Biochem. J. (2010) 428, 147–161 (Printed in Great Britain)
147
doi:10.1042/BJ20100151
REVIEW ARTICLE
Cohesin: a regulator of genome integrity and gene expression
Katherine M. FEENEY1 , Christopher W. WASSON1 and Joanna L. PARISH2
Bute Medical School, Bute Building, University of St Andrews, St Andrews, Fife KY16 9TS, Scotland, U.K.
Following DNA replication, chromatid pairs are held together by
a proteinacious complex called cohesin until separation during
the metaphase-to-anaphase transition. Accurate segregation is
achieved by regulation of both sister chromatid cohesion
establishment and removal, mediated by post-translational
modification of cohesin and interaction with numerous accessory
proteins. Recent evidence has led to the conclusion that cohesin
is also vitally important in the repair of DNA lesions and control
of gene expression. It is now clear that chromosome segregation
is not the only important function of cohesin in the maintenance
of genome integrity.
INTRODUCTION
due to the intertwining or catenation of DNA strands during
replication and that DNA topoisomerase II was required for the
separation of chromatids at anaphase [4]. Indeed, inhibition of
topoisomerase II has shown that DNA catenation between sister
chromatids does exist and needs to be resolved to allow accurate
sister chromatid separation, but not mitotic exit [5–7]. Although
these studies show that DNA decatenation is required for efficient
chromatid segregation, cohesion of DNA molecules can, however,
be established in the absence of catenation [8]. More compelling
evidence for the mechanism of sister chromatid cohesion comes
from the isolation of protein complexes essential for cohesion
but that play no part in the mechanism by which catenations
are removed. These highly conserved protein complexes contain
multiple core structural proteins along with several associated
regulatory subunits.
The accurate separation of sister chromatids into daughter cells
is critical for maintaining genome integrity. A loss of genome
integrity via W-CIN (whole chromosome instability) has been
implicated in a variety of pathologies including reproductive
failure, and mental and physical abnormalities [1]. However, the
link between W-CIN and cancer susceptibility has been difficult
to define. Aberrant chromosome numbers, or aneuploidy, is one
of the molecular hallmarks of cancer cells but little evidence
has been provided to conclude that aneuploidy can lead to
cancer rather than it being a consequence of tumorigenesis [2].
Several mouse models of W-CIN have been used to determine
whether aneuploidy is a cause or effect of tumorigenesis. These
models have provided strong evidence that mutation of several
of the genes involved in the regulation of accurate chromosome
segregation results in higher cancer susceptibility and thus
provides a link between aberrant chromosome segregation and
cancer formation [3].
The mechanism by which replicated sister chromatids are
accurately segregated into daughter cells initially depends on their
physical association with each other, a process termed cohesion.
Sister chromatid cohesion facilitates capture by a bipolar mitotic
spindle and segregation to opposing poles, resulting in the
formation of two genetically identical daughter cells. Without
physical association of sister chromatids, the opposing forces
required for bipolar separation of chromatids would not exist.
In this case premature separation of chromatids could occur
before bipolar attachment resulting in inaccurate chromosome
segregation and genome instability. It was long thought that sister
chromatids were physically associated until the onset of anaphase
Key words: chromosome segregation, cohesion, DNA damage,
heterochromatin, transcriptional insulation.
MOLECULAR ARCHITECTURE OF THE COHESION COMPLEX
The core members of the protein complex that facilitates
chromosomal cohesion during mitosis were first isolated using
a yeast screen for temperature-sensitive mutants that lose
chromosomes at high frequency. Four genes were isolated using
this approach: two known genes belonging to the SMC (structural
maintenance of chromosomes) family of ATPases (SMC1 and
SMC3), and two novel genes, termed SCC1 and SCC3 due to
their involvement in sister chromatid cohesion [9]. A second
key finding of that study was that the Scc1 subunit of the
cohesin complex was specifically degraded at the metaphase-toanaphase transition by the APC (anaphase-promoting complex),
promoting sister chromatid separation [9]. At the same time,
Abbreviations used: APC, anaphase-promoting complex; APC/C, APC/cyclosome; ATM, ataxia telangiectasia mutated; Bub1, budding uninhibited by
benzimidazole 1; 3C, chromosome conformation capture; CDC, cell division cycle; CdLS, Cornelia de Lange syndrome; Chk1, checkpoint kinase 1;
ChIP, chromatin immunoprecipitation; Chl1, chromosome loss 1; CTCF, CCCTC-binding transcriptional insulator protein; CTF, chromosome transmission
fidelity; DMR, differentially methylated region; DSB, double-stand break; ECO/ESO/ESCO, establishment of mitotic (sister) chromatid cohesion; FRET,
fluorescence resonance energy transfer; H19, H19 fetal liver mRNA; Hcy, homocysteine; HR, homologous recombination; IGF2, insulin-like growth
factor 2; IR, ionizing radiation; KSHV, Kaposi’s sarcoma-associated herpes virus; LCR, locus control region; LTR, long terminal repeat; MAT, mating
type; Mcd, mitotic chromosome determinant; Mec1, mitosis checkpoint entry 1; Mre11, meiotic recombination 11; NHEJ, non-homologous end-joining;
NIPBL, Nipped-B homologue; PCNA, proliferating-cell nuclear antigen; PDS, precocious dissociation of sisters; Plk1, polo-like kinase 1; PP2A, protein
phosphatase 2A; RAD, radiation-sensitive; rDNA, ribosomal DNA; Rec8, recombination 8; RFC, replication factor C; SA/STAG, stromal antigen; SCC, sister
chromatid cohesion; Sgo1, Shugoshin 1; SIR, silencing protein; SMC, structural maintenance of chromosomes; Swi6, switching deficient 6; Tel1, telomere
maintenance 1; W-CIN, whole chromosome instability; Wpl/Wapl/Wapal, wings apart-like.
1
These authors contributed equally to this work.
2
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2010 Biochemical Society
148
Table 1
K. M. Feeney, C. W. Wasson and J. L. Parish
Cohesin subunits and associated proteins
Alternative names are given in parentheses. Cap, chromosome-associated protein; CUT, Cell untimely torn; Esp1, extra spindle pole bodies 1; Mis, minichromosome instability 4.
Process
S. cerevisiae
S. pombe
D. melanogaster
H. sapiens
Cohesin complex
Smc1
Smc3
Mcd1 (Scc1)
Scc3
Scc2
Scc4
Eco1 (Ctf7)
Pds5
Rad61
Pds1
Esp1
Psm1
Psm3
Rad21
Psc3
Mis4
Ssl3
Eso1
Pds5
Wpl1
Cut3
Cut1
Smc1
Smc3 (Cap)
Rad21 (Verthandi)
SA1 (Stromalin)
Nipped-B
Mau-2 (CG4203)
Eco
Pds5
Wapl
Pimples
Separase
Smc1
Smc3
Rad21
SA1/SA2/STAG3
NIPBL
Scc4
Esco1/Esco2
Pds5
Wapal
Securin
Separase
Cohesin-loading proteins
Cohesion establishment/anti-establishment, protection
Cohesin cleavage
an independent study also identified Scc1 [also known as
Mcd1 (mitotic chromosome determinant 1) in Saccharomyces
cerevisiae (budding yeast) and as Rad21 (radiation-sensitive 21)
in Schizosaccharomyces pombe (fission yeast) and vertebrates] as
a Smc1-associated protein involved in sister chromatid cohesion
[10]. It was later shown that the products of these four genes,
Smc1, Smc3, Scc1 and Scc3, exist in a complex called cohesin
that is central to the mechanism of sister chromatid cohesion.
Table 1 gives the names of the cohesin subunits and associated
protein in yeasts, Drosophila and mammals.
The SMC family of proteins are highly conserved in
prokaryotes and eukaryotes and have the same general structure.
SMC proteins are large (1000 –1300 amino acids) and contain
two nucleotide-binding motifs, known as Walker A and Walker
B motifs, situated at opposing ends of the polypeptide. A large
coiled-coil motif is situated between the Walker A and Walker
B motifs and this folds into an antiparallel formation creating
an ATP-binding head domain. At the other end of the folded
protein is a hinge domain that facilitates either homo- or heterodimerization of SMC molecules creating a V-shaped structure
[11,12]. Electron micrographs of SMC dimers show that the
coiled-coil arms of each molecule are approx. 50 nm in length
and that a high degree of flexibility exists in the hinge, resulting
in a wide variability of conformational states between dimers
[12,13]. The cohesin complex consists of a heterodimer of the
Smc1 and Smc3 subunits, assembled as described above. The
kleisin family member Scc1 binds to both the Smc1 and Smc3
head domains creating a closed ring structure that is large enough
to encircle chromatin fibres, while the fourth subunit of cohesin,
Scc3, associates with the complex via interaction with Scc1 [11]
(Figure 1). There are known homologues of Scc3 in vertebrate
cells, SA1 and SA2 (stromal antigen 1 and 2; also known as
STAG 1 and 2; a third homologue STAG3 functions in meiosis
and is only expressed in germinal cells). Interestingly, it has been
shown that cohesin complexes contain either SA1 or SA2, but
not both [14,15], although the function of these distinct protein
complexes has not been defined. However, it has been suggested
that SA1 and SA2 play a role in the establishment of cohesion at
the telomeres and centromeres respectively, potentially revealing
distinct roles for SA1- and SA2-associated cohesin [16].
ASSOCIATION OF THE COHESIN RING WITH DNA
The first studies looking at the mechanism by which cohesin
associates with DNA showed that the SMC proteins alone can associate with DNA in an ATP-independent manner [17,18]. A further study showed that cohesin can facilitate intermolecular asso
c The Authors Journal compilation c 2010 Biochemical Society
Figure 1
Structure of the cohesin complex
The SMC proteins contain nucleotide-binding motifs situated at opposing ends of the
polypeptide, which fold creating a hairpin structure with a 50 nm coiled-coil domain situated
between the ATP-binding head domain and hinge domain. Smc1 (green) and Smc3 (orange)
together form a V-shaped heterodimer via hinge–hinge interaction. The C-terminal domain (C)
of Scc1/Rad21 (purple) associates with the head domain of Smc1, whereas the N-terminal
domain (N) associates with the head domain of Smc3, thus forming a closed ring structure. The
forth subunit of cohesin (Scc3/SA1/SA2; brown) binds directly to Scc1.
ciation of DNA molecules rather than the intramolecular knotting
directed by the related condensin complex [19]. These results
led to the hypothesis that the Smc1–Smc3 heterodimer acts
as an intermolecular bridge between two DNA molecules,
providing the cohesive force that holds chromosomes together.
Initially, this intermolecular bridging was suggested to be
regulated or maintained by the non-SMC subunits unique to
cohesin, thus explaining the intermolecular bridging facilitated by
cohesin rather than the intramolecular interactions facilitated by
condensin. However, the bridging hypothesis has been modified
further following structural and biochemical studies of the cohesin
complex that strongly indicate the formation of a ring-like
structure between the V-shaped SMC heterodimer, joined at either
end by the cleavable Scc1 subunit [11,20,21]. From this it was
proposed that the cohesin complex functions by encircling DNA
strands, trapping them together rather than forming a molecular
bridge between strands.
The requirement for ATP-binding and hydrolysis in cohesion
establishment and release has also been a contentious debate.
Cohesin regulation and function
Although some in vitro studies have shown that SMC proteins
can associate with DNA via the C-terminal domains in an ATPindependent manner [17,18], in vivo studies have shown that
ATP hydrolysis is required for the association of cohesin with
DNA [22,23]. Evidence suggests that two ATP molecules are held
between the SMC heads, which separate and bind a single Scc1
molecule upon ATP hydrolysis [23]. This could be triggered by the
presence of DNA and therefore result in the entrapment of DNA
strands within the now tripartite ring. Although the cohesionspecific Smc1–Smc3 heterodimer has been shown to bind to
DNA in the absence of other proteins and ATP, this association
has a very low affinity suggesting that it does not occur in vivo,
whereas inclusion of the Scc1 subunit in the complex substantially
increases binding affinity [24]. In addition, the heterotrimeric
complex associates with cruciform DNA with a much higher
affinity than unstructured double-stranded DNA, indicating that
the presence of secondary structure is important for the association
of cohesin with DNA [24]. Indeed, an independent study showed
that the cohesin complex bound to chromatin with a much higher
affinity than the Smc1–Smc3 heterodimer alone, indicating that
the holocomplex requires secondary structure elements to bind
to DNA with high affinity [25]. Electron micrographs of SMC
heterodimers show that the arms of the molecule are approx. 50–
60 nm in length. Inclusion of the Scc1 subunit to form a tripartite
ring could allow the encircling of nucleosomal DNA, which is
approx. 10 nm in diameter [11]. The topological entrapment of
DNA within cohesin rings is supported by studies showing that
cleavage of the Scc1 or Smc3 subunits results in dissociation
of cohesin from DNA and premature loss of cohesion [26].
Whether one cohesin ring embraces two chromatin fibres to hold
them together or each fibre is encircled by a cohesin ring that
interconnect holding strands together has been hard to determine
(Figure 2).
The ‘ring’ model (Figure 2A) has historically dominated.
This model suggests that one single cohesin ring can encircle
two DNA strands thus trapping them together [20,27,28] and is
supported by experimental results showing that linearization of
circular minichromosomes in yeast, by restriction digest, causes
dissociation of cohesin from DNA [8]. In addition, interactions
between multiple cohesin complexes were undetectable using
FRET (fluorescence resonance energy transfer) technology and
in co-immunoprecipitation experiments in cells expressing two
differently tagged copies of the same cohesin subunit [11,29].
Furthermore, chemical cross-linking of the tripartite cohesin
rings produces DNA–cohesin complexes that are insensitive to
protein denaturation, thus providing strong evidence that cohesion
between DNA molecules is mediated by a single cohesin ring that
embraces both sister chromatids [30].
An alternative model of how cohesin physically holds
chromatids together is the two-ring ‘handcuff’ model (Figure 2B)
in which each sister is encircled by one cohesin ring and two
rings then associate with each other or interconnect resulting
in cohesion of DNA strands. Although the ring model has
been favoured in recent years, new evidence has resurrected the
handcuff model. In contrast with the studies described above,
self interaction of the Scc1 subunit was detected in a yeast twohybrid assay and was confirmed in mammalian cells expressing
tagged proteins, suggesting the presence of a higher-order cohesin
complex [31]. This is consistent with a study showing that cohesin
is able to embrace only one sister chromatid at transcriptionally
silent heterochromatin regions, which have a larger diameter due
to greater chromatin compaction [32]. However, it is possible that
this mechanism of cohesion is specific to transcriptionally silent
regions and that transcriptionally active regions can be encircled
by a single ring.
Figure 2
149
Models of sister chromatid cohesion
(A) Ring model: one cohesin ring encircles two sister chromatids (the DNA is wrapped around
nucleosomes and is 10 nm in diameter). (B) Handcuff model: each sister chromatid is encircled
by separate cohesin rings and two rings then associate with each other (1,2) or interconnect (3)
resulting in cohesion of sister chromatids.
COHESIN LOADING
In lower eukaryotes cohesin associates with DNA at the end of G1 phase [9,10,33]. In contrast, cohesin has been shown to assemble
on to DNA in mammalian cells following nuclear envelope
reformation during telophase [34]. This difference may be due
to the different regulatory mechanisms in place to maintain and
remove cohesin, or may be due to differences in the functions of
cohesin outside of its canonical role in chromosome segregation
(discussed below). It has been demonstrated that the interaction of
cohesin with DNA in both higher and lower eukaryotes becomes
more stable as cells progress from G1 to S to metaphase [35,36]
and although the timing differs, cohesin loading in all organisms
appears to require ATP hydrolysis [22,23,37] and the Scc2–Scc4
protein complex (Nipped-B in Drosophila) [38–41]. Although
the specific mechanism of Scc2–Scc4-mediated cohesin loading
remains unclear, it has been proposed that the Scc2 protein
c The Authors Journal compilation c 2010 Biochemical Society
150
K. M. Feeney, C. W. Wasson and J. L. Parish
interacts transiently with cohesin to promote ATP hydrolysis by
the SMC heads, leading to SMC dimer hinge opening or Scc1
dissociation [22]. Interestingly, the Scc2–Scc4 complex does
not play a role in the maintenance of cohesin association with
DNA, as cohesin rapidly and independently moves away from
chromosomal-loading sites to areas of convergent transcription
[33].
Although the loading of cohesin is generally Scc2–Scc4
dependent, there are some instances where cohesin loading
appears independent of Scc2–Scc4. In human cells the Scc2–
Scc4 complex has been shown to dissociate from chromosomes
during mitosis, which is presumed to inhibit the reassociation
of cohesin until progression through to telophase, when Scc2–
Scc4 and cohesin rebind [34,41]. However, during pre-anaphase
centromere ‘breathing’, a process that occurs as sister chromatids
transiently split under tension from the mitotic spindle, cohesin is
reloaded on to transiently separated chromatids in an Scc2–Scc4independent manner [42], suggesting that an alternative and as
yet undetermined mechanism of cohesin loading may exist at this
stage of the cell cycle.
COHESION ESTABLISHMENT DURING DNA REPLICATION
Traditional sister chromatid cohesion can only be established
during DNA replication. However, cohesin associates with DNA
early in the cell cycle and therefore cohesion establishment
is thought not to require new cohesin loading [43]. Cohesion
establishment, but not maintenance, has been shown to be
dependent on the lysine acetyltransferase Eco1 [establishment
of cohesion 1; known as Eso in S. pombe and, Esco1 and Esco2
in humans)][44,45]. Eco1 physically associates with numerous
members of the DNA replication complex, including two highly
homologous RFC (replication factor C) clamp loader proteins,
Ctf18 (chromosome transmission fidelity 18) and Elg1 (enhanced
level of genomic instability 1), which each play an important
role in cohesion establishment [46,47], and PCNA (proliferatingcell nuclear antigen) [48], which genetically associates with
Ctf4 and is part of the replication progression complex [49].
Interestingly, Eco1 has also been shown to bind to the XPD
DNA-helicase family member, Chl1 [chromosome loss 1; DDX11
(DEAD/H box polypeptide 11) in humans], which plays a role
in cohesion establishment [50–53]. Like Eco1, Chl1 associates
with members of the DNA replication complex, such as RFC
and PCNA [48,52,54]. These results provide strong evidence that
cohesion establishment is intimately coupled to PCNA-dependent
DNA replication and that Eco1 may function to allow dissociation
of cohesin rings, or passage of the replication machinery through
cohesin rings, ensuring cohesion establishment and completion
of DNA replication. In support of this, it has been shown
that acetylation of cohesin is required for replication fork
processivity and in the absence of RFCCTF18 both replication and
cohesin acetylation are severely impaired, demonstrating that the
acetylation of cohesin is required for replication fork progression
[55]. Whether Eco1 alone or in collaboration with Chl1 facilitates
opening of a cohesin ring and passage of the replication fork
followed by resealing of the ring to encircle both sisters (the ring
model), or facilitates the tethering of new cohesin rings deposited
in S-phase to pre-loaded rings as the replication machinery passes
(the handcuff model) remains to be determined [56]. Interestingly,
it has been suggested that although cohesion establishment, or
rather re-establishment, during centromere breathing is Scc2–
Scc4 independent, Eco1 is required, providing evidence that Eco1dependent cohesion establishment can also occur in the absence
of DNA replication [42].
c The Authors Journal compilation c 2010 Biochemical Society
Eco1 was shown to acetylate components of the cohesin
complex in vitro some years ago [57], but the requirement for
this activity in cohesion establishment was controversial [58].
However, recent studies have shown that the Smc3 subunit of
cohesin is acetylated by Eco1 in vivo at two highly conserved
lysine residues [59–61]. Mutation of both of these lysine residues
within Smc3 inhibits acetylation and results in a lethal cohesion
defect. Conversely, mutation of both lysine residues to asparagine
or glutamine, which mimic the acetylated state, abolishes the
requirement for Eco1 expression, an otherwise essential protein
[61]. Acetylation of Smc3 by Eco1 only occurs at the onset
of S-phase and is suppressed during the G1 , G2 and mitotic
phases [60]. This may provide a mechanism by which cohesin
only becomes cohesive as the sister chromatid is synthesized. In
addition to Eco1, a second acetyltransferase called San, which
exists only in metazoans, has been shown to be important for
stabilizing centromeric sister chromatid cohesion during mitosis
[62]. Whether this acetyltransferase is important for establishment
or maintenance of centromeric cohesion remains unclear. It is
also possible that the acetyltransferase activity of San is required
for the maintenance of centromeric cohesion during centromere
breathing, a hypothesis that should be tested.
The cohesin-associated proteins Wpl1 [wings apart-like 1;
Rad61 in S. cerevisiae, Wapl in Drosophila and Wapal in
vertebrates] and Pds5 (precocious dissociation of sisters 5)
together form a complex [63,64] and play a role in the inhibition
of cohesion establishment, so called anti-establishment. In
S. cerevisiae, it has been shown that the Wpl1–Pds5 complex
associates with cohesin rings until Eco1-induced acetylation of
the Smc3 subunit occurs, leading to the hypothesis that the Wpl1–
Pds5 complex inhibits establishment when bound to cohesin and
that Eco1 functions to inhibit this interaction by acetylating Smc3
during S-phase [65,66]. This function of the Wpl1–Pds5 complex
appears conserved in human cells as processive fork movement
has been shown to require acetylation-induced dissociation of
Pds5 and Wapal [55]. In addition, depletion of Wapal has been
shown to prevent removal of cohesin from chromosome arms
during prophase, indicating that Wapal also functions to inhibit
cohesion re-establishment in higher organisms [63,64]. Studies
of Xenopus and human protein complexes have shown that Wapal
associates with Pds5 and cohesin via FGF (Phe-Gly-Phe) motifs,
which are predicted to bind HEAT [huntingtin, elongation factor
3, the PR65/A subunit of PP2A (protein phosphatase 2A) and the
lipid kinase Tor] repeat motifs found in Pds5 and the SA1/SA2
family of proteins. It is hypothesized that association of Wapal
with cohesin in this manner may induce conformation changes
in cohesin rings thereby facilitating release from chromatid arms
during prophase, a function that may not exist in yeast [67].
COHESIN REMOVAL DURING MITOSIS
Once all chromatids are captured by the bipolar mitotic spindle
and aligned on the metaphase plate, cohesion is dissolved by proteolytic cleavage of Scc1 by the activated cysteine protease separase, in a DNA-dependent manner [26,68]. Activation of
separase results from degradation of its inhibitory binding
partner securin by the ubiquitin ligase APC/C (APC/cyclosome),
which is activated following satisfaction of the spindle assembly
checkpoint [69] (see [70] for a review on the spindle assembly checkpoint). Although this has long been the established
model of cohesin removal during mitosis, much of the work
was performed in lower eukaryotes where evidence suggests that
cohesin remains associated with chromatids until anaphase onset.
In metazoa, cohesin is removed in a more complex and highly
Cohesin regulation and function
regulated two-step process. As cells progress from prophase
to metaphase the cohesion on chromosome arms is lost in a
pathway that is thought to be independent of Rad21 (Scc1)
cleavage by separase. However, centromeric cohesion is protected
and remains bound until anaphase onset when it is cleaved
by activated separase to allow segregation of chromosomes to
opposing spindle poles [34,71]. Loss of arm cohesion in metazoa
is dependent on the activity of Plk1 (polo-like kinase 1) [72–
74]. Plk1 can phosphorylate both the Rad21 and SA2 (Scc3)
subunits of cohesin in vitro [72,73]. In S. cerevisiae it has been
shown that phosphorylation of Scc1 by the Plk1 homologue Cdc5
(cell division cycle 5) enhances cleavage of Scc1 by separase
[75,76] and hyperphosphorylation of Scc1 at the centromeres
of metaphase chromosomes in Xenopus extracts has also been
reported [77]. It has also been reported that phosphorylation of
SA2 (Scc3) by Plk1 in vertebrate cells is important for the reduced
affinity and dissociation of cohesin from chromosome arms during
prophase [78].
Although the prophase pathway of cohesin removal was thought
absent in lower eukaryotes, a recent study has shown that this
may not be the case. Upon entry into mitosis, cohesin dynamics
in S. pombe appear to be related to those observed in metazoa
as the loss of a fraction of cohesin from chromosomes has been
observed as these cells enter mitosis [79]. In higher eukaryotes,
cohesin associates with chromatin in two distinct binding modes:
one stable, established in S-phase and removed by Scc1 cleavage,
and the other highly dynamic and thought to be required for
transcription regulation and DNA damage repair, as discussed
below [36]. This suggests that the existence of two distinct binding
modes of cohesin throughout the cell cycle may be conserved
in different model organisms. However, more work needs to be
done to determine the role of these binding modes of cohesin
in lower eukaryotes and whether similar pathways of cohesin
removal during mitosis are in place in these model systems. In
vertebrate cells, the cohesion-associated protein sororin may have
an important role in regulating the stable association of cohesin
and chromatin. Sororin has been shown to be important in sister
chromatid cohesion and is degraded by the APC/C in G1 of the
cell cycle [80]. Overexpression of sororin prevents resolution of
the sister chromatids at anaphase onset and depletion results in
precocious sister chromatid separation. In addition, depletion of
sororin results in a significant reduction in the stably bound pool
of cohesin that exists following cohesin establishment at S-phase
[81]. In the light of this result, it has been suggested that sororin
may function in the establishment of stably bound cohesin and
that degradation as cells enter G1 -phase might prevent the reestablishment of this stable pool of cohesin before initiation of
DNA replication.
The active protection of centromeric cohesin until anaphase
onset requires the recruitment of the ‘guardian spirit’ protein
Sgo1 [Shugoshin 1; Mei-S332 (meiotic from via Salaria 332)
in Drosophila] to the centromeres during prophase [82,83].
Besides its role in the protection of centromeric cohesion, Sgo1
is important for both microtubule stabilization and microtubuledependent tension between sisters [83] and has been shown to
accumulate on kinetochores until anaphase onset when bipolar
attachment and inner kinetochore tension is achieved. The role of
Sgo1 in tension sensing may be through its ability to recruit Plk1
to the kinetochores as it has recently been shown that depletion of
Sgo1 results in reduced accumulation of Plk1 on the kinetochores
and a subsequent reduction in the Plk1-dependent 3F3/2 phosphoepitope, important in the tension-sensing mechanism [84]. The
recruitment of Sgo1 to centromeres requires both the spindle
checkpoint kinase Bub1 (budding uninhibited by benzimidazole
1) and the Aurora B kinase [84–88], which has been shown
151
to phosphorylate Sgo1 in vitro [84]. Interestingly, depletion of
Bub1 or Aurora B causes Sgo1 to localize along the length of
the chromosomes rather than concentrate at the centromeres,
suggesting that these kinases are required for defining the region
on each chromosome where cohesion is to persist [84,86].
Interestingly, Aurora B has also been shown to play a role in
the recruitment of separase to the chromosomes at early stages
of mitosis [89]. Although the exact role of separase recruitment
early in mitosis is unclear, the authors of that study suggest that
separase may be required for the removal of arm cohesin during
the prophase pathway, but until further studies are completed this
remains speculative.
Centromeric cohesin is lost following depletion of Sgo1 by
RNA interference, resulting in premature chromatid separation.
Expression of a non-phosphorylatable SA2 protein in these cells
prevents loss of centromeric cohesion, providing evidence that
Sgo1 prevents phosphorylation of cohesin situated at the centromere [82]. Sgo1 binds to PP2A with high affinity and it is
thought that Sgo1 prevents the phosphorylation of cohesin by recruiting PP2A, rapidly reversing phosphorylation of cohesin by
Plk1, thereby inhibiting release [90–92]. Interestingly, prevention
of Scc1 phosphorylation by PP2A has been shown to block
cleavage of Scc1 only in the presence of Sgo1 [93] and it has been
suggested that Sgo1-dependent recruitment of PP2A to unattached
kinetochores inhibits separase activity [94]. Sgo1 protein levels
peak in mitosis and it undergoes APC/C-dependent degradation
at the onset of anaphase; it has been shown that APC/Cmediated Sgo1 degradation is necessary for mitotic exit and
accurate chromosome segregation [95]. However, a recent study
contradicts these findings and suggests that Sgo1 degradation
is not crucial for progression through mitosis [96]. Alternative
mechanisms to inactivate Sgo1 upon anaphase onset may exist and
should be investigated before the true role of Sgo1 degradation in
mitotic progression can be elucidated.
COHESIN AND THE CONTROL OF GENE EXPRESSION
In addition to its role in chromosomal segregation cohesin has
long been thought to have other cellular functions. This initial
hypothesis was sparked from evidence, at least in vertebrate
cells, that cohesin re-associates with chromosomes at the end
of mitosis, when conventional thinking would assume cohesin is
absent from the DNA [34]. This re-association of cohesin occurs
well before sister chromatid cohesion is established in the next cell
cycle and suggests that cohesin may have a function independent
of its role in cohesion. Furthermore, human diseases such as
CdLS (Cornelia de Lange syndrome) and Roberts syndrome
are associated with mutation of cohesin subunits such as Smc1
and Smc3, or the cohesin loader NIPBL [Nipped-B homologue
(Drosophila)] (Scc2) [97–99]. These diseases have phenotypes
that include growth and mental retardation, craniofacial anomalies
and microcephaly, but surprisingly defects in cell cycling are
limited and do not solely account for the severe phenotypes
observed, suggesting that other functions of cohesin are disrupted.
In addition, these mutations pose no increased predisposition
to carcinoma, providing further evidence that cohesin has other
cellular functions aside from its role in sister chromatid cohesion
and accurate chromosome segregation.
The initial evidence that cohesin is important in the
control of gene transcription came from a study showing
that the positioning of cohesin on chromatin was dramatically
affected by transcriptional activity in S. cerevisae. ChIP
(chromatin immunoprecipitation) experiments have revealed that
the initiation of transcription at cohesin-binding sites results
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K. M. Feeney, C. W. Wasson and J. L. Parish
in local disassociation of cohesin from the chromatin and that
cohesin subunits relocate from their initial loading sites to places
of convergent transcription [33,100]. These results suggest that as
the transcription machinery travels along the DNA and encounters
bound cohesin rings, the machinery is simply too large to pass
through the rings and therefore pushes them until transcriptional
termination. This phenomenon has been used to explain why
cohesin molecules in yeast accumulate at sites of convergent
transcription.
An active function for the accumulation of cohesin at sites
of convergent transcription was later discovered in S. pombe
[101]. This study suggests that the accumulation of cohesin rings
may act as a barrier between coding regions to aid efficient
transcriptional termination. Convergent genes in S. pombe have
been shown to generate overlapping transcripts in G1 due to
inefficient transcriptional termination, whereas in G2 , when
cohesin is present, overlapping transcripts are undetectable. In
support of this, loss of the HP1 (heterochromatin protein 1)
family member Swi6 (switching deficient 6), required for proper
targeting of cohesin to the centromeres, or cohesin itself in
G2 -phase, results in the presence of overlapping transcripts that are
normally absent at this stage of the cell cycle [101]. Transcription
termination is particularly important for genes arranged in a
convergent orientation because failure of polymerase II complexes
to terminate transcription results in inhibition of gene expression
due to collision of the elongating complexes [102].
Chromatin silencing
Heterochromatin regions are, for the most part, transcriptionally
silent and are believed to serve several functions from gene
regulation to the protection of the integrity of chromosomes.
These areas of DNA are transcriptionally silent because they
are inaccessible to DNA-modifying enzymes and therefore
transcription elongation is blocked [103]. In S. cerevisiae there
are three classes of silent domains: the silent mating-type loci,
HMR and HML, telomeres and rDNA (ribosomal DNA). The
HMR and HML silent domains contain cryptic copies of the
mating locus alleles MATa and MATα respectively. These genes
at HML and HMR are not expressed but serve as a repository
of genetic information, which upon transposition by the HO
endonuclease causes yeast to switch from one mating type to
another. The HMR and HML domains remain silent by the
action of flanking regulatory elements known as silencers [104].
rDNA consists of a tandem array of 9.1 kb units repeated 100 –
200 times on chromosome XII [105]. Such repeated DNA is
subject to recombination and excessive recombination of the
rDNA domain can be deleterious. Regulation of recombination
is thought to involve Sir2 (silencing protein 2) [106]. Four
SIR genes exist in yeast and were identified by mutations
that activate the silent mating-type genes [107]. Sir2, Sir3
and Sir4 are essential for silencing because of their role as
structural components of silenced chromatin [108]. Once these
SIR proteins are recruited to a silencer, co-operative interactions
enable them to spread throughout the silent locus, with Sir3 and
Sir4 binding to the deacetylated tails of histones H3 and H4
[109].
Cohesin-binding sites have been discovered at silent loci in
S. cerevisiae, including areas adjacent to telomeres and the
HMR locus [110]. Interestingly, cohesin enrichment has also
been reported at the LTR (long terminal repeat) of the Ty2
retrotransposon, which has been shown to contribute to boundary
element function in the HMR boundaries [110]. The accumulation
of cohesin at silent domains has led researchers to hypothesize
c The Authors Journal compilation c 2010 Biochemical Society
that cohesin plays an important function in chromatin silencing.
Indeed, there is accumulating evidence that cohesin plays a
positive role in the formation of silent chromatin in yeast. Initially,
it was shown that cohesin is recruited to the HMR locus in a Sirprotein-dependent manner [32]. This is in agreement with results
showing that Sir2 mediates the association of Scc1 with rDNA
[111]. The genes that encode rRNA are clustered in LTRs and
genes with such a repeated structure are, in general, thought
to be unstable because of a high frequency of recombination
events. Sir2 plays an important role in decreasing the frequency
of recombination of rDNA [106]. Interestingly, a decrease in
the association of the cohesin subunit Scc1 to rDNA has been
reported in Sir2 mutant strains [111]. Therefore it was proposed
that Sir2 prevents unequal sister-chromatid recombination,
probably by forming DNA structures with the aid of cohesin.
Silent chromatin-mediated recruitment of cohesin might serve a
similar function at the quiescent mating-type loci where postreplicative recombination between homologous sequences at
HMR, HML and MAT could lead to unprogrammed matingtype switching or lethal chromosomal deletions. In support
of this, cohesin mutant cells have shown such rearrangements
[112].
The idea that cohesin may stabilize silent chromatin and prevent
recombination has also been demonstrated in S. pombe [113].
The mating-type region contains three linked loci called MAT1,
MAT2 and MAT3, where MAT2 and MAT3 serve as donors
for MAT1 switching in a similar mechanism to the HMR and
HML loci in S. cerevisiae. It has been shown that a discrete
20 kb region containing MAT2 and MAT3 is assembled into a
heterochromatic structure and cohesin is recruited to the region
in a Swi6-dependent manner. Although cohesin does not seem to
affect Swi6 localization and silencing at the MAT locus, it has
been shown that cohesin mutants produce switching-defective
colonies at a rate nearly 100-fold that of the wild-type [113]. In
addition, defects in the recruitment of cohesin in Swi6 mutants
might be partly responsible for the increased rearrangement at
the MAT locus. Taken together, these results suggest that cohesin
is recruited to silent chromatin to regulate recombination and is
essential to prevent unwanted recombination at these unstable
regions (Figure 3).
Controversially, cohesin has also been shown to inhibit the
establishment of silent chromatin. A study by Lau et al. [114]
showed that silencing of the HMR domain is prevented by the
presence of Scc1 during G2 /M, even in the presence of
the Sir proteins. Cleavage or repression of Scc1 during the cell
cycle results in silencing of the HMR domain and inhibition of
Scc1 cleavage represses silencing [114]. In addition it has been
hypothesized that sequences near the border of silent and nonsilent chromatin domains would include a boundary element. To
test this, a number of target boundary elements were deleted to
determine whether neighbouring reporter genes became inactive
due of the spread of the silent chromatin from the HMR locus.
Of the numerous chromosome structure proteins tested, only
mutations in the cohesin subunits Smc1 and Smc3 were found to
have significant effects on boundary function [115]. Smc proteins
are reported to be part of the nuclear scaffold and are thought to
be involved in chromosome loop organization [116], thus linking
the insulator function of cohesin at the HMR with models in
which insulator element function is dependent on chromosome
architecture (Figures 4 and 5). The study by Donze et al. [115]
suggests that proteins involved in higher-order chromosome
structure might be involved in the functional delineation of
the chromosome and that the formation of large multiprotein
complexes at the boundary of silent chromatin prevents the spread
of heterochromatin (Figure 3).
Cohesin regulation and function
Figure 3
153
Model for the role of cohesin at silent domains
Top panel: Scc1 associates with the silent domain preventing the establishment of silencing until the creation of cohesin-associated boundary elements. This prevents the unwanted spread of silent
heterochromatin. Middle panel: following establishment of the boundary elements, Scc1 is cleaved and the spread of Sir proteins can occur. Bottom panel: once silent heterochromatin is developed,
cohesin (orange rings) re-associates to stabilize the domain, preventing unwanted recombination.
Figure 4
Structure of the β -globin locus
Looping of the β -globin locus is created by the interaction of CTCF (green) and cohesin (orange
rings), allowing for long-range spatial interactions between the LCR (dark blue) and active
genes. The association of cohesin rings at the CTCF-binding sites within the 5 and 3 DNase
I-hypersensitive regions creates as a barrier, preventing interaction of the β -globin locus and
regulatory elements of neighbouring genes.
Control of transcription
In mammalian cells cohesin is not enriched at sites of convergent
transcription but at DNase I-hypersensitive sites [117]. These
sites have also been shown to contain an over-representation of
the consensus sequence CTCF (CCCTC-binding transcriptional
insulator protein) [117,118]. CTCF is a ubiquitously expressed
protein that contains 11 zinc-finger motifs. Its expression is cellcycle regulated, peaking at the S- to G2 -phase [119]. Interestingly,
depletion of CTCF disrupts the specific positioning of the cohesin
subunits Rad21 (Scc1) and Smc3, but depletion of cohesin
does not affect the positioning of CTCF [117]. CTCF-binding
sites often flank groups of genes that are transcriptionally coregulated, suggesting that the majority of CTCF-binding sites
function as insulators [120]. Insulators are genetic elements
near the chromatin domain boundaries that are involved in the
alteration of gene expression. They act as barriers that shield
genes from position effects to prevent the spread of repressive
heterochromatin from one domain to the next [121]. In addition,
they can prevent communication between distant elements, such
as enhancers, to influence gene expression, a function known as
enhancer blocking [122].
There are a number of genes shown to be insulated by
CTCF. The first set of genes that CTCF was found to insulate
in an enhancer-blocking fashion was the β-globin gene locus
which contains a number of developmentally regulated erythroidspecific genes. CTCF was found at the 5 and 3 chromatin
boundaries of the chicken β-globin locus [123] and CTCFbinding sites have also been found to flank the equivalent
human and mouse gene loci [124]. Enhancer-blocking assays
showed that CTCF binding at the boundaries of the human and
mouse gene loci results in transcriptional insulation by preventing
inappropriate interactions between β-globin regulatory elements
and those of neighbouring domains [124]. There is also evidence
that CTCF mediates long-range chromatin looping in the βglobin locus (Figure 4) [125]. In mice, three CTCF-binding sites,
two upstream and one downstream of the locus, are involved
in spatial interactions between the LCR (locus control region)
and the active β-globin genes, forming an active chromatin
hub [126]. Disruption of CTCF binding at the downstreambinding site results in destabilization of the loop [125]. To
determine whether cohesin is important for the insulator function
of CTCF, cells were depleted of either CTCF or Rad21 protein
and transcriptional changes measured by DNA-chip technology.
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K. M. Feeney, C. W. Wasson and J. L. Parish
Figure 5 Arrangement of the IGF2 /H19 locus and structural rearrangements
resulting in maternal and paternal imprinting
(A) Situated between the IGF2 and H19 genes are three DMRs. Each gene is transcribed from
a unique promoter (arrows). (B) The maternal locus contains unmethylated DMR allowing
association of CTCF (green) and cohesin rings (orange rings) with these domains. This leads to
the formation of a loop between DMR1 and DMR, blocking the access of IGF2 to the enhancer
elements resulting in insulation of the gene. (C) The paternal locus contains methylated DMRs,
which results in abrogation of CTCF and cohesin binding. This allows interaction of DMR2 and
DMR resulting in enhancement of IGF2 gene expression from cis -acting enhancer elements.
Similar transcriptional changes were observed after downregulation of CTCF or Rad21, and genes within 25 kb of cohesinrich sites had a higher tendency to be up-regulated rather than
down-regulated, consistent with the conclusion that cohesin
mediates CTCF insulator function [118]. It has also been shown
that cohesin plays a role in CTCF insulator function by the
insertion of two copies of a known CTCF-dependent insulator
between a neomycin gene and an associated SV40 (simian
virus 40) enhancer and promoter elements. Depletion of CTCF
in cells containing this construct resulted in an enhancement
of neomycin expression and surprisingly a similar result was
obtained following depletion of Rad21 [117]. This shows that
cohesin is involved in CTCF-dependent transcription insulation
via the creation of a barrier at CTCF sites, where cohesin
association blocks the gene from associated enhancer elements.
The fact that it requires two of the CTCF-dependent insulator
sequences between the gene and enhancer/promoter elements to
generate insulation suggests that cohesin creates the insulation by
the creation of a loop structure within the DNA (Figure 4).
CTCF is also involved in imprinting of the IGF2 (insulinlike growth factor 2)/H19 (H19 fetal liver mRNA) mouse locus.
The IGF2 and H19 genes are located on chromosome 7 within
c The Authors Journal compilation c 2010 Biochemical Society
90 kb of each other. These two genes are reciprocally imprinted
in developing embryos with paternal expression of IGF2 and
maternal expression of H19 [127]. A set of enhancers downstream
of the H19 gene are involved in the regulation of expression of
both IGF2 and H19 expression. It is thought that CTCF insulates
this locus by binding to DMRs (differentially methylated regions)
thus insulating the maternal IGF2 locus [128] (Figure 5B). The
DMRs within the IGF2 locus bind to one another via interactions
with CTCF and other proteins, creating loops that are involved
in the insulation of the locus [129,130]. In support of this, the
association of CTCF with DNA has been shown to be methylationsensitive as methylation of DMR in the paternal locus prevents
CTCF binding leading to a bypass of the H19 locus and activation
of IGF2 expression (Figure 5C) [131]. To confirm these results,
the effect of methylation on the H19/IGF2 locus was explored by
Hcy (homocysteine) treatment, which induces hypomethylation
of the DNA. Hcy treatment resulted in the hypomethylation of
CTCF-binding sites upstream of the H19 locus and an increase
in H19 mRNA along with decreased expression of IGF2 mRNA
[132].
Cohesin is also involved in the CTCF-dependent insulation
of the IGF2/H19 gene locus [133]. Using methylation-sensitive
PCR, the cohesin subunit SA1 has been shown to associate with
the CTCF-binding sites within the IGF2/H19 gene locus in an
allele-specific manner. CTCF has previously been shown to be
involved in interchromosomal looping at the β-globin [125] and
IGF2/H19 loci [129]. Therefore it is likely that cohesin helps
to create these loops by creating a bridge via the handcuff
model discussed above (Figures 2, 4 and 5). The creation of
interchromosomal loops at the IGF2/H19 locus by cohesin and
CTCF has been confirmed using 3C (chromosome conformation
capture) assays and RNA interference depletion of cohesin
[134]. When cohesin was depleted, the higher-order chromatin
conformation at the locus was disrupted, which in turn disrupted
insulation of the IGF2 gene [134]. In addition, it has been
shown that cohesin is required for long-range CTCF-dependent
cis interactions at the developmentally regulated IFNG (interferon
γ ) cytokine locus [135]. Using 3C-based assays it was shown
that the IFGN locus contains topological loops and depletion
of either CTCF or cohesin resulted in the loss of these loops,
thus demonstrating that CTCF and cohesin are both important
for long-range interactions between DNA sites [135]. CTCF and
cohesin are also required for transcriptional insulation of the IL-3
(interleukin 3) and GM-CSF (granulocyte-macrophage colonystimulating-factor) gene cluster. These genes are responsive to the
same signals but are regulated independently by tissue-specific
enhancers. Situated between the two genes are CTCF-binding
sites that recruit both CTCF and cohesin to block interactions
between the various enhancer and promoter elements allowing
independent regulation of gene expression [136].
Interestingly, a similar CTCF-dependent role for cohesin is seen
in KSHV (Kaposi’s sarcoma-associated herpes virus). CTCFbinding sites exist within the major latency control region of
KSHV and cohesin co-localizes with CTCF within this region,
suggesting that CTCF and cohesin function as insulators in
the KSHV genome [137]. Depletion of CTCF results in a loss
of cohesin association and inappropriate expression of lytic
genes. Interestingly, the expression of genes regulated by CTCF
and cohesin is controlled in a cell-cycle-dependent manner and
mutation of CTCF-binding sites results in a disruption of phasedependent gene expression. This suggests that CTCF and the
cohesin complex together play a role in regulating the control
of cell-cycle-dependent gene expression during latency thus
preventing disruption of host cell proliferation [138]. A similar
role for CTCF in the prevention of inappropriate expression of
Cohesin regulation and function
lytic genes has been shown in HSV1 (herpes simplex virus type 1),
suggesting this may be a conserved mechanism that DNA viruses
use to control gene expression [139,140].
COHESIN AND DNA DAMAGE REPAIR
In recent years it has become apparent that cohesin plays
an important role in DNA damage repair. Eukaryotes have
evolved two mechanisms for DNA damage repair: NHEJ (nonhomologous end-joining) and HR (homologous recombination).
HR involves the use of a homologous template to repair the
damage. Typically the template is a sister chromatid, therefore
HR occurs from S- to G2 -phase. NHEJ entails the direct rejoining
of DNA ends and is therefore error-prone, unlike HR. It is
easy to envisage how cohesin can facilitate HR by keeping
the homologous template in close proximity to the damaged
DNA strand. Indeed, studies in yeast demonstrated that cohesion
established in S-phase is required for efficient DNA damage
repair in G2 -phase [141]. Furthermore, DNA damage induces
post-replicative cohesin loading in the region of a DSB (doublestrand break) [142,143]. This was surprising as it had been
shown previously that cohesin loading and cohesion establishment
occurred in late G1 - and S-phases respectively [38,144].
So what are the molecular components required for this
process? Perhaps not surprisingly, the cohesin loading protein
Scc2 is required for this damage-induced post-replicative
cohesin loading [142,143]. In addition, the primary checkpoint
kinases Mec1 (mitosis entry checkpoint 1) and Tel1 (telomere
maintenance 1) are required, but act in a redundant manner
[143]. Phosphorylation of histone H2AX by Mec1/Tel1 generates
what is known as γ -H2AX. The γ -H2AX signal extends at least
60 kb from the break, whereas cohesin extends 40 –50 kb from
the break. Strains expressing a non-phosphorylatable H2AX fail
to recruit cohesin thus suggesting that γ -H2AX may act as
a signal for cohesin assembly. Mre11 (meiotic recombination
11), a component of the DNA-damage-sensing complex MRX
[Mre11/Rad50/Xrs2 (X-ray-sensitive 2)], is also required for
cohesin assembly around the DSB but it does not abrogate
γ -H2AX formation, indicating that Mre11 is involved in an
independent pathway [143]. In addition to cohesin loading to the
site of a DSB, DNA damage also induces post-replicative cohesion
establishment [142,145] and it has been shown that DNA-damageinduced cohesion is established genome-wide and is not restricted
to the break site [146,147]. Curiously, damage-induced cohesion
requires Eco1 and Scc2 suggesting that newly loaded cohesin is
used to establish damage-induced cohesion.
In S. cerevisiae, mitotic and meiotic cohesin contains different
kleisin subunits, Scc1 and Rec8 (recombination 8) respectively.
Substitution of Scc1 with Rec8 in mitotic cells results in impaired
DSB repair [148]. Crucially Rec8 cohesin fails to become
cohesive after DNA damage. Further analysis revealed that Chk1
(checkpoint kinase 1)-mediated phosphorylation of Ser83 of Scc1
is a critical determinant for DSB-induced cohesion. DSB-induced
phosphorylation of Ser83 primes Scc1 for acetylation by Eco1
and antagonizes the anti-establishment activity of Wpl1 (see
above) [149]. S-phase cohesion, on the other hand, depends on
acetylation of Smc3. Thus Eco1-mediated acetylation of distinct
targets, depending on the cell cycle phase, facilitates cohesion
establishment by antagonizing Wpl1.
During an unperturbed cell cycle, the active site of separase is
blocked by the protein securin and the destruction of securin at
anaphase facilitates cohesin cleavage [69]. In response to DNA
damage the yeast homologue of securin, Pds1, is stabilized, which
presumably preserves cohesion until the cell-cycle checkpoint is
155
activated and/or the damage repaired [150]. Pds1 stabilization is
mediated by the cell-cycle checkpoint kinases Chk1 and Rad53.
Chk1-dependent phosphorylation prevents Pds1 ubiquitination by
the APC, whereas Rad53 prevents Pds1 from interacting with the
APC-associated protein Cdc20 [151]. It has also been suggested
that dephosphorylation of Pds1 occurs following repair of DNA
damage [151].
As mentioned previously, DNA damage induces cohesin
loading and cohesion establishment. These studies were
performed using S. cerevisiae in which a single DSB was
generated within the MAT locus on chromosome III by induction
of the HO endonuclease [143]. However, a recent study in
S. pombe found that an epitope-tagged version of Rad21 (Scc1)
was not loaded around the DSB site [152]. Whether this truly
reflects divergence in the DNA damage response between fission
and S. cerevisiae or if it reflects differences in assay conditions is
unclear. It will be interesting to see if DSB-induced cohesion
occurs in S. pombe. Adachi et al. [152] identified multiple
Rad21 phosphorylation sites in extracts from asynchronous
and mitotically arrested cells. One of these phosphorylation
sites, Ser553 , may have a role in the DNA damage response.
Following exposure to UV damage Ser553 was phosphorylated
in a Rad3 (which is homologous to Mec1)-dependent manner.
Phosphorylation of Ser553 was observed ∼ 2 h later than the
γ -H2AX signal, which suggests that phosphorylation of Ser553
may have a role in re-entry into the cell cycle after checkpoint
arrest or mitotic exit. Rad3-dependent phosphorylation of Rad21
following UV exposure had been demonstrated previously,
although the phosphorylation sites were not determined [153].
Although the signalling pathway may vary, Rad21/Mcd1
phosphorylation following DNA damage occurs in both fission
and S. cerevisiae. It remains to be seen if damage-induced Rad21
phosphorylation mediates Eco1 acetylation and subsequent
cohesion, as has been illustrated for S. cerevisiae [149].
In vertebrate cells there is evidence that cohesin is recruited
to DSBs. Remarkably, cohesin recruitment to DSBs is dependent
on another SMC complex, Smc5–Smc6 [154]. In S. cerevisiae
the Smc5–Smc6 complex has also been shown to co-localize
with cohesin on undamaged chromosomes [155]. Furthermore,
Smc5–Smc6 is required for DNA damage repair and has a
specific role in HR [156]. However, studies using fission and
S. cerevisiae present conflicting evidence on the requirement
of the Smc5–Smc6 complex for cohesin recruitment to DSBs
[146,157]. In S. cerevisiae Smc6 is required for damage-induced
genome-wide cohesion, but not for damage-induced cohesin
loading, whereas in S. pombe, cohesin is still recruited to
DSB regions in the absence of functional Smc6 [146,157]. The
interpretation of these studies is further complicated by the finding
that depletion of Smc5 [and Mms21 (mitochondrial splicing
system 21) a non-SMC subunit of the Smc5–Smc6 complex]
results in severe chromosome misalignments, whereas Smc6
depletion does not. This suggests that Smc5 function is not
always dependent on its interaction with Smc6 [158]. Further
study of this somewhat obscure Smc5–Smc6 complex will help
to clarify its role in DSB repair and cohesin dynamics (reviewed in
[159]).
There are other lines of evidence that highlight the role of
cohesin at DSBs. Cohesin recruitment to DNA damage was
illustrated by immunofluorescent staining of Smc1 and SA1 at
the site of laser-induced DNA damage [160]. However, in a
subsequent report cohesin accumulation could not be detected
following DNA damage induced by IR (ionizing radiation), but
was only detected when a high-energy laser was used [161].
The authors suggest that such a high laser intensity induces
local disruption of the nuclear structure and does not reflect
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K. M. Feeney, C. W. Wasson and J. L. Parish
classical IRIF (IR-induced foci). Indeed, when moderate IR
doses (3–10 Gy) were used, focus formation of DNA damage
checkpoint proteins [e.g. γ -H2AX, ATM (ataxia telangiectasia
mutated), BRCA1 (breast cancer early-onset 1) and 53BP1 (p53binding protein 1)] was observed, but not cohesin subunits
[161]. This seems to contradict the study by Potts et al. [154],
which demonstrated Smc5–Smc6-dependent cohesin recruitment
to DSBs in an ChIP assay; however, these conflicting results
may reflect a difference in sensitivity between the assays used.
A recent study has shown that cohesin subunits localize to IRinduced DSBs only during S- and G2 -phases and not at G1 [162].
This suggests an active role of the cohesin complex in the repair
of DSBs that may be unique to the G2 -phase of the cell cycle
and hence replicated chromatin [162]. That study may offer some
explanation for the discrepancies described above and the cellcycle-specific recruitment of cohesin to DSBs should be more
carefully studied.
Interestingly, it has been shown that Smc1 is phosphorylated
at Ser957 and Ser966 following IR and UV damage, mediated by
ATM and ATR (ATM- and Rad3-related) respectively [163–166].
Phosphorylated Smc1 localizes to γ -H2AX foci, which mirrors
the finding in S. cerevisiae that cohesin is loaded on to DSBs
in the same region as the γ -H2AX signal [143,161,167,168].
Phosphorylation of Smc3 also occurs following IR in an ATMdependent manner [169]. Mutations in Smc1 or Smc3 that prevent
phosphorylation result in abrogated DNA damage responses,
although the exact mechanism for this remains unclear [167,169].
It is tempting to speculate that ATM-dependent phosphorylation
of cohesin subunits generates sister chromatid cohesion; however,
evidence of DNA-damage-induced cohesion in mammalian cells
has been lacking. A recent study shows that inter-sister chromatid
distances were reduced following DSB induction [170]. Although
a direct role for cohesin could not be demonstrated, disruption
of the ATM locus resulted in inter-sister chromatid distances
similar to undamaged cells [170]. This implies that following
DNA damage, ATM phosphorylates cohesin leading to cohesion
establishment. Further research is required to test this, and
indeed whether the proposed DNA-damage-induced cohesion is
restricted to the site of the DSB.
Like Pds1, securin is also involved in the DNA damage
response. Securin interacts with p53 and represses p53
transcriptional activity [171]. Securin also interacts with the
NHEJ proteins Ku70 and Ku80 [172]. In contrast with yeast,
DNA damage induces degradation of human securin [173,174].
Given that securin degradation triggers release of separase and
subsequent cohesin cleavage it may seem surprising that securin is
degraded following DNA damage. However, securins are required
for separase activity and localization, suggesting that securin is
not merely an inhibitor of separase [175,176]. Indeed, securin has
a well established role in the pathogenesis of pituitary tumours.
The involvement of securin in tumour development is perhaps a
product of its roles in cohesin dynamics, DNA damage repair and
its more recently identified role as a transcription factor which
specifically regulates the G1 -to-S transition [177].
COHESIN AND HUMAN DISEASES
Two human diseases known as cohesinopathies are characterized
by mutations in cohesin subunits and cohesin establishment
factors: CdLS and Roberts syndrome. An interesting feature
of these diseases is that mutation of the cohesion complex
has a limited affect on sister chromatid cohesion and cell
cycling and therefore may be caused by changes in gene
expression. This hypothesis is supported by a study in
c The Authors Journal compilation c 2010 Biochemical Society
S. cerevisiae in which mutations were made that recapitulate the
mutations found in cohesinopathies. These mutations had little
effect on sister chromatid cohesion or cell viability but led to
defects in transcription control, gene silencing and sub-nuclear
organization of chromatin [178]. CdLS is caused by mutations
in either the NIPBL gene, which, as described above, encodes
the human orthologure of Nipped B/Scc2 [98], or in the Smc1or Smc3-encoding genes [99]. In Drosophila, Nipped B
has been shown to alleviate cohesin-mediated blocking
of enhancer–promoter communication by assisting in long-range
enhancer–promoter interactions. Nipped B and cohesin colocalize
to active transcription units and examination of cohesin-binding
sites has shown that cohesin preferentially binds to the promoter
region of genes [179]. Binding of cohesin to these sites is
significantly reduced in NIPBL-mutant CdLS patient samples
resulting in up-regulation of cohesin-associated genes [180].
Mapping of the Smc1 and Smc3 mutations found in a number
of CdLS patients on to the known structures of the SMC
hinge domain from Thermotoga maritima [181] and the S.
cerevisiae Smc1 head domain [182] has shown that the mutations
predominantly mapped to the coiled-coil and hinge domains of
the proteins [97]. These mutations are thought to either cause
angulation of the proteins disrupting the approximation of the
SMC head domains, or interfere with the binding of accessory
proteins to the cohesin ring, such as Scc2, which would abolish
loading of cohesin on to chromatin [97].
Roberts syndrome is caused by the mutations in the ESCO2
gene, the human homologue of S. cerevisiae Eco1 [183].
Cells derived from Roberts syndrome patients have been
shown to have defects in sister chromatid cohesion [184],
characterized by heterochromatin repulsion [185], suggesting
that Esco2 may be important in the establishment of cohesin
at heterochromatic regions. Interestingly, the genetic mutations
found in Roberts syndrome patient samples are predominantly
associated with loss of the acetyltransferase activity of Esco2
[186], providing evidence that this activity is indeed important
for cohesion establishment. Interestingly, replication forks in
Roberts syndrome cells are slowed, demonstrating that fork
progression is regulated by acetylation of cohesin and loss of
this mechanism results in the accumulation of DNA damage,
which could contribute to the abnormalities observed in Roberts
syndrome [55].
SUMMARY
For a number of years cohesin has been seen as the master
regulator of chromosome segregation. This tightly regulated
function of cohesin has been explored in depth but there are
still many unanswered questions as to how cohesin is loaded
on to and associates with DNA. Whether ATP hydrolysis is
required for hinge opening, Scc1 binding or Scc1 dissociation
needs to be definitively answered. In vitro single molecule studies
of fluorescently tagged proteins would enable determination of
whether ATP binding and hydrolysis occur simultaneously with
hinge opening and/or Scc1 dissociation. In addition, careful
analysis of the timing of ATP hydrolysis within the cell cycle
would help to determine whether hydrolysis is required for
loading on to chromatin prior to S-phase or for replicationcoupled cohesion establishment. Determining the stage at which
ATP hydrolysis occurs may also shed light on how cohesin holds
DNA strands together. If ATP hydrolysis is indeed required for
hinge opening and occurs during DNA replication, one could
hypothesize that this reflects ring opening as the replication fork
progresses and would imply sister chromatid cohesion by single
Cohesin regulation and function
cohesin rings rather than a higher-order complex as described in
the handcuff model. The molecular trigger for ATP hydrolysis
is also unclear. Determining the molecular events that lead
to ATP binding and hydrolysis will provide new insight into
the requirement for ATP in the multiple functions of cohesin.
Possible stimuli could include post-translational modification of
one of more cohesin complex subunits or interaction with other
proteins at specific stages of the cell cycle. Indeed interaction
with the cohesin loader Scc2/Nipped-B appears to promote ATP
hydrolysis, although the specific mechanism of action remains
unknown. It is important to determine whether Scc2 has inherent
enzyme activity or whether it functions in the cohesin loading
pathway simply by recruiting other proteins to the cohesin
complex. It would also be interesting to determine the timing
and localization of the Scc2 interaction with cohesin and whether
this only occurs during DNA replication.
Whether the handcuff or ring model, or both, accurately reflect
the mechanism by which chromatin is captured by cohesin
in vivo remains unresolved. FRET and cross-linking studies would
suggest the ring model to be most accurate, but such biochemical
assays could be misinterpreted in the context of the whole cell.
Studies showing self interaction of cohesin subunits suggest that
cohesin forms a higher-order complex and thus holds chromatids
together via the handcuff model. It is of course possible, and
highly likely, that both models are correct and that cohesin
can associate with chromatin in multiple ways depending on
chromatin structure and specific function (cohesion, transcription
control or DNA damage repair). The biochemical assays that have
been used to study the molecular architecture of cohesin rings
on DNA should be utilized to specifically isolate the structure
of cohesin complexes involved in transcriptional control and
gene silencing compared with cohesin rings that are involved
in sister chromatid cohesion. Any differences observed will help
to determine whether cohesin rings do indeed self-associate to
hold chromatin together and whether this is a feature of cohesin
specifically involved in sister chromatid cohesion. We hypothesize
that there are multiple forms of cohesin that each have their own
function and differ in their specific mechanism of association with
DNA.
In addition to sister chromatid cohesion, cohesin has a major
role in DNA damage repair and gene regulation. The mounting
evidence shows that cohesin accumulates at sites of damage
and mutation of cohesin subunits results in impairment of
repair. The majority of these studies have been performed in
yeast and work in mammalian cells has produced conflicting
results. This is probably due to differing complexity in the
pathways involved in DNA damage repair between higher and
lower eukaryotes because the cohesin proteins themselves appear
highly conserved between yeast and humans. Differences in
the mechanism of cohesin-mediated transcriptional insulation
in higher and lower eukaryotes have also been highlighted.
Whereas in lower eukaryotes cohesin is enriched at sites of
convergent transcription, in higher eukaryotes, cohesin appears
to be actively recruited to DNase I-hypersensitive sites by CTCF,
a protein that has no known homologue in lower eukaryotes.
The discovery that the association of cohesin with chromatin is
vitally important in the control of gene transcription has helped
explain the defects observed in cohesinopathies, such as CdLS
and Roberts syndrome. As these syndromes are characterized by
developmental defects, with little or no association with cancer
predisposition, it is hypothesized that the insulator function of
cohesin is significantly disturbed, contributing to the phenotypes
seen, whereas the impact on sister chromatid cohesion and
chromosome segregation is less severe. These newly emerging and
novel functions demonstrate that cohesin is not only essential for
157
chromosome segregation, but is the master regulator of multiple
aspects of genome integrity in all eukaryotes.
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Received 28 January 2010/12 March 2010; accepted 18 March 2010
Published on the Internet 13 May 2010, doi:10.1042/BJ20100151
c The Authors Journal compilation c 2010 Biochemical Society