The
concept that the transcriptional machinery constrains chromatin (Fig. 5D,E) is consistent withthe classic transcription factory hypothesis(Cook 1999; Edelman and Fraser 2012; Feuerborn and Cook 2015), where clusters of RNAPII
and other transcription factors immobilize genome chromatin to facilitate transcription. Recent studies also suggest that RNAPII, Mediator,and other factors form condensate/clusters
upon transcription (Cho et al. 2018; Chonget al. 2018; Lu et al. 2018; Sabari et al. 2018).
Association with Chromatin Domains

Other prominent chromatin constraining factors are the proteins associated with heterochromatin, where the chromatin is less mobile (Fig.
4F; Chubb et al. 2002; Shinkai et al. 2016; Nozaki
et al. 2017; Nagashima et al. 2019; Lerner et al.
2020). The nuclear periphery lamina-associatedchromatin domains (LADs) enriched regions(van Steensel and Belmont 2017) have more heterochromatin marker di- and trimethylation ofhistone H3 lysine 9 (H3K9me2/3) and HP1 protein (Machida et al. 2018). Inner nuclear membrane proteins (Pawar and Kutay 2021) such as
lamins (Reddy 2021), together with HP1, mayconstrain chromatin movement (Fig. 4F; Chubb
et al. 2002; Shinkai et al. 2016; Nozaki et al. 2017;
Nagashima et al. 2019; Lerner et al. 2020). The
pericentric heterochromatin regions in mouse
cells (Maison et al. 2010) are also enrichedwith the H3K9me2/3 and HP1 protein, which
can cross-link nucleosomes (Machida et al.

2018) and restrict chromatin motion (Fig. 4F;
Nozaki et al. 2017; Lerner et al. 2020). Whereas
HP1 was suggested to form liquid droplets by
LLPS (Larson et al. 2017; Strom et al. 2017),
Erdel et al. (2020) recently reported that LLPS
may not be a prerequisite for the chromatincross-link function of HP1 (for details, see
Rippe 2021). Whether condensates/clusters/liquid droplets of other nuclear bodies can stabilize
the genome chromatin for their functions remains unclear and an area to be investigated.
PERSPECTIVES

The physical nature of chromatin in the nucleus,
discussed in this review, is functionally important for gene regulation and control of gene expression. Therefore, defects in the physical nature may be relevant for physiological andpathological processes (Misteli 2010; see Khan
2021). It is known that mutations and transcriptional misregulation of several “global genome
organizers,” such as the cohesin complex (Misteli 2010; Kline et al. 2018), are linked to humandiseases. Interestingly, some mutations in core
histones act as oncogenic drivers, creating theterm “oncohistones” (Bennett et al. 2019; Nacev
et al. 2019). For example, several mutations in
the histone variant H3.3 have been reported inpediatric gliomas (Schwartzentruber et al. 2012;
Wu et al. 2012). Among these, the dominantnegative H3.3 K27M mutant, where the 27th Lys
is replaced by Met, inhibits the activity of the
Polycomb repressor complex 2 (PRC2) to methylate H3K27, resulting in impaired transcriptional silencing (Chan et al. 2013; Lewis et al.
2013). H2B Glu76 and H3.1 Glu97 mutations
are also frequently found in cancer cells, andreplacement of these residues with Lys leads tonucleosome instability (Arimura et al. 2018).
Furthermore, mutations in the genes that encode several linker histone H1 isoforms arehighly recurrent in B-cell lymphomas (Yusufova
et al. 2021). Whereas how these mutations enhance tumorigenesis remains unclear, theirresultant altered behaviors may be involved in
the process. Such alterations might affect transcription, DNA repair efﬁciency, or plasticity ofchromatin in the nucleus (Shimamoto et al.

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

181

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.

2017; Stephens et al. 2017; Nagashima et al.
2019; also see Miroshnikova and Wickström
2021). Discovering how altering the physicalproperties of chromatin contributes to creatingcell abnormalities and subsequent human disorders, including tumorigenesis, is an exciting
area for future research.
ACKNOWLEDGMENTS

We are grateful to Dr. Y. Hiromi, Dr. Y. Itoh, and
Dr. K.M. Marshall for critical reading and editing of this manuscript. We thank Dr. M. Sasai,Dr. Belmont, Dr. Rando, and Maeshima laboratory members for helpful discussions andsupport. We apologize that space limitations
rendered us unable to mention many importantworks and papers on chromatin structure anddynamics. This work was supported by the Japan Society for the Promotion of Science (JSPS)
and MEXT KAKENHI Grants (19H05273 and
20H05936), a Japan Science and Technology
Agency CREST Grant (JPMJCR15G2), theTakeda Science Foundation, and the Uehara
Memorial Foundation.
REFERENCES
Reference is also in this collection.

Agbleke AA, Amitai A, Buenrostro JD, Chakrabarti A, Chu
L, Hansen AS, Koenig KM, Labade AS, Liu S, Nozaki T, etal. 2020. Advances in chromatin and chromosome research: perspectives from multiple ﬁelds. Mol Cell 79:
881–901. doi:10.1016/j.molcel.2020.07.003
Albiez H, Cremer M, Tiberi C, Vecchio L, Schermelleh L,
Dittrich S, Küpper K, Joffe B, Thormeyer T, von Hase J, etal. 2006. Chromatin domains and the interchromatincompartment form structurally deﬁned and functionallyinteracting nuclear networks. Chromosome Res 14: 707–
733. doi:10.1007/s10577-006-1086-x
Amitai A, Seeber A, Gasser SM, Holcman D. 2017. Visualization of chromatin decompaction and break site extrusion as predicted by statistical polymer modeling of
single-locus trajectories. Cell Rep 18: 1200–1214. doi:10.1016/j.celrep.2017.01.018
Andrews JO, Conway W, Cho WK, Narayanan A, Spille JH,
Jayanth N, Inoue T, Mullen S, Thaler J, Cissé II. 2018. qSR:
a quantitative super-resolution analysis tool reveals thecell-cycle dependent organization of RNA polymerase I
in live human cells. Sci Rep 8: 7424. doi:10.1038/s41598018-25454-0
Arai R, Sugawara T, Sato Y, Minakuchi Y, Toyoda A, Nabeshima K, Kimura H, Kimura A. 2017. Reduction in chromosome mobility accompanies nuclear organization dur-

182

ing early embryogenesis in Caenorhabditis elegans. Sci
Rep 7: 3631. doi:10.1038/s41598-017-03483-5
Arimura Y, Ikura M, Fujita R, Noda M, Kobayashi W, Horikoshi N, Sun J, Shi L, Kusakabe M, Harata M, et al. 2018.
Cancer-associated mutations of histones H2B, H3.1 andH2A.Z.1 affect the structure and stability of the nucleosome. Nucleic Acids Res 46: 10007–10018.
Asakura S, Oosawa F. 1954. On interaction between twobodies immersed in a solution of macromolecules. J
Chem Phys 22: 1255–1256. doi:10.1063/1.1740347
Ashwin SS, Nozaki T, Maeshima K, Sasai M. 2019. Organization of fast and slow chromatin revealed by singlenucleosome dynamics. Proc Natl Acad Sci 116: 19939–
19944. doi:10.1073/pnas.1907342116
Ashwin SS, Maeshima K, Sasai M. 2020. Heterogeneous ﬂuid-like movements of chromatin and their implications
to transcription. Biophys Rev 12: 461–468. doi:10.1007/
s12551-020-00675-8
Ausio J, Dong F, van Holde KE. 1989. Use of selectively
trypsinized nucleosome core particles to analyze therole of the histone “tails” in the stabilization of the nucleosome. J Mol Biol 206: 451–463. doi:10.1016/0022-2836(89)90493-2
Babokhov M, Hibino K, Itoh Y, Maeshima K. 2020. Local
chromatin motion and transcription. J Mol Biol 432: 694–
700. doi:10.1016/j.jmb.2019.10.018
Bajpai G, Jain I, Inamdar MM, Das D, Padinhateeri R. 2017.
Binding of DNA-bending non-histone proteins destabilizes regular 30-nm chromatin structure. PLoS Comput
Biol 13: e1005365. doi:10.1371/journal.pcbi.1005365
Baldi S, Korber P, Becker PB. 2020. Beads on a string-nucleosome array arrangements and folding of the chromatinﬁber. Nat Struct Mol Biol 27: 109–118. doi:10.1038/
s41594-019-0368-x
Bednar J, Garcia-Saez I, Boopathi R, Cutter AR, Papai G,
Reymer A, Syed SH, Lone IN, Tonchev O, Cruciﬁx C, etal. 2017. Structure and dynamics of a 197 bp nucleosome
in complex with linker histone H1. Mol Cell 66: 729.
doi:10.1016/j.molcel.2017.05.018
Bennett RL, Bele A, Small EC, Will CM, Nabet B, Oyer JA,
Huang X, Ghosh RP, Grzybowski AT, Yu T, et al. 2019. A
mutation in histone H2B represents a new class of oncogenic driver. Cancer Discov 9: 1438–1451. doi:10.1158/
2159-8290.CD-19-0393
Bian Q, Belmont AS. 2012. Revisiting higher-order andlarge-scale chromatin organization. Curr Opin Cell Biol
24: 359–366. doi:10.1016/j.ceb.2012.03.003
Bianconi E, Piovesan A, Facchin F, Beraudi A, Casadei R,
Frabetti F, Vitale L, Pelleri MC, Tassani S, Piva F, et al.
2013. An estimation of the number of cells in the humanbody. Annals Hum Biol 40: 463–471. doi:10.3109/
03014460.2013.807878
Bintu B, Mateo LJ, Su JH, Sinnott-Armstrong NA, Parker M,
Kinrot S, Yamaya K, Boettiger AN, Zhuang X. 2018. Super-resolution chromatin tracing reveals domains andcooperative interactions in single cells. Science 362:
eaau1783. doi:10.1126/science.aau1783
Cai S, Chen C, Tan ZY, Huang Y, Shi J, Gan L. 2018. Cryo-ET
reveals the macromolecular reorganization of S. pombe
mitotic chromosomes in vivo. Proc Natl Acad Sci 115:
10977–10982. doi:10.1073/pnas.1720476115

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

Physical Nature of Chromatin

Chan KM, Fang D, Gan H, Hashizume R, Yu C, Schroeder
M, Gupta N, Mueller S, James CD, Jenkins R, et al. 2013.
The histone H3.3K27M mutation in pediatric glioma reprograms H3K27 methylation and gene expression.
Genes Dev 27: 985–990. doi:10.1101/gad.217778.113
Chen B, Gilbert LA, Cimini BA, Schnitzbauer J, Zhang W, Li
GW, Park J, Blackburn EH, Weissman JS, Qi LS, et al.
2013. Dynamic imaging of genomic loci in living humancells by an optimized CRISPR/Cas system. Cell 155:
1479–1491. doi:10.1016/j.cell.2013.12.001
Chen C, Lim HH, Shi J, Tamura S, Maeshima K, Surana U,
Gan L. 2016. Budding yeast chromatin is dispersed in a
crowded nucleoplasm in vivo. Mol Biol Cell 27: 3357–
3368. doi:10.1091/mbc.E16-07-0506
Chen H, Levo M, Barinov L, Fujioka M, Jaynes JB, Gregor T.
2018. Dynamic interplay between enhancer-promoter topology and gene activity. Nat Genet 50: 1296–1303. doi:10.1038/s41588-018-0175-z
Cho WK, Spille JH, Hecht M, Lee C, Li C, Grube V, Cisse II.
2018. Mediator and RNA polymerase II clusters associate
in transcription-dependent condensates. Science 361:
412–415. doi:10.1126/science.aar4199
Chong S, Dugast-Darzacq C, Liu Z, Dong P, Dailey GM,
Cattoglio C, Heckert A, Banala S, Lavis L, Darzacq X, etal. 2018. Imaging dynamic and selective low-complexity
domain interactions that control gene transcription. Science 361: eaar2555. doi:10.1126/science.aar2555
Chubb JR, Boyle S, Perry P, Bickmore WA. 2002. Chromatin
motion is constrained by association with nuclear compartments in human cells. Curr Biol 12: 439–445. doi:10.1016/S0960-9822(02)00695-4
Church GM, Gao Y, Kosuri S. 2012. Next-generation digital
information storage in DNA. Science 337: 1628. doi:10.1126/science.1226355
Cole A. 1967. Chromosome structure. Theoret Biophys 1:
305–375.
Collepardo-Guevara R, Schlick T. 2014. Chromatin ﬁber
polymorphism triggered by variations of DNA linker
lengths. Proc Natl Acad Sci 111: 8061–8066. doi:10.1073/pnas.1315872111
Cook PR. 1999. The organization of replication and transcription. Science 284: 1790–1795. doi:10.1126/science.284.5421.1790
Cremer T, Cremer C. 2001. Chromosome territories, nuclear
architecture and gene regulation in mammalian cells. Nat
Rev Genet 2: 292–301. doi:10.1038/35066075
Davey CA, Sargent DF, Luger K, Maeder AW, Richmond TJ.
2002. Solvent mediated interactions in the structure of thenucleosome core particle at 1.9Å resolution. J Mol Biol
319: 1097–1113. doi:10.1016/S0022-2836(02)00386-8
Dekker J. 2008. Mapping in vivo chromatin interactions inyeast suggests an extended chromatin ﬁber with regionalvariation in compaction. J Biol Chem 283: 34532–34540.
doi:10.1074/jbc.M806479200
 Dekker J. 2021. 3D chromatin organization. Cold Spring
Harb Perspect Biol doi:10.1101/cshperspect.a040147
Dekker J, Heard E. 2015. Structural and functional diversityof topologically associating domains. FEBS Lett 589:
2877–2884. doi:10.1016/j.febslet.2015.08.044

Dion V, Gasser SM. 2013. Chromatin movement in themaintenance of genome stability. Cell 152: 1355–1364.
doi:10.1016/j.cell.2013.02.010
Dion V, Kalck V, Seeber A, Schleker T, Gasser SM. 2013.
Cohesin and the nucleolus constrain the mobility of spontaneous repair foci. EMBO Rep 14: 984–991. doi:10.1038/
embor.2013.142
Dixon JR, Selvaraj S, Yue F, Kim A, Li Y, Shen Y, Hu M, Liu
JS, Ren B. 2012. Topological domains in mammalian genomes identiﬁed by analysis of chromatin interactions.
Nature 485: 376–380. doi:10.1038/nature11082
Dorigo B, Schalch T, Bystricky K, Richmond TJ. 2003. Chromatin ﬁber folding: requirement for the histone H4
N-terminal tail. J Mol Biol 327: 85–96. doi:10.1016/
S0022-2836(03)00025-1
Dubochet J, Adrian M, Schultz P, Oudet P. 1986. Cryo-electron microscopy of vitriﬁed SV40 minichromosomes: theliquid drop model. EMBO J 5: 519–528. doi:10.1002/j.1460-2075.1986.tb04241.x
Earnshaw WC, Laemmli UK. 1983. Architecture of metaphase chromosomes and chromosome scaffolds. J Cell
Biol 96: 84–93. doi:10.1083/jcb.96.1.84
Edelman LB, Fraser P. 2012. Transcription factories: genetic
programming in three dimensions. Curr Opin Genet Dev
22: 110–114. doi:10.1016/j.gde.2012.01.010
Eltsov M, Maclellan KM, Maeshima K, Frangakis AS, Dubochet J. 2008. Analysis of cryo-electron microscopy images does not support the existence of 30-nm chromatinﬁbers in mitotic chromosomes in situ. Proc Natl Acad Sci
105: 19732–19737. doi:10.1073/pnas.0810057105
Erdel F, Rademacher A, Vlijm R, Tunnermann J, Frank L,
Weinmann R, Schweigert E, Yserentant K, Hummert J,
Bauer C, et al. 2020. Mouse heterochromatin adopts digital compaction states without showing hallmarks of
HP1-driven liquid-liquid phase separation. Mol Cell 78:
1236–249.e7.
Feuerborn A, Cook PR. 2015. Why the activity of a genedepends on its neighbors. Trends Genet 31: 483–490.
doi:10.1016/j.tig.2015.07.001
Finch JT, Klug A. 1976. Solenoidal model for superstructurein chromatin. Proc Natl Acad Sci 73: 1897–1901. doi:10.1073/pnas.73.6.1897
Fussner E, Strauss M, Djuric U, Li R, Ahmed K, Hart M, Ellis
J, Bazett-Jones DP. 2012. Open and closed domains in themouse genome are conﬁgured as 10-nm chromatin ﬁbres.
EMBO Rep 13: 992–996. doi:10.1038/embor.2012.139
Germier T, Kocanova S, Walther N, Bancaud A, Shaban HA,
Sellou H, Politi AZ, Ellenberg J, Gallardo F, Bystricky K.
2017. Real-time imaging of a single gene reveals transcription-initiated local conﬁnement. Biophys J 113: 1383–
1394. doi:10.1016/j.bpj.2017.08.014
Gibson BA, Doolittle LK, Schneider MWG, Jensen LE, Gamarra N, Henry L, Gerlich DW, Redding S, Rosen MK.
2019. Organization of chromatin by intrinsic and regulated phase separation. Cell 179: 470–484.e21. doi:10.1016/j.cell.2019.08.037
Gómez-García PA, Portillo-Ledesma S, Neguembor MV,
Pesaresi M, Oweis W, Rohrlich T, Wieser S, Meshorer
E, Schlick T, Cosma MP, et al. 2021. Mesoscale modelingand single-nucleosome tracking reveal remodeling of
clutch folding and dynamics in stem cell differentiation.
Cell Rep 34: 108614. doi:10.1016/j.celrep.2020.108614

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

183

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.

Gorisch SM, Wachsmuth M, Toth KF, Lichter P, Rippe K.
2005. Histone acetylation increases chromatin accessibility. J Cell Sci 118: 5825–5834. doi:10.1242/jcs.02689
Gu B, Swigut T, Spencley A, Bauer MR, Chung M, Meyer T,
Wysocka J. 2018. Transcription-coupled changes in nuclear mobility of mammalian cis-regulatory elements. Science 359: 1050–1055. doi:10.1126/science.aao3136
Guo YE, Manteiga JC, Henninger JE, Sabari BR, Dall’Agnese
A, Hannett NM, Spille JH, Afeyan LK, Zamudio AV,
Shrinivas K, et al. 2019. Pol II phosphorylation regulates
a switch between transcriptional and splicing condensates. Nature 572: 543–548. doi:10.1038/s41586-0191464-0
Hajjoul H, Mathon J, Ranchon H, Goiffon I, Mozziconacci J,
Albert B, Carrivain P, Victor JM, Gadal O, Bystricky K, etal. 2013. High-throughput chromatin motion tracking inliving yeast reveals the ﬂexibility of the ﬁber throughout
the genome. Genome Res 23: 1829–1838. doi:10.1101/gr.157008.113
Hancock R. 2007. Packing of the polynucleosome chain ininterphase chromosomes: evidence for a contribution of
crowding and entropic forces. Semin Cell Dev Biol 18:
668–675. doi:10.1016/j.semcdb.2007.08.006
Hansen JC. 2002. Conformational dynamics of the chromatin ﬁber in solution: determinants, mechanisms, andfunctions. Annu Rev Biophys Biomol Struct 31: 361–
392. doi:10.1146/annurev.biophys.31.101101.140858
Hansen JC, Connolly M, McDonald CJ, Pan A, Pryamkova
A, Ray K, Seidel E, Tamura S, Rogge R, Maeshima K. 2018.
The 10-nm chromatin ﬁber and its relationship to interphase chromosome organization. Biochem Soc Trans 46:
67–76. doi:10.1042/BST20170101
Hendzel MJ, Boisvert F, Bazett-Jones DP. 1999. Direct visualization of a protein nuclear architecture. Mol Biol Cell
10: 2051–2062. doi:10.1091/mbc.10.6.2051
Heun P, Laroche T, Shimada K, Furrer P, Gasser SM. 2001.
Chromosome dynamics in the yeast interphase nucleus.
Science 294: 2181–2186. doi:10.1126/science.1065366
Hihara S, Pack CG, Kaizu K, Tani T, Hanafusa T, Nozaki T,
Takemoto S, Yoshimi T, Yokota H, Imamoto N, et al.
2012. Local nucleosome dynamics facilitate chromatin
accessibility in living mammalian cells. Cell Rep 2:
1645–1656. doi:10.1016/j.celrep.2012.11.008
Hoffman DP, Shtengel G, Xu CS, Campbell KR, Freeman M,
Wang L, Milkie DE, Pasolli HA, Iyer N, Bogovic JA, et al.
2020. Correlative three-dimensional super-resolution andblock-face electron microscopy of whole vitreously frozen
cells. Science 367: eaaz5357. doi:10.1126/science.aaz5357
Hsieh TH, Weiner A, Lajoie B, Dekker J, Friedman N, Rando
OJ. 2015. Mapping nucleosome resolution chromosome
folding in yeast by micro-C. Cell 162: 108–119. doi:10.1016/j.cell.2015.05.048
Hu Y, Kireev I, Plutz M, Ashourian N, Belmont AS. 2009.
Large-scale chromatin structure of inducible genes: transcription on a condensed, linear template. J Cell Biol 185:
87–100. doi:10.1083/jcb.200809196
Hudson DF, Vagnarelli P, Gassmann R, Earnshaw WC.
2003. Condensin is required for nonhistone protein assembly and structural integrity of vertebrate mitotic chromosomes. Dev Cell 5: 323–336. doi:10.1016/S1534-5807(03)00199-0

184

Hyman AA, Weber CA, Jülicher F. 2014. Liquid–liquid
phase separation in biology. Ann Rev Cell Dev Biol 30:
39–58. doi:10.1146/annurev-cellbio-100913-013325
Ide S, Imai R, Ochi H, Maeshima K. 2020. Transcriptional
suppression of ribosomal DNA with phase separation. Sci
Adv 6: eabb5953. doi:10.1126/sciadv.abb5953
Imai R, Nozaki T, Tani T, Kaizu K, Hibino K, Ide S, Tamura
S, Takahashi K, Shribak M, Maeshima K. 2017. Density
imaging of heterochromatin in live cells using orientation-independent-DIC microscopy. Mol Biol Cell 28:
3349–3359. doi:10.1091/mbc.e17-06-0359
Itoh Y, Iida S, Tamura S, Nagashima R, Shiraki K, Goto T,
Hibino K, Ide S, Maeshima K. 2021. 1,6-hexanediol rapidly immobilizes and condenses chromatin in living human cells. Life Sci Alliance 4: e202001005. doi:10.26508/
lsa.202001005
Joti Y, Hikima T, Nishino Y, Kamada F, Hihara S, Takata H,
Ishikawa T, Maeshima K. 2012. Chromosomes without a
30-nm chromatin ﬁber. Nucleus 3: 404–410. doi:10.4161/
nucl.21222
Kan PY, Caterino TL, Hayes JJ. 2009. The H4 tail domain
participates in intra- and internucleosome interactions
with protein and DNA during folding and oligomerization of nucleosome arrays. Mol Cell Biol 29: 538–546.
doi:10.1128/MCB.01343-08
 Khan J. 2021. Chromatin in cancer. Cold Spring Harb Perspect Biol doi:10.1101/cshperspect.a040956
Kieffer-Kwon KR, Nimura K, Rao SSP, Xu J, Jung S, Pekowska A, Dose M, Stevens E, Mathe E, Dong P, et al. 2017.
Myc regulates chromatin decompaction and nuclear architecture during B cell activation. Mol Cell 67: 566–578.
e10. doi:10.1016/j.molcel.2017.07.013
Kimura H, Cook PR. 2001. Kinetics of core histones in living
human cells: little exchange of H3 and H4 and some rapid
exchange of H2B. J Cell Biol 153: 1341–1354. doi:10.1083/
jcb.153.7.1341
Kline AD, Moss JF, Selicorni A, Bisgaard AM, Deardorff
MA, Gillett PM, Ishman SL, Kerr LM, Levin AV, Mulder
PA, et al. 2018. Diagnosis and management of Cornelia deLange syndrome: ﬁrst international consensus statement.
Nat Rev Genet 19: 649–666. doi:10.1038/s41576-0180031-0
Kornberg RD. 1974. Chromatin structure: a repeating unit ofhistones and DNA. Science 184: 868–871. doi:10.1126/
science.184.4139.868
Krietenstein N, Rando OJ. 2020. Mesoscale organization of
the chromatin ﬁber. Curr Opin Genet Dev 61: 32–36.
doi:10.1016/j.gde.2020.02.022
Larson AG, Elnatan D, Keenen MM, Trnka MJ, Johnston JB,
Burlingame AL, Agard DA, Redding S, Narlikar GJ. 2017.
Liquid droplet formation by HP1α suggests a role forphase separation in heterochromatin. Nature 547: 236–
240. doi:10.1038/nature22822
Lerner J, Gomez-Garcia PA, McCarthy RL, Liu Z, Lakadamyali M, Zaret KS. 2020. Two-parameter mobility assessments discriminate diverse regulatory factor behaviors inchromatin. Mol Cell 79: 677–688.e6. doi:10.1016/j.molcel.2020.05.036
Levi V, Ruan Q, Plutz M, Belmont AS, Gratton E. 2005.
Chromatin dynamics in interphase cells revealed bytracking in a two-photon excitation microscope. Biophys
J 89: 4275–4285. doi:10.1529/biophysj.105.066670

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

Physical Nature of Chromatin

Lewis PW, Müller MM, Koletsky MS, Cordero F, Lin S,
Banaszynski LA, Garcia BA, Muir TW, Becher OJ, Allis
CD. 2013. Inhibition of PRC2 activity by a gain-of-function H3 mutation found in pediatric glioblastoma. Science
340: 857–861. doi:10.1126/science.1232245
Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev
M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ,
Dorschner MO, et al. 2009. Comprehensive mapping of
long-range interactions reveals folding principles of thehuman genome. Science 326: 289–293. doi:10.1126/sci
ence.1181369
Lin Y, Protter DS, Rosen MK, Parker R. 2015. Formation andmaturation of phase-separated liquid droplets by RNAbinding proteins. Mol Cell 60: 208–219. doi:10.1016/j.molcel.2015.08.018
Lu H, Yu D, Hansen AS, Ganguly S, Liu R, Heckert A,
Darzacq X, Zhou Q. 2018. Phase-separation mechanism
for C-terminal hyperphosphorylation of RNA polymerase II. Nature 558: 318–323. doi:10.1038/s41586-0180174-3
Luger K, Mäder AW, Richmond RK, Sargent DF, RichmondTJ. 1997. Crystal structure of the nucleosome core particle
at 2.8 Å resolution. Nature 389: 251–260. doi:10.1038/
38444
Ma H, Tu LC, Chung YC, Naseri A, Grunwald D, Zhang
S, Pederson T. 2019. Cell cycle- and genomic
distance-dependent dynamics of a discrete chromosomal
region. J Cell Biol 218: 1467–1477. doi:10.1083/jcb.201807162
Machida S, Takizawa Y, Ishimaru M, Sugita Y, Sekine S,
Nakayama JI, Wolf M, Kurumizaka H. 2018. Structural
basis of heterochromatin formation by human HP1. Mol
Cell 69: 385–397.e8. doi:10.1016/j.molcel.2017.12.011
Maeshima K, Hihara S, Eltsov M. 2010. Chromatin structure:
does the 30-nm ﬁbre exist in vivo? Curr Opin Cell Biol 22:
291–297. doi:10.1016/j.ceb.2010.03.001
Maeshima K, Imai R, Tamura S, Nozaki T. 2014. Chromatin
as dynamic 10-nm ﬁbers. Chromosoma 123: 225–237.
doi:10.1007/s00412-014-0460-2
Maeshima K, Kaizu K, Tamura S, Nozaki T, Kokubo T,
Takahashi K. 2015. The physical size of transcription factors is key to transcriptional regulation in the chromatin
domains. J Phys 27: 064116.
Maeshima K, Ide S, Hibino K, Sasai M. 2016a. Liquid-likebehavior of chromatin. Curr Opin Genet Dev 37: 36–45.
doi:10.1016/j.gde.2015.11.006
Maeshima K, Rogge R, Tamura S, Joti Y, Hikima T, Szerlong
H, Krause C, Herman J, Seidel E, DeLuca J, et al. 2016b.
Nucleosomal arrays self-assemble into supramolecularglobular structures lacking 30-nm ﬁbers. EMBO J 35:
1115–1132. doi:10.15252/embj.201592660
Maeshima K, Matsuda T, Shindo Y, Imamura H, Tamura S,
Imai R, Kawakami S, Nagashima R, Soga T, Noji H, et al.
2018a. A transient rise in free Mg2+ ions released from
ATP-Mg hydrolysis contributes to mitotic chromosome
condensation. Curr Biol 28: 444–451.e6. doi:10.1016/j.cub.2017.12.035
Maeshima K, Tamura S, Shimamoto Y. 2018b. Chromatin asa nuclear spring. Biophys Physicobiol 15: 189–195. doi:10.2142/biophysico.15.0_189
Maeshima K, Tamura S, Hansen JC, Itoh Y. 2020. Fluid-likechromatin: toward understanding the real chromatin or-

ganization present in the cell. Curr Opin Cell Biol 64: 77–
89. doi:10.1016/j.ceb.2020.02.016
Maison C, Quivy JP, Probst AV, Almouzni G. 2010. Heterochromatin at mouse pericentromeres: a model for de novo
heterochromatin formation and duplication during replication. Cold Spring Harb Symp Quant Biol 75: 155–165.
doi:10.1101/sqb.2010.75.013
Marenduzzo D, Finan K, Cook PR. 2006. The depletion
attraction: an underappreciated force driving cellular organization. J Cell Biol 175: 681–686. doi:10.1083/jcb.200609066
Marshall WF, Straight A, Marko JF, Swedlow J, Dernburg A,
Belmont A, Murray AW, Agard DA, Sedat JW. 1997. Interphase chromosomes undergo constrained diffusionalmotion in living cells. Curr Biol 7: 930–939. doi:10.1016/
S0960-9822(06)00412-X
McDowall AW, Smith JM, Dubochet J. 1986. Cryo-electron
microscopy of vitriﬁed chromosomes in situ. EMBO J 5:
1395–1402. doi:10.1002/j.1460-2075.1986.tb04373.x
Meister P, Towbin BD, Pike BL, Ponti A, Gasser SM. 2010.
The spatial dynamics of tissue-speciﬁc promoters during
C. elegans development. Genes Dev 24: 766–782. doi:10.1101/gad.559610
Meshorer E, Yellajoshula D, George E, Scambler PJ, Brown
DT, Misteli T. 2006. Hyperdynamic plasticity of chromatin proteins in pluripotent embryonic stem cells. Dev Cell
10: 105–116. doi:10.1016/j.devcel.2005.10.017
Miron E, Oldenkamp R, Brown JM, Pinto DMS, Xu CS, Faria
AR, Shaban HA, Rhodes JDP, Innocent C, de Ornellas S,
et al. 2020. Chromatin arranges in chains of mesoscaledomains with nanoscale functional topography independent of cohesin. Sci Adv 6: eaba8811. doi:10.1126/sciadv.aba8811
 Miroshnikova YA, Wickström SA. 2021. Mechanical forcesin nuclear organization. Cold Spring Harb Perspect Biol
doi:10.1101/cshperspect.a039685
Misteli T. 2010. Higher-order genome organization in human disease. Cold Spring Harb Perspect Biol 2: a000794.
doi:10.1101/cshperspect.a000794
Misteli T. 2020. The self-organizing genome: principles ofgenome architecture and function. Cell 183: 28–45. doi:10.1016/j.cell.2020.09.014
Misteli T, Gunjan A, Hock R, Bustin M, Brown DT. 2000.
Dynamic binding of histone H1 to chromatin in living
cells. Nature 408: 877–881. doi:10.1038/35048610
Mitrea DM, Kriwacki RW. 2016. Phase separation in biology;functional organization of a higher order. Cell Commun
Signal 14: 1. doi:10.1186/s12964-015-0125-7
Morales C, Losada A. 2018. Establishing and dissolving cohesion during the vertebrate cell cycle. Curr Opin Cell Biol
52: 51–57. doi:10.1016/j.ceb.2018.01.010
Nacev BA, Feng L, Bagert JD, Lemiesz AE, Gao J, Soshnev
AA, Kundra R, Schultz N, Muir TW, Allis CD. 2019. The
expanding landscape of “oncohistone” mutations in human cancers. Nature 567: 473–478. doi:10.1038/s41586019-1038-1
Nagashima R, Hibino K, Ashwin SS, Babokhov M, Fujishiro
S, Imai R, Nozaki T, Tamura S, Tani T, Kimura H, et al.
2019. Single nucleosome imaging reveals loose genome
chromatin networks via active RNA polymerase II. J Cell
Biol 218: 1511–1530. doi:10.1083/jcb.201811090

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

185

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.

Nasmyth K, Haering CH. 2005. The structure andfunction of SMC and kleisin complexes. Annu Rev Biochem 74: 595–648. doi:10.1146/annurev.biochem.74.082803.133219
Nishino Y, Eltsov M, Joti Y, Ito K, Takata H, Takahashi Y,
Hihara S, Frangakis AS, Imamoto N, Ishikawa T, et al.
2012. Human mitotic chromosomes consist predominantly of irregularly folded nucleosome ﬁbres without a
30-nm chromatin structure. EMBO J 31: 1644–1653.
doi:10.1038/emboj.2012.35
Nishiyama T. 2019. Cohesion and cohesin-dependent chromatin organization. Curr Opin Cell Biol 58: 8–14. doi:10.1016/j.ceb.2018.11.006
Nora EP, Lajoie BR, Schulz EG, Giorgetti L, Okamoto I,
Servant N, Piolot T, van Berlum NL, Meisig J, Sedat JW,
et al. 2012. Spatial partitioning of the regulatory landscapeof the X-inactivation centre. Nature 485: 381–385. doi:10.1038/nature11049
Nozaki T, Imai R, Tanbo M, Nagashima R, Tamura S, Tani T,
Joti Y, Tomita M, Hibino K, Kanemaki MT, et al. 2017.
Dynamic organization of chromatin domains revealed bysuper-resolution live-cell imaging. Mol Cell 67: 282–293.
e7. doi:10.1016/j.molcel.2017.06.018
Nozaki T, Hudson DF, Tamura S, Maeshima K. 2018. Dynamic chromatin folding in the cell. In Nuclear architecture and dynamics (ed. C Lavelle, JM Victor), pp. 101–
122. Academic, New York.
Nozawa RS, Gilbert N. 2019. RNA: nuclear glue for folding
the genome. Trends Cell Biol 29: 201–211. doi:10.1016/j.tcb.2018.12.003
Nozawa RS, Boteva L, Soares DC, Naughton C, Dun AR,
Buckle A, Ramsahoye B, Bruton PC, Saleeb RS, Arnedo
M, et al. 2017. SAF-A regulates interphase chromosome
structure through oligomerization with chromatin-associated RNAs. Cell 169: 1214–1227.e18. doi:10.1016/j.cell.2017.05.029
Ohno M, Ando T, Priest DG, Kumar V, Yoshida Y, Taniguchi Y. 2019. Sub-nucleosomal genome structure reveals
distinct nucleosome folding motifs. Cell 176: 520–534.
e25. doi:10.1016/j.cell.2018.12.014
Olins AL, Olins DE. 1974. Spheroid chromatin units(v bodies). Science 183: 330–332. doi:10.1126/science.183.4122.330
Olins DE, Olins AL. 2003. Chromatin history: our view fromthe bridge. Nat Rev Mol Cell Biol 4: 809–814. doi:10.1038/
nrm1225
Olins DE, Olins AL. 2018. Epichromatin and chromomeres:a “fuzzy” perspective. Open Biol 8: 180058. doi:10.1098/
rsob.180058
Ou HD, Phan S, Deerinck TJ, Thor A, Ellisman MH, O’Shea
CC. 2017. ChromEMT: visualizing 3D chromatin structure and compaction in interphase and mitotic cells. Science 357: eaag0025. doi:10.1126/science.aag0025
 Pawar S, Kutay U. 2021. The diverse cellular functions of
inner nuclear membrane proteins. Cold Spring Harb Perspect Biol doi:10.1101/cshperspect.a040477.
Pecqueur C, Oliver L, Oizel K, Lalier L, Vallette FM. 2013.
Targeting metabolism to induce cell death in cancer cells
and cancer stem cells. Int J Cell Biol 2013: 805975. doi:10.1155/2013/805975

186

Pepenella S, Murphy KJ, Hayes JJ. 2014a. Intra- and internucleosome interactions of the core histone tail domains
in higher-order chromatin structure. Chromosoma 123:
3–13. doi:10.1007/s00412-013-0435-8
Pepenella S, Murphy KJ, Hayes JJ. 2014b. A distinct switch ininteractions of the histone H4 tail domain upon salt-dependent folding of nucleosome arrays. J Biol Chem 289:
27342–27351. doi:10.1074/jbc.M114.595140
Qian Y, Wang X, Liu Y, Li Y, Colvin RA, Tong L, Wu S, Chen
X. 2014. Extracellular ATP is internalized by macropinocytosis and induces intracellular ATP increase and drug
resistance in cancer cells. Cancer Lett 351: 242–251.
doi:10.1016/j.canlet.2014.06.008
Rao SS, Huntley MH, Durand NC, Stamenova EK, Bochkov
ID, Robinson JT, Sanborn AL, Machol I, Omer AD,
Lander ES, et al. 2014. A 3D map of the human genome
at kilobase resolution reveals principles of chromatin
looping. Cell 159: 1665–1680. doi:10.1016/j.cell.2014.11.021
Rao SSP, Huang SC, Glenn St Hilaire B, Engreitz JM, Perez
EM, Kieffer-Kwon KR, Sanborn AL, Johnstone SE, Bascom GD, Bochkov ID, et al. 2017. Cohesin loss eliminates
all loop domains. Cell 171: 305–320.e24. doi:10.1016/j.cell.2017.09.026
 Reddy K. 2021. Nuclear lamina. Cold Spring Harb Perspect
Biol doi:10.1101/cshperspect.a040113.
Ricci MA, Manzo C, García-Parajo MF, Lakadamyali M,
Cosma MP. 2015. Chromatin ﬁbers are formed by heterogeneous groups of nucleosomes in vivo. Cell 160: 1145–
1158. doi:10.1016/j.cell.2015.01.054
 Rippe K. 2021. Phase separation in chromatin. Cold Spring
Harb Perspect Biol doi:10.1101/cshperspect.a040683
Risca VI, Denny SK, Straight AF, Greenleaf WJ. 2017. Variable chromatin structure revealed by in situ spatially correlated DNA cleavage mapping. Nature 541: 237–241.
doi:10.1038/nature20781
Robinett CC, Straight A, Li G, Willhelm C, Sudlow G,
Murray A, Belmont AS. 1996. In vivo localizationof DNA sequences and visualization of large-scalechromatin organization using lac operator/repressor recognition. J Cell Biol 135: 1685–1700. doi:10.1083/jcb.135.6.1685
Robinson PJ, Fairall L, Huynh VA, Rhodes D. 2006. EM
measurements deﬁne the dimensions of the “30-nm”
chromatin ﬁber: evidence for a compact, interdigitatedstructure. Proc Natl Acad Sci 103: 6506–6511. doi:10.1073/pnas.0601212103
Sabari BR, Dall’Agnese A, Boija A, Klein IA, Coffey
EL, Shrinivas K, Abraham BJ, Hannett NM, Zamudio
AV, Manteiga JC, et al. 2018. Coactivator condensation
at super-enhancers links phase separation and gene
control. Science 361: eaar3958. doi:10.1126/science.aar3958
Sanborn AL, Rao SS, Huang SC, Durand NC, Huntley MH,
Jewett AI, Bochkov ID, Chinnappan D, Cutkosky A, Li J,
et al. 2015. Chromatin extrusion explains key features ofloop and domain formation in wild-type and engineered
genomes. Proc Natl Acad Sci 112: E6456–E6465. doi:10.1073/pnas.1518552112
Sanulli S, Trnka MJ, Dharmarajan V, Tibble RW, Pascal BD,
Burlingame AL, Grifﬁn PR, Gross JD, Narlikar GJ. 2019.
HP1 reshapes nucleosome core to promote phase separa-

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

Physical Nature of Chromatintion of heterochromatin. Nature 575: 390–394. doi:10.1038/s41586-019-1669-2
Schalch T, Duda S, Sargent DF, Richmond TJ. 2005. X-ray
structure of a tetranucleosome and its implications for thechromatin ﬁbre. Nature 436: 138–141. doi:10.1038/na
ture03686
Schwartzentruber J, Korshunov A, Liu XY, Jones DT, Pfaff E,
Jacob K, Sturm D, Fontebasso AM, Quang DA, Tönjes M,
et al. 2012. Driver mutations in histone H3.3 and chromatin remodelling genes in paediatric glioblastoma. Nature 482: 226–231. doi:10.1038/nature10833
Seeber A, Hauer MH, Gasser SM. 2018. Chromosome dynamics in response to DNA damage. Annu Rev Genet 52:
295–319. doi:10.1146/annurev-genet-120417-031334
Sexton T, Yaffe E, Kenigsberg E, Bantignies F, Leblanc B,
Hoichman M, Parrinello H, Tanay A, Cavalli G. 2012.
Three-dimensional folding and functional organization
principles of the Drosophila genome. Cell 148: 458–472.
doi:10.1016/j.cell.2012.01.010
Shaban HA, Barth R, Recoules L, Bystricky K. 2020. Hi-D:
nanoscale mapping of nuclear dynamics in single living
cells. Genome Biol 21: 95. doi:10.1186/s13059-02002002-6
Shimamoto Y, Tamura S, Masumoto H, Maeshima K. 2017.
Nucleosome–nucleosome interactions via histone tails
and linker DNA regulate nuclear rigidity. Mol Biol Cell
28: 1580–1589. doi:10.1091/mbc.e16-11-0783
Shinkai S, Nozaki T, Maeshima K, Togashi Y. 2016. Dynamic
nucleosome movement provides structural informationof topological chromatin domains in living human cells.
PLoS Comput Biol 12: e1005136. doi:10.1371/journal.pcbi.1005136
Sinha D, Shogren-Knaak MA. 2010. Role of direct interactions between the histone H4 tail and the H2A core inlong range nucleosome contacts. J Biol Chem 285: 16572–
16581. doi:10.1074/jbc.M109.091298
Song F, Chen P, Sun D, Wang M, Dong L, Liang D, Xu RM,
Zhu P, Li G. 2014. Cryo-EM study of the chromatin ﬁber
reveals a double helix twisted by tetranucleosomal units.
Science 344: 376–380. doi:10.1126/science.1251413
Soutoglou E, Misteli T. 2007. Mobility and immobility ofchromatin in transcription and genome stability. Curr
Opin Genet Dev 17: 435–442. doi:10.1016/j.gde.2007.08.004
Stephens AD, Banigan EJ, Adam SA, Goldman RD, Marko
JF. 2017. Chromatin and lamin A determine two different
mechanical response regimes of the cell nucleus. Mol Biol
Cell.
Strickfaden H, Tolsma TO, Sharma A, Underhill DA, Hansen JC, Hendzel MJ. 2020. Condensed chromatin behaves
like a solid on the mesoscale in vitro and in living cells.
Cell 183: 1772–1784.e13. doi:10.1016/j.cell.2020.11.027
Strom AR, Emelyanov AV, Mir M, Fyodorov DV, Darzacq X,
Karpen GH. 2017. Phase separation drives heterochromatin domain formation. Nature 547: 241–245. doi:10.1038/
nature22989
Takata H, Hanafusa T, Mori T, Shimura M, Iida Y, Ishikawa
K, Yoshikawa K, Yoshikawa Y, Maeshima K. 2013. Chromatin compaction protects genomic DNA from radiation
damage. PLoS ONE 8: e75622. doi:10.1371/journal.pone.0075622

Tasan I, Sustackova G, Zhang L, Kim J, Sivaguru M,
HamediRad M, Wang Y, Genova J, Ma J, Belmont AS,
et al. 2018. CRISPR/Cas9-mediated knock-in of an
optimized TetO repeat for live cell imaging of endogenous loci. Nucleic Acids Res 46: e100. doi:10.1093/nar/gky501Toth KF, Knoch TA, Wachsmuth M, Frank-Stohr M, Stohr
M, Bacher CP, Muller G, Rippe K. 2004. Trichostatin Ainduced histone acetylation causes decondensation of interphase chromatin. J Cell Sci 117: 4277–4287. doi:10.1242/jcs.01293
Turner AL, Watson M, Wilkins OG, Cato L, Travers A,
Thomas JO, Stott K. 2018. Highly disordered histone
H1-DNA model complexes and their condensates. Proc
Natl Acad Sci 115: 11964–11969. doi:10.1073/pnas.1805943115
van Steensel B, Belmont AS. 2017. Lamina-associated domains: links with chromosome architecture, heterochromatin, and gene repression. Cell 169: 780–791. doi:10.1016/j.cell.2017.04.022
Visvanathan A, Ahmed K, Even-Faitelson L, Lleres D, Bazett-Jones DP, Lamond AI. 2013. Modulation ofhigher order chromatin conformation in mammaliancell nuclei can be mediated by polyamines and divalentcations. PLoS ONE 8: e67689. doi:10.1371/journal.pone.0067689
Wolffe AP, Hayes JJ. 1999. Chromatin disruption and modiﬁcation. Nucleic Acids Res 27: 711–720. doi:10.1093/nar/
27.3.711
Woodcock CL, Safer JP, Stanchﬁeld JE. 1976. Structural repeating units in chromatin. I: Evidence for their generaloccurrence. Exp Cell Res 97: 101–110. doi:10.1016/00144827(76)90659-5
Woodcock CL, Frado LL, Rattner JB. 1984. The higherorder structure of chromatin: evidence for a helicalribbon arrangement. J Cell Biol 99: 42–52. doi:10.1083/
jcb.99.1.42
Woodcock CL, Skoultchi AI, Fan Y. 2006. Role of linker
histone in chromatin structure and function: H1 stoichiometry and nucleosome repeat length. Chromosome Res
14: 17–25. doi:10.1007/s10577-005-1024-3
Wu G, Broniscer A, McEachron TA, Lu C, Paugh BS,
Becksfort J, Qu C, Ding L, Huether R, Parker M, et al.
2012. Somatic histone H3 alterations in pediatricdiffuse intrinsic pontine gliomas and non-brainstem
glioblastomas. Nat Genet 44: 251–253. doi:10.1038/ng.1102
Wutz G, Várnai C, Nagasaka K, Cisneros DA, Stocsits RR,
Tang W, Schoenfelder S, Jessberger G, Muhar M, Hossain
MJ, et al. 2017. Topologically associating domains andchromatin loops depend on cohesin and are regulatedby CTCF, WAPL, and PDS5 proteins. EMBO J 36:
3573–3599. doi:10.15252/embj.201798004
Xu J, Ma H, Jin J, Uttam S, Fu R, Huang Y, Liu Y. 2018.
Super-resolution imaging of higher-order chromatin
structures at different epigenomic states in single mammalian cells. Cell Rep 24: 873–882. doi:10.1016/j.celrep.2018.06.085
Yusufova N, Kloetgen A, Teater M, Osunsade A, Camarillo
JM, Chin CR, Doane AS, Venters BJ, Portillo-Ledesma S,
Conway J, et al. 2021. Histone H1 loss drives lymphoma

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.

187

This is a free sample of content from The Nucleus, Second Edition.
Click here for more information on how to buy the book.

K. Maeshima et al.
by disrupting 3D chromatin architecture. Nature 589:
299–305. doi:10.1038/s41586-020-3017-y
Zheng C, Lu X, Hansen JC, Hayes JJ. 2005. Salt-dependent
intra- and internucleosomal interactions of the H3 taildomain in a model oligonucleosomal array. J Biol Chem
280: 33552–33557. doi:10.1074/jbc.M507241200
Zidovska A, Weitz DA, Mitchison TJ. 2013. Micron-scalecoherence in interphase chromatin dynamics. Proc

188

Natl Acad Sci 110: 15555–15560. doi:10.1073/pnas.1220313110
Zuin J, Dixon JR, van der Reijden MIJA, Ye Z, Kolovos P,
Brouwer RWW, van de Corput MPC, van de Werken
HJG, Knoch TA, van IJcken WFJ, et al. 2014. Cohesin
and CTCF differentially affect chromatin architectureand gene expression in human cells. Proc Natl Acad Sci
111: 996–1001. doi:10.1073/pnas.1317788111

Cite this article as Cold Spring Harb Perspect Biol doi: 10.1101/cshperspect.a040675© 2022 by Cold Spring Harbor Laboratory Press. All rights reserved.