REVIEWS
EPIGENETICS
Chromatin dynamics during the
cell cycle at centromeres
Sebastian Müller1,2 and Geneviève Almouzni1,2
Abstract | Centromeric chromatin undergoes major changes in composition and architecture
during each cell cycle. These changes in specialized chromatin facilitate kinetochore formation
in mitosis to ensure proper chromosome segregation. Thus, proper orchestration of centromeric
chromatin dynamics during interphase, including replication in S phase, is crucial. We provide
the current view concerning the centromeric architecture associated with satellite repeat
sequences in mammals and its dynamics during the cell cycle. We summarize the contributions
of deposited histone variants and their chaperones, other centromeric components — including
proteins and their post-translational modifications, and RNAs — and we link the expression and
deposition timing of each component during the cell cycle. Because neocentromeres occur at
ectopic sites, we highlight how cell cycle processes can go wrong, leading to neocentromere
formation and potentially disease.
α-Satellite sequences
Tandem repeat DNA
sequences found at
centromeres. The sequences
are highly divergent among
species.
Neocentromeres
Ectopic centromeres that are
formed at loci other than the
usual α-satellite sequence.
Institut Curie, PSL Research
University, CNRS, UMR3664,
Equipe Labellisée Ligue
contre le Cancer,
F-75005 Paris, France.
2
Sorbonne Universités,
UPMC Université Paris 06,
CNRS, UMR3664, F-75005
Paris, France.
1
Correspondence to G.A.
[email protected]
doi:10.1038/nrg.2016.157
Published online 31 Jan 2017
Chromosomes undergo major changes in structure and
organization during the cell cycle (FIG. 1). They condense
during mitosis, and during this stage, as first described by
Walther Flemming in 1882 (REF. 1), human centro­meres
become visible as chromosomal constrictions. The specialized nature and environment of centromeric chromatin enables the assembly of the kinetochore, which
is a large, multi-protein complex that attaches to microtubules during cell division (for reviews, see REFS 2,3),
thereby ensuring equal partitioning of genetic material
between daughter cells. Following each cell division,
chromatin decondenses, the structure and biochemical
composition of centromeres change, and kinetochores
disassemble. During mitosis, this decompaction is visualized by weak DNA staining on individual chromosomes (FIG. 1). In interphase, specialized densely stained
chromocentres become visible in mouse cells. They correspond to pericentric ­heterochromatin (PHC) bringing
several chromosomes together.
The smallest characterized centromere to date is
the ‘point centromere’ in Saccharomyces cerevisiae,
which captures one spindle microtubule4 and measures
125 bp of DNA of a unique sequence dictating centro­
mere location (BOX 1). At the other extreme, species such
as Caenorhabditis elegans have evolved ‘holocentro­
meres’, in which microtubule attachment sites extend
along large portions of the chromosome5. These holo­
centromeres are considered to be point centromeres
that are dispersed across the entire chromosome 6.
This shows the high plasticity of the organization of
centro­meres across different organisms (BOX 1) and the
existence of various means to segregate chromosomes.
‘Regional centromeres’, which are found in most other
eukaryotes, can span up to several mega base pairs (Mb)
and attach to several microtubules7. In most eukaryotes,
they are found at repetitive DNA sequences, known as
α-satellite sequences in humans8. These repeats vary in
length and sequence between species9. In species with
regional centromeres, a single centromere is normally
active on each chromosome: if two centromeres occur
on a chromosome, this leads to aberrant segregation.
The mammalian X and Y chromosomes harbour
repetitive sequences that differ from those of the other
chromosomes10, and under rare circumstances these
repetitive sequences can form centromeres at ectopic
sites. The existence of neocentromeres at sequences
that lack the typical centromeric repeats11 led to the
hypothesis that epigenetic features could determine
centromere location.
Most eukaryotic centromeres are marked by the
histone H3 variant centromere protein A (CENP‑A)12.
Earnshaw and Rothfield identified CENP‑A in 1985
as one of the proteins detected by autoantibodies from
patients with CREST (calcinosis, Reynaud syndrome,
oesophageal dysmotility, sclerodactyly, telangiectasia),
together with the centromeric proteins CENP‑B and
CENP‑C13. In 1991, Palmer et al. described CENP‑A
as a histone H3 variant 14. Although there are continued debates about the most appropriate nomen­
clature for mammalian CENP‑A12,15–17, in this article
we use our favoured term CenH3CENP‑A, which acknow­
ledges the original name of CENP‑A while facilitating
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cross-­comparisons with homologous centromere-­specific
histone H3 variants (CenH3) in non-mammalian s­ pecies
by using different superscript protein names.
Centromeric chromatin undergoes a series of
changes during the cell cycle. In S phase at the time
of DNA replication, following passage of the replication
a Mitosis
Equal distribution of DNA to both daughter cells
DAPI
Prophase
Metaphase
Anaphase
Prophase
Chromatin
condensation
Telomere
Minor satellite (centromere)
Major satellite (PHC)
Long chromosome arm
Metaphase
M
b Interphase
G2–mitosis
Kinetochore
assembly on
centromeric
chromatin
DAPI
fork, CenH3CENP‑A is diluted. A chain of events ensues,
including progressive chromosome condensation
(FIG. 1), and one kinetochore assembles on each centro­
mere. Centromeric CenH3CENP‑A incorporation is tightly
controlled and occurs between telophase and early
G1 in humans and several other eukaryotes18. Other
centro­mere and kinetochore components also exhibit
a particular turnover during the cell cycle at distinct
times. Thus, challenges during the cell cycle, such as
replication stress and DNA damage, can impinge on
these events. Recent technical advances have facilitated
analyses of dynamics in centromere organization during
the cell cycle. In particular, the specialized, engineered
protein tags SNAP-tag and CLIP-tag enable the specific
labelling of newly synthesized protein. This approach
can be used to visualize the cell cycle timing of new
CenH3CENP‑A deposition18,19 and other centromere components20–22. In addition, distinct and complementary
engineered systems have proved to be powerful for
studying de novo centromere formation: human artificial chromosomes (HACs)23,24 could provide a means
to modify the sequence and chromatin environment
at will, and the LacO–LacI targeting system25–28 allows
study of the dynamics of LacI-fused proteins targeted
to a LacO sequence inserted at a specific location.
Although repeat sequences are notoriously difficult to
study in depth using sequencing technologies, modern bioinformatic and sequencing approaches are
now being used to elucidate the biochemical features
of centromeres8,10.
G2
c S phase
BrdU
G1
S
Major satellite Merge
Mid S phase
Minor satellite
Late S phase
Mid S phase
Replication of PHC
Late S phase
Replication of centromeric chromatin
Figure 1 | Centromeric chromatin dynamics during the
cell cycle. Chromatin undergoes major changes during
the cell cycle, which can be visualized by microscopy.
Here, chromatin of mouse 3T3 cells is shown83. a | During
mitosis, chromatin undergoes compaction with distinct
staining patterns in prophase (distinct regions
corresponding to chromosomes appear under
4ʹ,6‑diamidino‑2‑phenylindole (DAPI) staining),
metaphase (chromosome pairs are clearly visible) and
anaphase (chromosome separation to the new daughter
cells with single chromosome arms is visible). White boxes
highlight a single chromosome in metaphase and anaphase,
respectively. Below the metaphase DAPI image is an
immunofluorescence image of metaphase chromosomes:
the long chromosome arm in blue was stained with DAPI,
major satellite DNA was labelled with a fluorescence in situ
hybridization (FISH) probe and coloured in green
(corresponding to pericentric heterochromatin (PHC)) and
the minor satellite DNA was labelled with another FISH
probe coloured in red (corresponding to centromeres).
This organization is shown diagrammatically on the right.
b | During interphase, DNA (stained by DAPI) shows
DAPI-dense regions corresponding to chromocentres, which
constitute heterochromatin. c | During S phase, DNA is
replicated. PHC is replicated in mid S phase and centromeric
chromatin in late S phase. Bromodeoxyuridine (BrdU) shows
distinct patterns in mid and late S phase, and replication
timing was determined by colocalization of BrdU with
fluorescent major or minor satellite probes in mouse 3T3
cells83 (scale bars represent 5 μm). Adapted with permission
from REF. 83, © Guenatri, M. et al., 2004. Originally published
in J. Cell Biol. http://dx.doi.org/10.1083/jcb.200403109.
Nature Reviews | Genetics
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Homotypic
A nucleosome that has copies
of the same variant of H3, H2A
or H2B.
Here, we focus on mammals, and more specifically
humans, to describe the changes in centromeric organization, architecture and protein content during the cell
cycle, which are orchestrated by a network of molecular
players, including histone variants, histone chaperones,
chromatin-remodelling factors and chromatin-­modifying
enzymes12 (TABLE 1). First, starting from centromere organization as the basis for kinetochore assembly, we highlight
the central role of CenH3CENP‑A in centromere identity.
We also introduce a series of key factors associated with
centro­meric chromatin as building blocks or dynamic
players. For each of them we link their expression profiles and availability with structural changes at different
cell cycle stages. Then, we provide the current knowledge
about the cell cycle-coupled mechanisms that ensure
centromere inheritance. This leads us to discuss circumstances in which these processes can go awry, which can
lead to neocentromere formation and potentially disease.
Availability of centromeric chromatin factors
H3 histone variants. During interphase, centromeric
chromatin undergoes major changes coupled to the
expression — that is, the availability — of centromeric
components. The interspersed organization of human
centromeric chromatin comprises homotypic nucleo­
somes containing the histone H3 variants CenH3CENP‑A,
H3.1 and H3.3 (REF. 29) (FIG. 2), and it is important to
consider when these components become available during the cell cycle30. Expression of replicative histones
such as H3.1 peaks during S phase, thus providing the
major histone supply at that time. Their deposition is
coupled to DNA synthesis. Expressed independently
of S phase, replacement H3 variants get incorporated
independently of DNA synthesis. The replacement
variant H3.3 is expressed in all cell cycle phases and in
quiescent cells. Expression of the replacement variant
CenH3CENP‑A peaks during G2 phase in human cells,
Box 1 | Architecture of centromeres in different model organisms
The figure shows centromeres in Saccharomyces
cerevisiae, Schizosaccharomyces pombe, Mus musculus
and Homo sapiens. The point centromere of
S. cerevisiae has only one CenH3Cse4 nucleosome and
three conserved DNA elements (CDEs) (see the figure,
part a). Cohesins map to the surroundings of the
centromere in S. cerevisiae, forming a region
functionally reminiscent of a pericentric region172.
The regional centromere of S. pombe has a core DNA
sequence, flanked by innermost repeat (imr) sequences
and several CenH3Cnp1 nucleosomes and H3
nucleosomes (see the figure, part b). The centromere is
flanked by heterochromatin consisting of the outer
repeat (otr) sequences. The mouse centromere has a
centric region flanked by pericentric heterochromatin
(PHC) at one side (see the figure, part c). The centric
region is transcribed into minor satellite RNA and PHC
is transcribed into major satellite RNA. Both RNAs have
structural roles in their respective regions. The major
satellite RNA is required to recruit heterochromatin
protein 1 (HP1). Importantly, mouse heterochromatin
shows the highest transcriptional activity at the G1–S
transition and in early S phase, and is then silenced
during G2 phase173, which is regulated by distinct
transcription factors at PHC. PHC is characterized by
HP1 and SUV39H1, which is a histone-lysine
N‑methyltransferase that is responsible for
establishing histone H3 lysine 9 trimethylation
(H3K9me3) at PHC. The human centromere has long
stretches of α-satellite repeats and several CenH3CENP‑A,
H3.1 and H3.3 nucleosomes (see the figure, part d).
Centromeres are flanked by PHC. The centric
α‑satellite region is transcribed and the transcripts
might have a structural or functional role. PHC might
also be transcribed in a similar way to that observed in
mice. HP1 and SUV39H1 characterize PHC.
Throughout the figure, to distinguish the inner part
of the centromere from its flanking regions, we
designate them as centric regions and PHC,
respectively. Notably, the term ‘centromeric’, although
often used as a synonym for the centric region, usually
covers a broader centromere area encompassing both
the centric and the pericentric regions.
a S. cerevisiae
Regional centromere
CenH3Cse4 nucleosome
H3 nucleosome
DNA
CDE1
CDE2
CDE3
125 bp
b S. pombe
Regional centromere
CenH3Cnp1 nucleosome
H3 nucleosome
DNA
otr
imr
Heterochromatin
imr
Core
otr
Heterochromatin
Centric region up to 3.5–11.0 × 105 bp
c M. musculus
Regional centromere
CenH3CENP-A nucleosome
H3 nucleosome (H3.2 and H3.3)
DNA
PHC
Major satellite RNA
HP1
SUV39H1
Major satellite
repeat
Minor
satellite
repeat
Chromatin
Centric region up to 1.0 × 106 bp
Minor satellite RNA
d H. sapiens
Regional centromere
H3.3 nucleosome
CenH3CENP-A
nucleosome
H3.1 nucleosome
DNA
PHC
RNA?
Centric region up to 5.0 × 106 bp
HP1
α-satellite
α-satellite RNA
SUV39H1
repeat
PHC
RNA?
HP1
SUV39H1
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Table 1 | Centromeric and kinetochore factors
Function
H3 variants
Factor
H3 (canonical)
H3.3
(metazoan)
CenH3
Licensing
CENP‑B
CENP‑C
RBAP46,
RBAP48
MIS18α,
MIS18β
MIS18BP1
CDK1, CDK2
Deposition
Post-­
translational
and DNA
modifications
Kinetochore
components
or regulators
Condensin I,
condensin II
HJURP (Scm3,
Cal1)*
Conservation
•Hs, Mm: H3.1, 3.2
•Dm, Xl: H3.2
Sc, Sp: H3
Hs, Mm, Dm, Xl: H3.3
•Hs, Mm, Xl: CENP‑A
•Dm: CID
•Sc: Cse4
•Sp: Cnp1
•At: HTR12
•Ce: HCP‑3
•Mm: CENP-B
•Sp: Abp1, Cbh1, Cbh2
•Sc: Cbf1
•Mm: CENP‑C
•Sc: Mif2
•Sp: Cnp3
Sp: Mis16
Sp: Mis18
Ce: KNL‑2 (Hs: MIS18BP1 is also known
as KNL2)
•Mm: CDK1, CDK2
•Sc: Cdc28
•Sp: Cdc2
Xl: condensin I, condensin II
•Hs, Mm, Xl: HJURP
•Sc, Sp: Scm3
•Dm: Cal1
NPM1
•Mm: NPM1
FACT complex
(SPT16, SSRP1)
•Sc: Spt16
•Sp: Pob3, Spt16
•Dm: Dre4
•Mm: CUL4B
CUL4–RING E3
ubiquitin ligase,
RBAP46
K124
Unknown
ubiquitylation
Hs CENP‑A S68 Unknown
Properties
Deposition in a DNA synthesis-dependent manner (replicative
histone H3 variant); the specific chaperone is CAF1
H3 can be deposited in DNA synthesis-dependent or ‑independent
manners
Deposition in a DNA synthesis-independent manner; the
responsible chaperones are HIRA and DAXX–ATRX
Centromeric H3 variant that is highly enriched at centromeres and
is the epigenetic determinant of centromeres
Stabilizes centromeres and kinetochores by forming another
interface with centromeric DNA, CenH3CENP‑A nucleosomes and
CENP‑C; not an essential gene in mice
Involved in a complex feedback mechanism with HJURP and the
MIS18 complex
General chaperones for H3–H4 and possibly CenH3; potentially
involved in altering chromatin acetylation status
Part of the Hs MIS18 complex, do not associate with CenH3CENP‑A
in vivo
Part of the Hs MIS18 complex, does not associate with CenH3CENP‑A
in vivo
Regulate centromeric cell cycle progression and phosphorylate
MIS18BP1; involved in the HJURP phosphorylation cycle controlling
HJURP localization to centromeres
Impose a chromatin structure on centromeres that is necessary for
successful incorporation of CenH3
CenH3CENP‑A-specific chaperone needed for its deposition at
centromeres; interacts directly with CENP‑C, the MIS18 complex
and DNA; involved in CENP‑C reorganization at centromeres and
has a role in establishing the correct organization of centromeric
and kinetochore factors
Identified as part of the HJURP complex, but its function in
centromeric establishment and maintenance is unclear
General chaperone for H3–H4, H2A–H2B and potentially other
centromeric factors
Refs
41
41
43,46
13
55,56
26,129
47,123
3,123,
128,135
123,130
19,133
144
19,26,
47,48,
79
47
118
Complex needed to control CenH3CENP‑A deposition at centromeres
142
Ubiquitylation of K124 is reported to be required for efficient Hs
CENP‑A deposition at centromeres
Ser68 phosphorylation controls the interaction between HJURP
and CenH3CENP‑A that determines CenH3CENP‑A deposition timing
Phosphorylation of these residues controls the timing of localization
of HJURP to centromeres; CDK1 is involved in this control
Methyltransferase for H3K9me3, which prevents CenH3CENP‑A
deposition if its function is upregulated at centromeres
142
Hs HJURP S412, Unknown
S448, S473
SUV39H1
•Mm: SUV39H1
•Dm: Su(var)3-9
•Sp: Suv39h1
Constitutive
CENP‑H (Sp: Sim4), CENP‑I (Sp: Mis6),
Multiprotein complex that functions as a structural core of the
centromere
CENP‑K,CENP‑L (Sp: SPAC4F10.12
kinetochore, either forming the inner kinetochore or recruiting
associated
(Fta1)), CENP‑M (Sp: Mis17), CENP‑N
kinetochore components
network
(Sp: Mis15), CENP‑O, CENP‑P, CENP‑Q,
(CCAN)
CENP‑R, CENP‑S, CENP‑T, CENP‑U
(CENP‑50, KLIP1), CENP‑X, CENP‑W
140
19
24
51,63,
71
A list of the factors of the human CenH3CENP‑A maintenance and deposition network, showing conservation across multiple species and the function of these factors in the
network. At, Arabidopsis thaliana; ATRX, α‑thalassaemia/mental retardation syndrome X‑linked; DAXX, death domain-associated protein 6; Dm, Drosophila melanogaster;
CAF1, chromatin assembly factor 1; Cal1, chromosome alignment defect 1; CDK, cyclin-dependent kinase; Ce, Caenorhabditis elegans; CENP, centromere protein;
CUL4, cullin 4; FACT, facilitates chromatin transcription; HJURP, Holliday junction recognition protein; Hs, Homo sapiens; Mm, Mus musculus; NPM1, nucleophosmin;
Sc, Saccharomyces cerevisiae; Scm3, suppressor of chromosome mis-segregation protein 3; Sp, Schizosaccharomyces pombe; Xl, Xenopus laevis. *Homologues of HJURP
exist in most vertebrates, including the Scm3 proteins in S. cerevisiae and S. pombe. Despite their conserved function in CenH3 deposition, HJURP/Scm3 homologues
exhibit high degrees of sequence divergence among species. However, HJURP and Scm3 have a conserved CenH3‑interacting amino‑terminal domain138, and the
structure of the S. cerevisiae CenH3Cse4–H4–Scm3 complex shows striking similarities to the human structure. Homologues of HJURP have not been identified in
D. melanogaster, C. elegans or plants. In D. melanogaster, the absence of HJURP is compensated for by the CenH3CID chaperone Cal1 (REF. 79). Cal1 localizes to
centromeres during the same time window in the cell cycle as HJURP, but the exact time of CenH3CENP‑A and CenH3CID deposition differs.
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Homo sapiens centromere
DNA
PHC
PHC
Centromere
Kinetochore assembly (G2–M)
MIS18BP1
CENP-T
α
CENP-C
β
HJURP
Protein expression
during the cell cycle
CENP-W
CENP-B
CENP-N
CENP-B
box (DNA
sequence)
H3.3
nucleosome
CenH3CENP-A
nucleosome
G2 phase
H3.1
nucleosome
S phase
All phases
CenH3CENP-A
H3.1
H3.3
Constitutive
Molecular interactions at centromeres
CENP-B
N terminus of CenH3CENP-A DNA (CENP-B box)
G2–M
CENP-C
C terminus of CenH3CENP-A (nucleosomal); HJURP;
CENP-H–CENP-I–CENP-K–CENP-M complex
G2, mitosis, G1 (highest)
CENP-N
Arg–Gly (RG) loop of CenH3CENP-A
All phases with a
peak in S phase
CENP-T
Transient
CENP-W
?
MIS18BP1 β
α
HJURP
CENP-H–CENP-I–CENP-K–CENP-M complex;
CENP-T–CENP-W part of a nucleosomal particle?
Late S–G2
Peak in G2
CENP-C; HJURP
CenH3CENP-A (pre-nucleosomal); HJURP; CENP-C
Peak in G2
Figure 2 | Molecular features of centromeric chromatin during the cell cycle. Human centromeres are flanked by
pericentric heterochromatin (PHC) (top). Below are the key components of centromeric chromatin
described
Nature
Reviewsin| this
Genetics
Review. Centromeric chromatin has CenH3CENP‑A nucleosomes, interspersed with H3.1 and H3.3 nucleosomes. Centromere
proteins (CENPs) are characteristic of centromeric chromatin, and the CENPs that are central to this Review are depicted:
CENP‑B, CENP‑C, CENP‑N and CENP‑T–CENP‑W, as well as Holliday junction recognition protein (HJURP) and the MIS18
complex. Key molecular interactions of these factors at centromeres are listed, as well as their expression patterns during
the cell cycle. The CENP‑T–CENP‑W complex is a key kinetochore component and might form nucleosome-like particles
together with CENP‑S and CENP‑X71 (a nucleosomal context is indicated by the grey comma shapes). Factors that are
found transiently at centromeres during the cell cycle are indicated. CENP‑T–CENP‑W and CENP‑N levels vary during the
cell cycle but are still constitutively present at centromeres. For full reviews of the constitutive centromere-associated
network (CCAN) and kinetochores, see REFS 174,175.
making it available for deposition later in mitosis (for
reviews on histone H3 dynamics and expression, see
REFS 12,31,32).
CenH3CENP‑A is deposited de novo at centromeres
between telophase and early G1 phase in humans18
and other mammals33. Orthologues of CenH3CENP‑A are
incorporated into the centromeric chromatin of diverse
eukaryotes, and all CenH3 orthologues can mark
centro­meres and form nucleosomes. Human cells can
incorporate S. cerevisiae CenH3Cse4 into centromeres,
which indicates that CenH3Cse4 can form functionally and structurally equivalent nucleosomal particles
despite sequence divergence34. Alternatively, CenH3Cse4
might aid functionality in combination with any residual
CenH3CENP‑A still present. Intriguingly, CenH3 deposition
timing differs between mammals and yeast12, but in all of
these organisms, CenH3 is essential35,36. In S. cerevisiae,
CenH3Cse4 deposition occurs during S phase37, whereas
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Telomere
CentroPericentric
mere heterochromatin
Gapfilling
Active genes Regulatory Telomere
and promoters elements
Replication
genome wide
Chaperone with
key role in deposition
ATRX–DAXX
CAF1 complex
HIRA complex
ATRX–DAXX
H3.3
H3.1
H3.3
H3.3
All phases
S phase
All phases
All phases
Histone H3 variant
Deposition during
cell cycle
Centromere
Chaperone
with key role
in deposition
HJURP
MCM2
CAF1
complex
HIRA
complex
FACT
CenH3CENP-A
Histone H3
variant
CenH3CENP-A
H3.3
CENP
protein
H3.1
H3.3
S phase
All phases
CENP-T
CENP-W
HJURP ?
CENP-B
CENP-C
CENP-N
H3.1
Deposition
during cell
cycle
Telophase–
early G1
S phase
Late S–G2
All phases? All phases?
G1–S
Figure 3 | Cell cycle timing of deposition and chaperones of histone H3 variants and key centromere
proteins.
Nature Reviews
| Genetics
Depiction of a chromosome with key regions indicated where histone H3 variants are deposited. The chaperones and
times of incorporation involved are shown. Centromeres consist of nucleosomes containing the centromeric histone H3
variant CenH3CENP‑A, interspersed with nucleosomes containing H3.1 and H3.3. These histone H3 variants are deposited
differentially in a cell cycle-dependent manner with the involvement of different histone chaperone complexes. Key
centromere proteins (CENPs) that form part of centromeric chromatin are also deposited in a cell cycle-dependent
manner. ATRX, α‑thalassaemia/mental retardation syndrome X‑linked; CAF1, chromatin assembly factor 1; DAXX, death
domain-associated protein 6; FACT, facilitates chromatin transcription; HJURP, Holliday junction recognition protein.
Figure adapted from REF. 32, Macmillan Publishers Ltd.
in Schizosaccharomyces pombe, CenH3Cnp1 deposition
starts in S phase but the bulk of deposition occurs in G2
phase38,39. In humans, mice, Drosophila melanogaster and
Xenopus laevis, centromeric CenH3 deposition occurs
outside S phase18. Although it is not entirely clear why
the system has evolved in that manner, the uncoupling
of CenH3CENP‑A deposition from S phase may give the
cell the opportunity to avoid adverse interference during replication and regulate the spatial deposition of
CenH3CENP‑A along with other components, which may
help to prevent ectopic localization.
Histone chaperone complexes. In addition to availability, one has to consider how each component can be
delivered and incorporated. Histone chaperone complexes that escort histone variants in the cell31,40 are
important for this. They help to transfer histones without necessarily being a component of the final chromatin product. Whereas the chromatin assembly factor 1
(CAF1) complex is dedicated to H3.1 (REF. 41) and H3.2
(REF. 42),
the HIRA complex 41 and the death domain-­
associated protein 6 (DAXX)–α‑thalassaemia/mental
retardation syndrome X‑linked (ATRX) complex 43 deals
with H3.3. HIRA promotes H3.3 deposition at active
genes, possibly at transient nucleosome-free regions44
and DNA damage sites45, and the DAXX–ATRX complex is important for H3.3 enrichment at PHC domains
that are located adjacent to the centric region. PHC,
which is considered to be a typical constitutive heterochromatic region, is decorated with distinct histone
marks that are also found close to telo­meres46, which
are specialized chromatin regions at the end of the
chromo­some arms (FIG. 3). In humans, Holliday junction
recognition protein (HJURP) is the dedicated chaperone for CenH3CENP‑A (REFS 47,48), which is enriched in
the centric region. HJURP interacts with the CENP‑A
targeting domain (CATD) of CenH3 CENP‑A (REF. 49).
Deposition of CenH3CENP‑A and the dynamics of nucleo­
somal particles are crucial to the organization of these
domains. During the cell cycle, just after DNA synthesis,
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it is important to understand how nucleosomal features
of the region are duplicated, and how they integrate
with higher-order architectures in interphase, and
ultimately mitosis, to faithfully segregate the d
­ aughter
chromosomes.
Centromere-associated and kinetochore proteins.
Between G2 phase and mitosis, the kinetochore assembles on centromeric chromatin. The kinetochore is a
complex network of proteins that are grouped into the
inner kinetochore, which comprises the constitutive
centromere-associated network (CCAN), and the outer
kinetochore. The CCAN components were initially
identified using CenH3CENP‑A‑based pull-down studies
to identify other centromere-associated proteins50–52.
The biochemical and structural nature of centromeric
chromatin, nucleated by CenH3CENP‑A nucleosomes,
is key for controlling kinetochore assembly. Whereas
H3.1 and H3.3 nucleosomal structures are almost identical, the more compact CenH3CENP‑A nucleosome only
wraps 121 bp of DNA53, which has an impact on higher-­
order chromatin structure (FIG. 4). These CenH3CENP‑A
nucleosomes help to connect the centromere to the
kinetochore through important chromatin-­associated
partners. CENP‑B interacts specifically with the centric α-satellite DNA54 and with CenH3CENP‑A nucleo­
somes through the CenH3CENP‑A amino‑terminal tail55.
Although a CenH3CENP‑A knockout is embryonically
lethal in mice35, CENP‑B‑deficient mice are viable56–58,
which suggests that CENP‑B might have a stabilizing
but not essential role at centromeres55. CENP‑C interacts with CenH3CENP‑A‑containing particles through the
carboxy‑terminal tail of CenH3CENP‑A and docks onto
the acidic patch of histone H2A and H2B59. CENP‑C
binding compacts the structure of the CenH3CENP‑A
nucleosome60 and serves as a binding platform for other
kinetochore components.
Most kinetochore components are recruited to
the centromere from late S phase onwards; recruitment continues through G2 phase until early mitosis.
It is unknown at what cell cycle stage new CENP‑C is
recruited, but it shows increased localization dynamics in S phase 61. Interestingly, the flexible ends of
CenH3CENP‑A nucleosomes prevent binding of histone H1
(REF. 62), which is a crucial structural component binding to H3.1 and H3.3 nucleosomes. This contributes to
the formation of a chromatin structure that is distinct
to other regions.
In late S phase, CENP‑T is deposited. It anchors
centro­meric chromatin to the kinetochore. The CENP‑T–
CENP‑W complex accumulates at centromeric H3.3
nucleosomes63,64 and not at CenH3CENP‑A nucleosomes
like CENP‑C. However, CenH3CENP‑A immunoprecipitates CENP‑T in humans51, and both CenH3CENP‑A and
CENP‑C are required for CENP‑T centro­meric localization26,63. An initial model proposed that CENP‑C and
CENP‑T constitute separate axes of kinetochore formation connecting to the outer kinetochore63,65. Indeed, in
chicken cells, CENP‑T can bind to the outer kinetochore
in the absence of CENP‑C66. Interestingly, both CENP‑C
and CENP‑T seem to recruit the outer kinetochore65.
However, recent evidence in human cells points towards
a dependence of the CENP‑T–CENP‑W complex on
CENP‑C, with the two being bridged by a complex
of CENP‑H, CENP‑I, CENP‑K and CENP‑M26,67; these
four proteins were also recently identified as being part
of a larger complex that also contains CenH3CENP‑A,
CENP‑C, CENP‑L and CENP‑N68. This co‑occurrence
in a complex argues for an interconnectivity between
CENP‑C and CENP‑T, which has been substantiated
by a study describing CENP‑C and CENP‑T as forming
a complex together with CENP‑B69. It is possible that a
strict hierarchical model does not reflect the nature of
the kinetochore, as suggested for the CCAN70. Taken
together, the extent to which CENP‑C and CENP‑T have
separate versus overlapping roles in kinetochore assembly is still an open debate in the field and will have to be
dissected further in future work. Another model posits
that the CENP‑T–CENP‑W complex, given its similarity
with H2A–H2B, could form a nucleosome-like particle
together with CENP‑S–CENP‑X71.
In addition to these post-replicative deposition
events before mitosis, there are exceptions. For instance,
CENP‑N is recruited to centromeres de novo at the G1–S
transition72. This is mediated by CENP‑N interaction
with CenH3CENP‑A through a loop region protruding
from the nucleosome.
Taken together, we have learned a lot about the
molecular interactions between individual partners
at centromeres, which has enabled us to reconstruct a
detailed structural view of the basis of centromeric chromatin and the kinetochore. This composition and structure changes throughout the cell cycle, which will have to
be considered in detail when studying the centromeric
locus in future studies.
Non-coding RNA and centromeric transcription.
Human centromeres have been suggested to produce
a non-coding RNA73 with an unknown function. This
RNA could be involved in higher-order chromatin structures, and given the nucleic-acid-binding domains of
CENP‑B55, HJURP19 and other centromeric components,
this possibility should be investigated in more detail.
Indeed, this RNA could be part of a mechanism that
influences centromeric architecture during the cell cycle.
Beyond searching for specific functional roles of the
RNA, it is important to consider whether the mere act
of its transcription might have functional implications;
indeed, a low level of transcription at the centromere
between telophase and early G1, has been suggested
to be required for CenH3 deposition in various model
systems. Centromeric transcription was first reported
in S. pombe74 and occurs in various species, including
humans75,76. In addition, transcription of PHC is important for the establishment and maintenance of PHC77.
This continuous transcription challenges the structure
and biochemical composition of centromeres throughout the cell cycle. In S. pombe, the sequence-­encoded
features of centromeric DNA can create an environment
of pervasive low-fidelity RNA polymerase II (Pol II)
transcription that is important for CenH3Cnp1 assembly 78. Treatment of human cells with Pol II inhibitors
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compromises centromere integrity 75. In D. melanogaster,
Pol II‑dependent transcription is required for CenH3CID
deposition by its chaperone chromosome alignment
defect 1 (Cal1)79 during mitosis: the requirement for a low
level of transcription may permit the formation of a specialized chromatin structure in mitosis that is required
for CenH3Cnp1 loading. Studies using HACs showed that
targeting transcriptional activators alters local chromatin structure and gene expression80. Furthermore, HAC
centro­meres can resist transcriptional silencing mediated
by histone H3 lysine 9 trimethy­lation (H3K9me3) and
H3K27me3 when the enhancer of zeste homologue 2
(EZH2) methyltransferase is tethered81, and reduction
of euchromatic histone marks in the centromere causes
a
Histone marks at the centromere
Constitutive
Transient
Modifications on the
CenH3CENP-A nucleosome
Modifications on the H3.1
or H3.3 nucleosome
Modifications on the H3.1
or H3.3 nucleosome
CenH3CENP-AS16P
CenH3CENP-AS18P
H3K4me2
H3K36me2
H3K9me2
H3K9me3
H4K20me3
CenH3CENP-AS7P (prophase)
H4K20me1 (early G1)
CenH3CENP-AK124ub (G1)
CenH3CENP-AK124ub (G1–S)
H3T11P (M phase)
H3S10P (prophase)
H3S28P (prophase)
H3.1
CenH3CENP-A
Histone variant
dynamics
Deposition
(telophase–early G1)
Deposition
(S phase)
Dilution
(S phase)
b
Histone marks at PHC
H3.3
Histone marks along the
chromosome arms with a role
in mitosis
Modifications on the H3.1
or H3.3 nucleosome
H3S10P (spreading in M phase)
H3S28P (spreading in M phase)
H3T3P (M phase)
H3T11P (M phase)
H3.3S31P (anaphase)
H2A.Z
Deposition
(early G1)
Deposition
(all phases)
Detected loss
(mitosis)
Histone tail
H3T3P
H3K4me2
H3K9me2/3
H3S10P
H3T11P
H3K27me3
H3S28P
H3K36me2
H3T3P
H3K4me2 H3K9me2/3
H3S10P H3T11P
H3K27me3
H3S28P
H3K36me2
H3T3P
CenH3CENP-AS7P
H3K9me2/3 H3K4me2
H3T11P H3S10P
CenH3CENP-AS16P
H3K27me3 H3S28P
H3.3S31P
CenH3CENP-AS18P
H3.3S31P
H3K36me2
CenH3CENP-AS7P
CenH3CENP-AS16P
CenH3CENP-AS18P
CenH3CENP-A
K124ub
CenH3CENP-A
K124ac
H4K20me3
H4K20me3
H4K20me1
DNA
DNA
H4K20me1
DNA
H4K20me3
H3.1 nucleosome core
particle (146 bp of DNA)
H3.3 nucleosome core
particle (146 bp of DNA)
Figure 4 | Cell cycle dynamics of histone modifications of centromeric and
pericentric chromatin. a | Depiction of characterized histone marks specific
for the centric region and pericentric heterochromatin (PHC) at human
centromeres. Constitutive and transient marks are indicated. Modifications
of CenH3CENP‑A nucleosomes are highlighted in blue, modifications of H3.3
nucleosomes are highlighted in green and modifications that are common to
both H3.1 and H3.3 nucleosomes have no highlighting. b | Crystal structures of
the H3.1, H3.3 and CenH3CENP‑A nucleosome core particles. The canonical
CenH3CENP-A nucleosome core
particle (121 bp of DNA)
nucleosome core particle consists of a tetramer containing two H3–H4 dimers
((H3–H4)2) flanked by two H2A–H2B dimers, all wrapped
by 146 b|pGenetics
of DNA.
Nature Reviews
Whereas H3.1 and H3.3 have minimal amino acid differences, CenH3CENP‑A
only shows ≈45% sequence conservation, in line with a rapid evolution of
centromere organization and its components174. The more compact
CenH3CENP‑A nucleosome is wrapped by only 121 bp of DNA53. Histone tail
modifications at centromeres or PHC are indicated. Red boxes indicate
phosphorylation. ac, acetylation; me, methylation; ub, ubiquitylation.
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a drop in CenH3CENP‑A levels in HACs82. It will be important to investigate centromeric transcription with respect
to the cell cycle and the impact on the biochemical composition and ­structure of centromeric chromatin at each
cell cycle stage.
The availability and dynamics of each of the described
centromere components, along with their interconnections, provide key parameters contributing to changes in
the overall architecture. Notably, centromeres of multi­
cellular eukaryotes are typically embedded in hetero­
chromatin, with distinct structural and biochemical
features. How this flanking PHC crosstalks with the
organization of the centric region plays a crucial part in
centromere dynamics.
Crosstalk between centric chromatin and PHC
PHC is a condensed region marked by histone H3K9
methylation and is enriched in proteins of the hetero­
chromatin protein 1 (HP1) family, which bind
H3K9me3. In mouse cells, the replication timing of
PHC precedes that of centromeres during S phase83
(FIG. 1). PHC is not just structurally but also functionally linked to the centric region. HP1 is implicated in
hetero­chromatin formation and maintenance, and
transcriptional regulation84. It is also involved in regu­
lating cohesin binding to centromeres, which is crucial for chromosome segregation during mitosis85.
Mammalian PHC is marked by the repressive histone
marks H3K9me2, H3K9me3 and H4K20me3, whereas
the centric region mainly carries the euchromatic marks
H3K4me2 and H3K36me (FIG. 4). H3K4me2 has a role
in HJURP recruitment during telophase and early G1
phase, and its levels are regulated by lysine-specific histone demethylase 1 (LSD1)86. At human and chicken
centromeres, H4K20me1 on CenH3CENP‑A nucleosomes
is a transient mark that is essential for the stability of
the kinetochore following CenH3CENP‑A deposition87. In
mouse cells the major satellite RNA is a structural component that is important for the accumulation of HP1 at
PHC88. HP1α is sumoylated, which facilitates its de novo
recruitment to PHC89.
Changes in the phosphorylation states of many factors, including several centromeric and peri­centric
components, are important events in the lead up to and
during cell division, and the phosphorylation of various
residues on CenH3CENP‑A, H3.1 and H3.3 at centromeres
and PHC is linked to key functions during mitosis. The
N‑terminal tail of CenH3CENP‑A is phosphorylated at
S16 and S18, which results in greater intranucleosome
associ­ations90. It is also phosphorylated transiently at S7
in mitosis, which facilitates interaction with CENP‑C91.
H3.1 and H3.3 nucleosomes are phosphorylated transiently in mitosis at the T3, S10, T11 and S28 positions,
which is crucial for proper chromosome segregation92.
A conserved feedback mechanism monitors mis-­
segregating chromosomes during anaphase through
the differential phosphorylation of histone H3.3 at S31
(REF. 93) (FIG. 4); this modification is predominantly found
in PHC during mitosis94. Thus, it is tempting to speculate that mutations in H3.3S31 in PHC may consequently
lead to defects in chromosome segregation.
S. pombe95 has been a useful model system for studying connections between chromosome segregation and
PHC96. In S. pombe, heterochromatin flanking the central
kinetochore domain is directed by RNA interference and
is required to promote CenH3CENP‑A deposition97. PHC
may inhibit the spreading of the centro­mere over neighbouring gene regions. This supports the idea of barriers
between PHC and the centromere core, as has been shown
for regional centromeres in S. pombe, which are regulated
by tRNAs and Pol III98. Point centromeres of S. cerevisiae
and holocentromeres in C. elegans are not flanked by
hetero­chromatin, but a boundary function might be carried out by centromere-­binding factor 1 (Cbf1) and Cbf3
in S. cerevisiae99, and by well-positioned canonical nucleo­
somes in C. elegans. Heterochromatin thus represents a
unique feature that is associated with regional centro­
meres. This is in line with the fact that in chicken and
human cells, the kinetochore covers the centric region,
but not PHC100, again suggesting a boundary function.
Disrupting PHC integrity by inhibiting histone methy­
l­ation was shown to increase centromeric CenH3CENP‑A
levels without changing its deposition timing in mouse
cells, highlighting that the integrity of both domains is
crucial for centromere functioning 33. The deposition
of core histone H2A at PHC occurs in a replication-­
dependent manner during mid to late S phase, whereas
the variant H2A.Z is incorporated during G1 phase33.
This is a potential signal to regulate the stoichiometry
of histone variants between the centric and pericentric
regions. Thus, chromatin surrounding centromeres can
have an impact on centric and pericentric composition
and ­structure during the cell cycle.
Centromeric dynamics during S phase
During DNA replication, chromatin undergoes disruption
and subsequent restoration following the passage of the
replication fork (FIG. 5). For reviews on cell cycle-­related
changes of PHC, see REFS 4,101. Notably, centromeric
CenH3CENP‑A is diluted during S phase and distributed
evenly to both daughter chromosomes in mammals18.
Three hypotheses could explain how cells duplicate the
organization to restore the pre-replication chromatin
state: nucleosome-free gaps left during S phase; nucleosomal particles with only one CenH3CENP‑A molecule; and
placeholder nucleosomes replacing gaps left by diluted
CenH3CENP‑A nucleosomes. The latter two hypotheses have
gathered experimental support, although the hypothesis
of nucleosome-free gaps cannot yet be formally excluded.
Previous studies provide arguments for the existence of
‘hemisomes’ containing one CenH3CENP‑A molecule102,103,
with different CenH3CENP‑A‑containing nucleosomal structures cycling from octamer to hemisome during the cell
cycle104, but the structural evidence is under debate105,106.
Whereas in vivo chemical cleavage data in S. cerevisiae
point towards the existence of hemisomes in that species, which could be specific for point centromeres107,
high-resolution imaging techniques point towards an
octameric structure108. Given the crystal structures of
CenH3CENP‑A‑containing particles53,60,109, octamers probably constitute most of the CenH3CENP‑A nucleosomes
in humans110.
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Recycled
Histones are evicted during
S phase and transcription, and
new histones can be deposited
de novo or the old histones can
be deposited again, meaning
that they are recycled.
Deposition of H3.1 and H3.3 at centromeres is
observed during S phase in human cells111, and the
detected loss of H3.3 only later in G1 phase, at the time
of CenH3CENP‑A deposition111, fuelled the hypothesis for
H3.3 as a CenH3CENP‑A placeholder. Although new H3.1
is deposited in a manner coupled to DNA replication
in S phase, it would chiefly contribute to maintaining
the interspersed organization of the centric region. For
each gap generated by diluting CenH3CENP‑A, the HIRA
complex could mediate a post-replicative mechanism
based on gap filling. Whereas H3.1 and H3.3 can be
deposited de novo and recycled at centromeres during
S phase, CenH3CENP‑A is exclusively recycled18. MCM2
is a chaperone that is involved in handling the recyc­
ling of old histones at the replication fork 112–114 to
antisilencing function 1 (ASF1), which facilitates depo­
sition of old histones after the replication fork on the
new DNA. Quantification and further analyses of the
dynamics of newly deposited and evicted histones at
centromeric chromatin will be required to understand
how other histone H3 variants can replace CenH3CENP‑A,
and to elucidate the connection between H3.1 and H3.3
depo­sition in S phase and the loss of H3.3 later during
mitosis111.
It is important to consider the supply of histone vari­
ants and their dedicated chaperones during S phase.
Depletion of CAF1 impairs replicative H3.1 incorporation, enabling compensatory H3.3 deposition at replication sites through HIRA44 using a post-replicative
deposition mechanism. In CenH3CENP‑A‑overexpressing
cells 115, the H3.3 chaperone DAXX promiscuously
handles CenH3CENP‑A, leading to its deposition at non-­
centromeric regions. Thus, the cell needs to control a
careful balance of H3 variants and histone chaperones
to ensure a correct biochemical composition at centro­
meres later in G2 phase and in mitosis. The distribution of old and new histone marks on centromeric H3.1
and H3.3 nucleosomes in S phase is an open question.
Although we have learned how chromatin maturation is
globally coupled to replication116,117, how this is operating
at this specific locus remains unknown. An interesting
hypothesis to consider is that histone marks could be
a sensor for the deposition of placeholder nucleosomes
and could indicate H3.3 nucleosomes to be replaced
later in mitosis. The interspersed nature of the centric
region also leaves space for fluctuations to accommodate
various amounts of CenH3CENP‑A. Determining which
other chaperone (or chaperones) could be involved in
H3.1 and H3.3 deposition at centromeres during S phase
and possible eviction during mitosis or G1 phase will
be a future challenge. Because H3.3 is found at regions
of high histone turnover and active genes, the involvement of H3.3 as a placeholder at centromeres leads to
­consideration of the role of transcription.
Furthermore, the cell cycle dynamics of other components are important. CENP‑T–CENP‑W deposition
at centromeres in late S or G2 phase20 might be coupled to transcription21. Notably, CENP‑T–CENP‑W21
harbours histone fold motifs that are reminiscent of
H2A–H2B. One histone chaperone that has been linked
to transcriptional events and H2A–H2B handling is the
facilitates chromatin transcription (FACT) complex 118,
and it also binds to CENP‑T–CENP‑W21. FACT localizes
to human centromeres handling H2A–H2B exchange
during transcription119. As the soluble H2A–H2B pool
is reduced and that of CENP‑T–CENP‑W increased20
at the end of S phase, FACT may switch from handling
H2A–H2B to CENP‑T–CENP‑W. FACT is also linked to
the assembly of new nucleosomes during replication (for
a review, see REF. 120). Taken together, this highlights
how not only the biochemical nature of interactions but
also the availability of factors at the right cell cycle stage
are critical for maintaining the domain. Recruitment of
CENP‑T–CENP‑W marks the onset of the recruitment
of ­kinetochore factors in G2 phase.
Licensing CenH3CENP‑A deposition at centromeres
Human centromeric chromatin is physically prepared
(licensed) for CenH3CENP‑A deposition121 during G2
phase, which allows subsequent CenH3CENP‑A deposition
between telophase and early G1 phase (FIG. 5). Licensing
involves chromatin remodelling 122 and a change of
the centromeric acetylation status82. In humans, the
histone-­binding proteins RBAP46 (also known as
RBBP7) and RBAP48 (also known as RBBP4) — which
are also subunits of many histone-modifying enzymes,
chromatin-­r emodelling and chromatin-assembly
­complexes — are recruited to centromeres with the
MIS18 complex (consisting of MIS18BP1 (also known
as KNL2), MIS18α and MIS18β)123 at anaphase onset,
and depletion of these components leads to a reduction
of centro­meric CenH3CENP‑A levels. Interestingly, tethering an acetyltransferase to a locus containing α-satellite
sequences and MIS18 stimulates CenH3CENP‑A deposition124. Collectively, these observations suggest that
the change in the centromeric acetylation status either
affects or is concomitant with the timely recruitment
of CenH3CENP‑A, and the detailed mechanisms of this
should be dissected in future work. Deciphering which
histone variants are acetylated will be important for our
understanding of centromeric architecture between G2
phase and mitosis.
In D. melanogaster, the existence of a trimolecular complex containing CenH3 CID, histone H4 and
RBAP46 or RBAP48 suggests a conservation of centro­
mere licensing in Diptera125. These players are implicated in controlling the centromeric acetylation status
in conjunction with acetyltransferases121. Tethering an
acetyltransferase at centromeres bypasses the MIS18
requirement for centromere licensing, suggesting that
it may be the acetylation status per se, and not the presence of these factors, that is required for CenH3CENP‑A
accumulation24. In humans, RBAP46 or RBAP48 is
part of a complex including HJURP47. A homologous
complex is involved in the licensing of centromeres
in S. pombe, where Mis16 (which is similar to human
RBAP46 and RBAP48) and Mis18 form a complex and
maintain the deacetylated state of centromeric histones.
In S. pombe, Mis18 oligo­merization is required for centromeric targeting 126, similar to the heterotetramer formation by MIS18α and MIS18β in humans127, where it
licenses centromeres for HJURP-mediated CenH3CENP‑A
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c Centromere priming (late G2 phase to late mitosis)
d CenH3CENP-A deposition (telophase to early G1 phase)
Decreased CDK1/2 activity, HJURP and MIS18 site-specific
dephosphorylation
P
P
α β
P
MIS18BP1 HJURP
MIS18BP1 HJURP
CDK1/2
CENP-T
CENP-N
PLK1
Recruitment:
MIS18BP1 in late
G2–early mitosis
CENP-N
CENP-W
Site-specific phosphorylation of MIS18BP1; HJURP dimerizes and
interacts with DNA to deposit CenH3CENP-A
CENP-N
P
CENP-T
MIS18BP
α β
CENP-W CENP-C
HJURP
CENP-B
Deposition
CenH3CENP-A–H4
HJURP and MIS18α/β
in late mitosis
CENP-C
H3.3 eviction?
CENP-B
P
Histone
mark?
M
e Kinetochore dissassembly (G1 phase)
b Kinetochore assembly (late S–G2 phase)
CENP-T–CENP-W recruitment
FACT
CENP-T
CENP-W
H2A–H2B
pool reduced
G1
CENP-T
CENP-T
CENP-N
G2
Kinetochore
assembly
Disassembly of the CenH3CENP-A
deposition machinery; degredation
of HJURP and MIS18?
CDK1/2
P
MIS18BP
P
α β
HJURP
CENP-W
Degradation?
CENP-C
CENP-W
CENP-C
CENP-N
CENP-B
S
CENP-B
f CENP-N recruitment (G1–S phase transition)
CENP-N recruitment facilitated by decompaction of
centromeric chromatin
a Replication of centromeric chromatin (late S phase)
CenH3CENP-A dilution: equal distribution of CenH3CENP-A to both daughter strands
CENP-N
?
H2A–H2B
CENP-T
Parental
H3–H4
dimers
Parental
H3–H4
tetramer
FACT
CENP-N CENP-W
CENP-B
(CenH3CENP-A–H4)2
(H3.1–H4)2
Old histone PTM
(H3.3–H4)2
Chaperone
Recycling
?
1
1
α-satellite DNA
2
Disruption
New
new H3.1–H4/ histone
H3.3–H4 dimers PTM
Gap?
CenH3CENP-A
MCM2
Recycling
H3.3
H3.1
Replication fork progression
MCM2–7
2
CENP-C
CAF1
De novo
complex assembly;
recycling?
H3.1
HIRA
De novo
complex assembly
H3.3
Gap?
De novo assembly
ASF1
H2A–H2B
Recycling;
de novo
assembly
H3.3
H3.1
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Reviews | Genetics
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◀ Figure 5 | Dynamics of centromeric chromatin composition during the cell cycle.
Centromeric chromatin undergoes major changes during the cell cycle. a | During late S
phase, centromeric chromatin is replicated. H3.1, H3.3 and CenH3CENP‑A nucleosomes are
disrupted as the replication fork progresses and old histones are recycled in a process
involving MCM2 and antisilencing function 1 (ASF1)113,114. New H3.1 (and possibly H3.3)
is deposited through mechanisms involving the dedicated chaperone complexes
indicated. CenH3CENP‑A is diluted onto both daughter strands, and H3.1 and/or H3.3 are
deposited to fill the gaps (gap-filling model). The H2A–H2B pool is depleted (possibly
handled by the facilitates chromatin transcription (FACT) complex). b | The CENP‑T–
CENP‑W complex is recruited de novo at late S phase–G2 phase20 in a process involving
the chaperone FACT21, possibly by a dynamic mechanism between H2A–H2B and
CENP‑T–CENP‑W binding to FACT21. c | Centromeres are licensed in late G2 phase in a
process involving the MIS18 complex. MIS18 and Holliday junction recognition protein
(HJURP) are dephosphorylated site-specifically and sequentially recruited concomitant
with reduced cyclin-dependent kinase 1 (CDK1) and CDK2 (CDK1/2) activity. d | MIS18α
and MIS18β are phosphorylated site-specifically by Polo-like kinase 1 (PLK1) and
recruited to centromeres. HJURP dimerizes and interacts with DNA and deposits
CenH3CENP‑A. It also interacts directly with CENP‑C and possibly helps to reposition
CENP‑C at centromeric chromatin. H3.3, as a placeholder for CenH3CENP‑A, is possibly
evicted in the process. e | Kinetochores are disassembled and MIS18 and HJURP leave the
centromere. f | CENP‑N has a high turnover at centromeres and interacts directly with
the Arg–Gly (RG)‑loop of CenH3CENP‑A nucleosomes during a conformational change.
PTM, post-translational modification.
deposition. In human cells, MIS18 recruitment relies
on phosphorylation ­mediated by Polo-like kinase 1
(PLK1)128.
In humans, CENP‑C interacts directly with
MIS18BP1 and may be involved in MIS18 recruitment 129. However, MIS18BP1 can associate with reconstituted human chromatin in vitro independently of
CENP‑C130. In C. elegans, CENP‑C recruitment follows
the recruitment of MIS18BP1 (known as KNL‑2 in
C. elegans)131, and HJURP is required for CENP‑C accumulation at synthetic centromeres in humans26. Whether
this is also the case for endogenous centromeres remains
to be elucidated. Interestingly, Cal1, the fly orthologue of
HJURP, is the limiting factor for CenH3CID and CENP‑C
deposition132. As it is not a homologue of HJURP, this
suggests that the feedback between CENP‑C and the
respective CenH3 chaperone has an important role.
Indeed, taken together, this argues for the existence of
a complex feedback mechanism involving CenH3CENP‑A,
HJURP, CENP‑C and the MIS18 complex that plays a
crucial role in the timely recruitment of CenH3CENP‑A.
Centromeric localization of human MIS18BP1 is coupled to its phosphorylation status133. Cyclin-dependent
kinase 1 (CDK1) phosphorylates MIS18BP1, keeping it
in an inactive state between the G1 and G2 phases. On
mitotic entry, MIS18BP1 becomes dephosphorylated,
which is required for its centromeric recruitment, akin
to HJURP19. We now take a look at cell cycle-dependent
HJURP dynamics and its involvement in CenH3CENP‑A
deposition during telophase and early G1 phase.
Heterotypic
A nucleosome that has
different variants of H3, H2A
or H2B.
HJURP-mediated CenH3CENP‑A deposition
The CCAN has a role in maintaining CenH3CENP‑A levels
at centromeres63 — particularly the complex of CENP‑H,
CENP‑I, CENP‑K and CENP‑M50 — and most CCAN
components are recruited to centromeres during G2
phase and mitosis. Because these factors are downstream
of CENP‑C134, they may stabilize CenH3CENP‑A rather
than contribute to its loading. Human HJURP localizes
to centro­meres at the time of CenH3CENP‑A deposition,
between telophase and early G1 phase48. The MIS18
complex may contribute to the timely recruitment of
HJURP to centromeres through its MIS18α and MIS18β
subunits19,133. MIS18β interacts with the first structural
repeat region of the C terminus of HJURP135, whereas
MIS18BP1 interacts with the central region of HJURP136.
CenH3CENP‑A deposition requires HJURP dimerization137,
possibly to facilitate the formation of a CenH3CENP‑A
nucleosome octamer. HJURP binds to CenH3CENP‑A using
its N‑terminal domain138, and then escorts CenH3CENP‑A
to the centromere for deposition47,48.
HJURP interacts with DNA through a specialized
domain19, which is essential to deposit CenH3CENP‑A at
centromeres (FIG. 5), highlighting that HJURP is not
merely escorting CenH3CENP‑A, but plays an active part
in CenH3CENP‑A deposition. Because HJURP has a propensity to interact with non-canonical DNA — that is,
DNA structures other than the B-helix 139 — dissecting
this interaction is an exciting avenue for future research.
HJURP interacts with CENP‑C26 and is implicated
in physically expanding and reordering centromeric
chromatin through an interaction with MIS18BP1 and
CENP‑C136. Thus, HJURP works across the nucleosomal
scale, not only escorting and depositing CenH3CENP‑A but
also influencing centromere and kinetochore structure by
reorganizing CENP‑C and MIS18 (REFS 26,136).
CDK kinases control the timely recruitment of HJURP
to centromeres in late mitosis by changing the phosphory­
lation status of the chaperone itself 19,135. CenH3CENP‑A
post-translational modifications are important for its
depo­sition. S68 phosphorylation of CenH3CENP‑A prevents
an interaction with HJURP in the pre-deposition complex,
helping to prevent premature loading140. The crystal structure at the interface of CenH3CENP‑A and HJURP shows that
this residue lies in the histone-­variant binding domain of
HJURP141. K124 ubiquitylation of CenH3CENP‑A has been
suggested to have a role in CenH3CENP‑A deposition by controlling the stability of the CenH3CENPA–HJURP complex
through a regulatory mechanism involving the cullin 4
(CUL4) ubiquitin ligase142, but the details of this must
be dissected further in future work. In D. melanogaster,
the stability of the CenH3CID–Cal1 complex is governed
by a ubiquitin-dependent mechanism143. Condensins
are additional factors that are required for CenH3CENP‑A
nucleosome assembly. They impose a certain structure on
chromatin, as demonstrated in Xenopus laevis144, in which
condensins are required for efficient CenH3CENP‑A deposition. We now examine scenarios in which these processes
go awry, leading to disease.
Centromere misregulation and disease links
In human cells overexpressing CenH3CENP‑A, DAXX is
involved in CenH3CENP‑A localization to ectopic loci115
(FIG. 6). This results in ectopic heterotypic (containing
both CenH3 CENP‑A–H4 and H3.3–H4) CenH3 CENP‑A
nucleosomes, whereas centromeric CenH3CENP‑A nucleo­
somes are usually homotypic (containing two copies
of CenH3CENP‑A–H4)53,109,115. The cell cycle regulation of
DAXX differs to that of HJURP, and thus the timing and
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a
Normal condition
HJURP-mediated CenH3CENP-A deposition at
centromeres coupled to telophase–G1 phase
CenH3CENP-A overexpression
DAXX-mediated CenH3CENP-A deposition at ectopic sites
not coupled to telophase–G1 phase
Histone pool
Histone pool
HJURP
Homotypic
CenH3CENP-A–
CenH3CENP-A
?
Increased
HJURP
pool
Homotypic
CenH3CENP-A–
CenH3CENP-A
HJURP
CenH3CENP-A–H4
DAXX
Heterotypic
CenH3CENP-A–H3.3
DAXX
H3.3–H4
H3.3–H4
Asymmetric
nucleosome
with possible
functional
consequence
b
Neocentromere formation
after genetic instability and
genome rearrangement
Neocentromere
Neocentromere formation and
cis-inactivation of endogenous centromere
Neocentromere formation
rescues chromosome after
endogenous centromere loss
Neocentromere
Endogenous
centromere
Endogenous
centromere
deactivated
Centromere
loss
Neocentromere
CENP‑A
Figure 6 | Ectopic CenH3CENP‑A deposition in cells overexpressing CenH3CENP‑A. a | Under normal Nature
conditions,
CenH3
Reviews
| Genetics
CENP‑A
is deposited by Holliday junction recognition protein (HJURP) at centromeres (homotypic CenH3
nucleosomes), whereas
death domain-associated protein 6 (DAXX) deposits homotypic H3.3 nucleosomes at other chromosome regions (left).
CenH3CENP‑A overexpression in human cells leads to ectopic enrichment at sites of active histone turnover in a process involving
a heterotypic tetramer containing CenH3CENP‑A–H4 with H3.3–H4. Ectopic localization of this particle depends on the H3.3
chaperone DAXX rather than the dedicated CenH3CENP‑A chaperone HJURP (right). b | Left panel, neocentromeres can form
during genetic instability and genome rearrangement. Neocentromeres can then rescue the acentric chromosome part.
Middle panel, neocentromeres can form on chromosomes, but only one centromere stays active. The other centromere is
deactivated by an unknown mechanism. Right panel, if endogenous centromeres are lost, neocentromeres can form at
ectopic regions to rescue the chromosome during cell division. Part a is adapted with permission from REF. 115, Elsevier.
control of ectopic and centromeric CenH3CENP‑A deposition might vary, arguing that neocentromere formation could be linked to cell cycle-dependent changes
in CenH3CENP‑A expression and/or deposition. Future
research should aim to analyse which histone modifications the particles carry on the different H3 variants
in one nucleosome. Later during the cell cycle, ectopic
heterotypic CenH3CENP‑A particles might be replaced by
homotypic particles, which could drive neocentromere
formation, or might be replaced by homotypic H3.3–H4
nucleosomes in other cases to prevent neocentromere
formation. This shows how changes in histone-­variant
levels can disrupt chromatin dynamics and lead to
different phenotypes by resulting in promiscuous functionalities of a histone chaperone and by causing changes
in chromatin landscape and architecture. Overexpression
of CenH3CENP‑A has been observed in several cancers145–148,
and CenH3 overexpression promotes its incorporation at
ectopic sites throughout chromatin in D. melanogaster 149
and S. cerevisiae150. In G1 phase, proteasome-mediated
degradation leads to the elimination of mislocalized
CenH3CID in D. melanogaster, and in S. cerevisiae the E3
ubiquitin ligase Psh1 is responsible for the eviction of
ectopic CenH3Cse4 (REF. 151) in G1 phase, demonstrating
that cells have another cell cycle-regulated layer of control
over the spatial distribution of CenH3.
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A copy number variation has been reported in
the genes encoding CenH3CENP‑A and HJURP in Mus
musculus 152 and, interestingly, the expression level of
the chaperone HJURP is connected to certain c­ ancer
pheno­types: its misregulation leads to chromosome
instability and mitotic defects139. Copy number vari­
ations have been linked to cancer propensity in mice153,
but aspects relating to cell cycle regulation in these
mice have yet to be determined. Proper chromosome
segregation relies on chromosomes harbouring a
­single centromere, and aberrations of this can lead to
aneuploidy (abnormal chromosome numbers), which
is one of the hallmarks of cancer 154. During M phase,
neocentromeres can rescue chromosome fragments in
cells with chromosomal rearrangements155. Genome
instability is often concomitant with neocentromere
formation (FIG. 6), potentially resulting in aneuploidy 156.
In one particular case, a neo­centromere-containing
giant chromosome that was devoid of α-satellites was
observed157: the additional sequences stemmed from
another chromosome that contained various oncogenes158. Other reports show that neocentromeres form
in the absence of major chromo­some rearrangements,
with little impact on human health. Neocentromeres
can be stably inherited through multiple generations159,
indicating mitotic and meiotic stability. Alternatively,
neocentromeres can form in the absence of chromosome alterations, concomitant with the inactivation of
the endogenous centromere160.
Neocentromeres are marked by CenH3 CENP‑A
(REF. 160) and contain all of the factors and features found
at centromeres, except for CENP‑B and ­α-satellites155.
Although centromere positions are generally stable,
genomic alterations can remove or disrupt centromere
function, and neocentromeres can restore faithful
chromo­some segregation161. In rare cases, neocentro­
meres form on otherwise normal chromosomes, without a deletion but concomitant with inactivation of
the endogenous centromere159. They are particularly
common on specific chromosome regions11, but among
neo­centromeres found on the same chromosomal band,
the sequences harbouring CenH3CENP‑A are unique from
case to case. A specialized chromatin environment
may favour neocentromere formation, and a threshold
amount of ectopic CenH3CENP‑A might be required to
establish neocentromeres29. It will be crucial to determine how the establishment and maintenance of these
sites are controlled during the cell cycle.
Synthetic neocentromeres — which can be established by using the LacO-system, for example28 — have
helped to elucidate the cell cycle dynamics of some (neo)
centromeric components. For example, this system
determined that CENP‑T recruitment follows that of
CENP‑C26. CenH3CID is sufficient to establish ectopic
kinetochores in D. melanogaster 28, and recruiting centromeric factors downstream of CenH3CENP‑A at ectopic sites
during the cell cycle leads to the recruitment of other
centromere components and a functional kinetochore28.
Neocentromeres have been engineered on endogenous
centromere removal162, and in chicken cells they typically form near the original centromere163 as a result
of ‘seeding’ events around the original excised site. In
D. melanogaster, ectopic centromeres have been generated following induced chromosome breakage, which
leads to neocentromere formation in heterochromatic
regions, including PHC164, suggesting that particular
chromatin environments favour de novo centromere
formation. Interestingly, CENP‑B was important for
the centromere establishment on a HAC165. A degree
of heterochromatinization is concomitant with de novo
centromere formation on HACs24, and neocentromere
formation requires functional heterochromatin proteins
in S. pombe162, which indicates that the chromatin environment at ectopic sites influences neocentromere establishment. Some HACs show aberrant timing of mitosis
and can cause an anaphase lag, suggesting that variable
centromeric DNA content or assembly can influence the
mitotic behaviour of artificial chromosomes166. Further
dissection of the nature of neocentromeric chromatin
and its cell cycle regulation in terms of establishment and
maintenance will be crucial in future studies.
Conclusions
Our knowledge about the architecture and cell cycle-­
dependent regulation of centromeres has greatly
expanded since the discovery of key centromeric factors.
A picture of centromere and kinetochore dynamics during the cell cycle, with its network of components, has
emerged thanks to powerful cellular assays. However,
one should also consider the evolutionary turnover of
kinetochore proteins. Although CenH3 nucleosomes
are key to the centromeric architecture in most eukary­
otes, some species do not have CenH3, whereas other
components of the centromere are conserved167,168. For
instance, in humans there are two parallel routes to
forming kinetochore–microtubule interactions through
CENP‑C or CENP‑T, but the CENP‑T route is absent
in D. melanogaster and most members of Diptera169,170.
Thus, the complex feedback mechanism between
the different components, which is regulated at each
cell cycle stage, can be exploited differently between
species, making some of the components redundant.
Furthermore, distinct cell types in a single organism
exhibit different cycle dynamics during development.
Future studies to deepen our understanding of the
higher-order organization of centromeres will therefore require technologies for examining distinct cell
cycle dynamics at the whole-animal level. Finally, going
beyond the protein and RNA level, it is becoming possible to unravel the biochemical features of underlying
DNA repeat sequences8,10.
The network of proteins at the centromere and
kinetochore underlines a complex feedback mechan­
ism coupled to the cell cycle stages. One has to consider the availability of the components during each cell
cycle phase, which is linked to transcription and gene
expression, and the biochemical mechanisms involved
in recruitment, such as cell cycle checkpoints involved in
the spatiotemporal recruitment of several centromeric
components19,133. It is important to consider that a component needs to be recruited at the right time with the
right biochemical environment to exert its function. In
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addition, although further dissection of the mechanism
of centromere and kinetochore regulation offers scope
for more discoveries, a major challenge lies in understanding how its higher-order organization integrates
with the dynamics of other cell cycle-regulated events,
such as duplication of centrioles, which are micro­
tubule-organizing centres involved in the formation of
the mitotic spindle171.
The involvement of centromeres in disease, such as
links to aberrant expression levels of some centromeric
factors, may mark these centromeric factors as actionable
1.Flemming, W. Zellsubstanz, Kern und Zelltheilung
(in German) (F.C.W. Vogel, Leipzig, 1882).
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Acknowledgements
We thank I. A. Drinnenberg, D. Ray-Gallet, D. Filipescu and D.
Fachinetti for useful discussion and proofreading of this Review.
We also thank H. Tachiwana for drawing the nucleosomal structure illustrations. S.M. thanks the R. Rodriguez laboratory for
support. This work was supported by la Ligue Nationale contre
le Cancer (Equipe labellisée Ligue), the European Commission
Network of Excellence EpiGeneSys (HEALTH‑F4‑2010‑257082),
t h e E u ro p e a n R e s e a rc h C o u n c i l ( a d va n c e d g ra n t
2009‑AdG_20090506 ‘Eccentric’), the European Commission
(large-scale integrating project FP7_HEALTH‑2010‑259743
‘MODHEP’), the French National Research Agency (ANR)
(‘ChromaTin’ ANR‑10‑BLAN‑1326‑03, ANR‑11‑LABX‑0044_
DEEP and ANR‑10‑IDEX‑0001‑02 PSL; and ‘CHAPINHIB’
ANR‑12‑BSV5‑0022‑02) and the Aviesan Instituts thématiques
multi-organismes (Aviesan-ITMO) cancer project ‘Epigenomics
of breast cancer’. S.M. was also supported by the Marie Curie
Initial Training Network (Nucleosome 4D), and La Fondation
pour la recherche médicale.
Competing interests statement
The authors declare no competing interests.
208 | MARCH 2017 | VOLUME 18
www.nature.com/nrg
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