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Cohesin: Its Roles
and Mechanisms
Kim Nasmyth1 and Christian H. Haering2
1
Department of Biochemistry, University of Oxford, Oxford OX1 3QU, United Kingdom;
email: [email protected]
2
European Molecular Biology Laboratory, 69115 Heidelberg, Germany;
email: [email protected]
Annu. Rev. Genet. 2009. 43:525–58
Key Words
First published online as a Review in Advance on
August 24, 2009
chromosome segregation, sister chromatid cohesion, mitosis, meiosis,
structural maintenance of chromosomes (Smc), ABC ATPase
The Annual Review of Genetics is online at
genet.annualreviews.org
This article’s doi:
10.1146/annurev-genet-102108-134233
c 2009 by Annual Reviews.
Copyright All rights reserved
0066-4197/09/1201-0525$20.00
Abstract
The cohesin complex is a major constituent of interphase and mitotic
chromosomes. Apart from its role in mediating sister chromatid cohesion, it is also important for DNA double-strand-break repair and
transcriptional control. The functions of cohesin are regulated by phosphorylation, acetylation, ATP hydrolysis, and site-specific proteolysis.
Recent evidence suggests that cohesin acts as a novel topological device
that traps chromosomal DNA within a large tripartite ring formed by
its core subunits.
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INTRODUCTION
The segregation of sister chromatids to opposite poles of the cell during mitosis is not only
one of the most dramatic events in the life of
cells but also one of the most important. Sister
DNAs produced during S phase must be segregated to opposite sides of the cell before it
divides. This is a remarkable feat, considering
that the uncoiled DNA of a single human cell
would stretch for 2 m and, as a random coil of
naked DNA, would occupy a volume of 2.6 ×
107 μm3 , which is several orders of magnitude
larger than the cells themselves. How do mitotic
cells organize their DNA into sausage-shaped
chromosomes only a few micrometers long?
How do they avoid tangling sister DNAs together, instead organizing them in largely separate domains (chromatids) lying side by side?
And how do identical chromatids know that
one sister has to move to one cell pole while
the other sister has to move to the opposite
pole, such that cytokinesis can eventually generate two genetically equal daughter cells? Key
to these processes is the physical connection of
sister DNAs, tenuous along chromosome arms
but more intimate at centromeres. This sister
chromatid cohesion is not merely pretty to look
at, it is essential for setting up the connections
between sister kinetochores and mitotic spindle
microtubules so that sisters are pulled in opposite directions. Loss of sister chromatid cohesion then triggers segregation of chromatids at
the metaphase-to-anaphase transition. A multisubunit protein complex named cohesin is
responsible for cohesion between sister chromatids. This review focuses on the molecular
mechanisms of cohesin function.
WHY IS SISTER CHROMATID
COHESION IMPORTANT?
One of the great mysteries of chromosome
segregation is how cells ensure that sister kinetochores attach to microtubules that extend to
opposite spindle poles, a process known as biorientation. It has long been supposed, but never
proven, that kinetochores form rather rigid
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polar structures, that sister chromatid cohesion
ensures that sister kinetochores are held back to
back with their microtubule-attaching surfaces
facing in opposite directions, and that attachment of one surface to stiff microtubules from
one pole would per se preclude attachment of
the sister to the same pole. As first recognized
by Nicklas (123, 124), back-to-back geometry
cannot explain how cells ensure that the maternal and paternal kinetochores of bivalent chromosomes attach to microtubules extending to
opposite poles during meiosis I. He suggested
that biorientation during meiosis I is achieved
instead by a process of error correction. More
specifically, kinetochore microtubule connections are unstable unless they generate tension,
which only occurs when maternal and paternal
kinetochores are pulled in opposite directions.
Meiotic sister chromatid cohesion is essential
not so much to hold kinetochores in the correct orientation but rather to resist the tendency
of microtubules to tear the bivalent apart and
thereby to create the tension necessary to stabilize correct attachments. The same principle
may also apply to mitotic cells.
Once biorientation has been achieved, destruction of sister chromatid cohesion, an
equally important process, must occur to initiate the separation of sister chromatids at
the metaphase-to-anaphase transition. Cohesion destruction is inhibited by “lagging” chromosomes, specifically those that have not yet
bioriented, in a process known as the spindle
assembly checkpoint (SAC) (121).
WHAT HOLDS SISTER
CHROMATIDS TOGETHER?
The first molecular explanation for sister chromatid cohesion was inspired by the realization
that DNA replication produces sister DNAs
that are wound around one another. It was proposed that decatenation mediated by Topoisomerase II (Topo II) is a regulated process that
is not completed until all chromosomes biorient (115). An opportunity to test this idea arose
with the remarkable discovery that addition
of a 150-bp centromere sequence to circular
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plasmid DNAs containing a replication origin
improves markedly the fidelity of their segregation in yeast cells (16). Gel electrophoresis demonstrated that minichromosomes within
cells that had replicated their DNA and arrested
in a mitotic state due to a spindle poison such
as nocodazole (which triggers the SAC) were
almost exclusively monomeric (84). In other
words, Topo II had already completed its task,
and decatenation had been fully achieved. The
conclusion was that something other than DNA
intertwining holds these particular sister DNAs
together and that this unknown mechanism
might be mediated by “one or more interesting proteins” (84, p. 1715).
Proteins Required for Sister
Chromatid Cohesion
The search for such proteins intensified with
the finding that destruction of sister chromatid
cohesion at the metaphase-to-anaphase transition is triggered by a ubiquitin protein ligase
named the anaphase-promoting complex or cyclosome (APC/C) (68, 76, 164). Genetic studies
revealed the first likely candidates (46, 108). We
know from these and subsequent studies that
sister chromatid cohesion requires the following components.
1. A core cohesin complex containing two
Smc proteins, Smc1 and Smc3, and two
non-Smc proteins, Scc1 (also known as
Mcd1 or Rad21) and Scc3 (known in
mammalian cells as SA1 and SA2). All
components of the cohesin complex are
essential for maintaining sister chromatid
cohesion in postreplicative yeast cells.
2. An accessory protein composed of HEAT
repeats named Pds5 that binds more
loosely to the cohesin complex but is
found at similar locations on the genome
as cohesin.
3. A separate complex containing the HEAT
repeat Scc2 protein (known as Nipbl in
mammalian cells) and the TPR repeat
Scc4 protein. This complex is essential for
cohesin’s association with chromosomes.
It is necessary to establish cohesion but
not to maintain it after DNA replication
has been completed.
4. An acetyl transferase named Eco1 (Esco1
and -2 in mammalian cells), which, like
the Scc2/4 complex, is needed for the establishment of cohesion during S phase
but not for its subsequent maintenance.
The vertebrate Sororin protein may also
fall into this category.
In addition to these proteins (Table 1),
which are essential for mitosis, many others are
necessary merely to improve the efficiency of
sister chromatid cohesion. Budding yeast mutants lacking these proteins are viable despite
having high rates of chromosome missegregation. Their patterns of “synthetic lethality”
have formed the basis for dividing many of the
nonessential cohesion proteins of Saccharomyces
cerevisiae into two groups (207). The first group
includes the putative DNA helicase Chl1, a
factor known as Ctf4 that forms a complex
with various DNA-replication proteins (e.g.,
DNA polymerase α, Mcm and GINS complexes, Cdc45, Mrc1, Tof1, and Csm3) (33), and
a complex formed between the Tof1 and Csm3
proteins that associates with replication forks
and regulates their response to being stalled
(132, 207). The second group includes Mrc1
(also associated with replication forks) (73) and
a Replication Factor C (RFC) complex, whose
Ctf1 subunit has been replaced by the Ctf1-like
protein Ctf18 and contains the additional subunits Dcc1 and Ctf8 (104). Most of these noncohesin proteins are thought to contribute to
sister chromatid cohesion during DNA replication. It was recently suggested that both ORC
(152) and condensin (90) have a role in maintaining cohesion in postreplicative cells. In the
case of ORC, the possibility that the loss of cohesion stems from defects during DNA replication has not been ruled out.
Dissolution of Sister
Chromatid Cohesion
The destruction of sister chromatid cohesion
takes place in two phases. The first commences
during prophase. Cohesin at centromeres is
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Table 1
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Genes encoding cohesin subunits and regulatory proteins
SMC
subunit
SMC
subunit
ECU04_0930
ECU09_1910
Cohesin complexes
Encephalitozoon cuniculi
Saccharomyces cerevisiae
Schizosaccharomyces pombe
Caenorhabditis elegans
Drosophila melanogaster
Xenopus laevis
Homo sapiens
Accesssory proteins
mit
mei
mit
mei
mit
mei
mit
mei
SMC1
SMC3
psm1 +
psm3 +
him-1
smc-3
SMC1
Cap
mit
smc1a
mei
smc1b
mit
SMC1A
mei
SMC1B
smc3
SMC3
Loading complex
Encephalitozoon cuniculi
kleisin
WHD subunit
HEAT repeat
subunit
ECU04_1370
ECU04_0870
MCD1 = SCC1
IRR1 = SCC3
REC8
rad21+
psc3 +
+
rec11+
rec8
scc-1 coh-1
scc-3
rec-8 coh-3
Rad21
SA
c(2)M
rad21
rec8
stag1 stag2
RAD21
STAG1 STAG2
REC8
STAG3
Cohesion establishment/release
?
SA-2
?
HEAT repeat
associated
PDS5
pds5 +
evl-14
pds5
pds5a pds5b
PDS5A PDS5B
Separase
Securin
ECU07_0350
?
Saccharomyces cerevisiae
SCC2
SCC4
ECO1
RAD61
ESP1
PDS1
Schizosaccharomyces pombe
mis4 +
ssl3 +
eso1 +
wpl1 +
cut1 +
cut2 +
Caenorhabditis elegans
pqn-85
mau-2
F08F8.4
wapl-1
sep-1
ify-1
Nipped-B
CG4203
san eco
wapl
Xenopus laevis
nipbl
kiaa0892
esco1
wapal
espl1
(LOC398156)
Homo sapiens
NIPBL
KIAA0892
ESCO1 ESCO2
WAPAL
ESPL1
PTTG1
Drosophila melanogaster
Sse
thr
pim
Gene names of mitosis (mit) or meiosis (mei) specific Smc, α-kleisin, or HEAT repeat subunits of cohesin complexes and
proteins that regulate cohesin’s association with chromosomes or establishment or release of cohesion (59, 120, 127, 131)
according to Saccharoymces Genome Database, S. pombe GeneDB, WormBase, Flybase, Xenbase, or HGNC.
spared this fate, which presumably contributes
to a characteristic feature of mitotic chromosomes, namely their central (centromeric) constriction. The second phase involves separase, a
highly conserved CD clan thiol protease, whose
cleavage of Scc1 removes cohesin from chromosomes and triggers (at least in yeast) sister
chromatid disjunction (130, 185, 187, 194).
HOW COHESIN CONNECTS
SISTER CHROMATIDS
The Structure of Cohesin
At the heart of the cohesin complex is a heterodimer formed between its Smc1 and Smc3
subunits. Each subunit, composed of a 50nm-long intramolecular antiparallel coiled coil,
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forms a rod-shaped protein with a globular “hinge” domain at one end and an ATP
nucleotide-binding domain (NBD) of the ABC
family at the other (50, 57, 109). Because of
the antiparallel nature of their coiled coils, each
Smc NBD is built from two halves, one from
C-terminal and the other from N-terminal
amino acids, whereas the hinge domain is encoded by amino acids from the center of the
polypeptide. Heterotypic interactions between
the hinge domains of Smc1 and Smc3 lead to the
formation of V-shaped Smc1/3 heterodimers
with an Smc1 NBD at the end of one coiledcoil arm and an Smc3 NBD at the end of the
other (Figure 1a). The interaction between
Smc1 and Smc3 hinge domains is tight, with
a low nanomolar KD (50). The crystal structure of a homodimeric bacterial hinge from
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Smc1
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b
Separase
N
C
Scc1
Scc3
ATP
hydrolysis
Smc3
ATP binding
c
K112/K113
Smc1
NBDs
N
C
Scc1
Scc3
Figure 1
Structure of the cohesin complex. (a) The α-kleisin subunit Scc1 connects the nucleotide-binding domains
(NBDs) situated at the apices of the V-shaped Smc1/3 heterodimer and recruits Scc3 to the complex. ATP
binding leads to NBD association, whereas ATP hydrolysis presumably drives NBD disengagement.
(b) Schematic of a crystal structure of the Smc hinge dimerization domain from Thermotoga maritima
(pdb 1GXL). The Smc protamers dimerize via two symmetric interfaces that are separated by a central
channel. (c) Schematic based on the crystal structure of the Smc1 NBD bound to the C-terminal WHD of
Scc1(pdb 1W1W). The second Smc1 NBD in the symmetric crystal structure was replaced by a homology
model of the Smc3 NBD. Two molecules of ATPγS are sandwiched between the NBD interfaces. Lysine
residues 112 and 113 in the Smc3 NBD are acetylated by the Eco1 acetyltransferase. Representations
generated with Pymol (DeLano Scientific).
Thermotoga maritima shows that two shallow
U-shaped hinge monomers interact to form a
twofold symmetric torus or ring with a shape
not unlike that of DNA polymerase clamps, albeit with a much smaller hole in the middle
(Figure 1b). The most remarkable feature of
the interaction is that the two monomers are
held together at equivalent but potentially independent surfaces. Each of the two contacts
can exist in the absence of the other because
the structure of a second crystal form reveals a
dimer in which one interaction surface is disturbed by crystal contacts and in which the
dimeric torroid is slightly twisted open. This
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bipartite nature of Smc hinge-hinge contacts
is curious but nevertheless highly conserved,
and it applies equally to eukaryotic Smc heterodimers, including cohesins Smc1/3. The 50nm-long antiparallel coiled coils that emerge
from cohesin’s hinge dimerization domains are
interrupted by a couple of short conserved domains that may contribute to the bent shape
of cohesin’s coiled-coil arms observed in rotary
shadowed electron micrographs of vertebrate
cohesin complexes (1).
Like other ABC-like NBDs, the Smc1/3
NBDs have a bilobed design wherein several
helices form a rigid helical domain (HD) that
is flexibly attached to a set of β sheets containing the nucleotide-binding Walker A and
B residues. The coiled coils of Smc proteins are
connected to the HD. The crystal structure of
the yeast Smc1 NBD complexed with a fragment of Scc1 (51) closely resembles the structures of Rad50 from Pyrococcus furiosus (61) and
an Smc protein from T. maritima (101). Like
Rad50 (61) and an ABC ATPase transporter
(156), the Smc1 NBD crystallizes as a dimer
in the presence of the slowly hydrolysable ATP
analog ATPγS (Figure 1c). The ABC signature
motif at the tip of one of the HD’s helices within
one Smc1 NBD interacts with the phosphates
of a nucleotide bound to a Walker A motif in its
partner. ATPγS is thereby sandwiched between
two NBD domains.
Within cohesin itself, ATP is presumably
sandwiched between Smc1 and Smc3 NBDs.
Indeed, Smc1/3 heterodimers cooperate in hydrolysing ATP when the Smc1 NBD is bound
by Scc1 (3). Hydrolysis (but not binding) of
ATP at either site is abolished by mutation of
highly conserved Walker B glutamate residues.
Cohesin complexes containing Smc1 or Smc3
proteins with these mutations assemble normally but cannot associate correctly with chromatin or build sister chromatid cohesion (2).
The precise function of ATP binding and/or
hydrolysis remains mysterious. One possibility is that ATP binding forces Smc1 and Smc3
NBDs together, whereas hydrolysis permits
their dissociation (Figure 1a), but it is currently unclear how association/dissociation per
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se of NBDs facilitates cohesin loading onto
chromatin.
The HD and coiled coils of Smc proteins
are directly connected to a short loop (the
Q loop), at one end of which is a highly
conserved glutamine residue that coordinates
the active-site magnesium ion in a manner
similar to that of other P-loop ATPases. This
glutamine is thought to couple the rotation of
HD domains to changes in ATP binding and
hydrolysis. Because of their proximity to the
coiled coils, changes in the rotation of Smc
HD domains may in turn alter the rotation or
angle of cohesin’s coiled coils as they emerge
from each NBD. The coiled-coil sequences
(especially those of Smc3) are remarkably
conserved for some distance as they emerge
from the NBDs, suggesting that they have very
specific properties. Indeed, the integrity of this
section of the coiled coil within Smc1 appears
essential because an insertion of five amino
acids inactivates cohesin (112).
A remarkable feature of cohesin’s structure
is that the NBDs of Smc3 and Smc1 are bound
tightly by Scc1’s N- and C-terminal domains,
respectively, creating a huge tripartite ring (50).
Both domains are found in members of the socalled kleisin family of proteins (147). This family includes (a) α-kleisins, orthologs of Scc1 and
its meiotic variant Rec8, (b) β-kleisins, equivalent subunits of condensin II, (c) γ-kleisins, subunits of condensin I, (d ) δ-kleisins, subunits of
Smc5/6 complexes, (e) ScpA-like proteins that
bind to bacterial Smc NBDs, and ( f ) MukF,
which binds to the bacterial Smc-like MukB
protein. The stable cross-linking of Smc1 and
Smc3 NBDs by α-kleisins is reminiscent of
the bacterial MalK ABC-like transporter whose
NBDs are also connected through a so-called
domain swap of their C-terminal domains.
A crystal structure of Smc1’s NBD bound to
Scc1’s C-terminal domain shows that the latter forms a winged helix domain (WHD) that
binds through extensive hydrophobic interactions to the two most C-terminal β strands of
the Smc1 NBD (51). This interaction alters the
structure of Smc1’s NBD in a manner that is
essential for ATP binding and hydrolysis (3).
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The three helices, followed by two β strands of
Scc1’s winged helix, correspond to the boundaries and secondary structure predictions for
the C-terminal domains of all kleisins (147). All
presumably bind to their Smc partners in a similar manner, and a recent analysis of the crystal
structure of MukF bound to MukB’s NBD confirms this hypothesis (205).
Interestingly, mutation of amino acids either
within the WHD or within the β strands of
Smc1 to which it binds abolishes all association
between Scc1 and Smc1/3 heterodimers in vivo
(51). This finding implies that the WHD-Smc1
interaction is responsible for recruiting Scc1 to
Smc1/3, and only after this has taken place can
the N-terminal domain of Scc1 bind to Smc3’s
NBD.
The interaction between Scc1’s conserved
N-terminal domain and Smc3’s NBD is less well
understood. It was first revealed by the finding
that a polypeptide encoding the N-terminal 115
amino acids of Scc1 copurifies with intact Smc3
protein (but not Smc1) when coexpressed in insect cells (44, 50), as does the N-terminal half of
Scc1’s meiotic counterpart Rec8 (44). Whether
this interaction occurs in vivo was tested by
analyzing the fate of cohesin after cleavage of
Scc1 by separase. The N-terminal Scc1 fragment produced by separase cleavage remains associated with the C-terminal fragment because
both are attached to Smc1 and Smc3 NBDs,
which are themselves connected by their associated hinge domains. Meanwhile, simultaneous
cleavage of Scc1 and Smc3’s coiled coil releases
Scc1’s N-terminal cleavage fragment associated
with Smc3’s NBD (44). Additional evidence for
the notion that Scc1’s N-terminal domain binds
to the Smc3 NBD is the finding that there is
greater fluorescence resonance energy transfer
(FRET) in vivo between cyan fluorescent protein (CFP) fused to the N terminus of Scc1 and
yellow fluorescent protein (YFP) fused to the C
terminus of Smc3 than to YFP fused to the C
terminus of Smc1 (106).
The functional importance of the interaction has been tested by mutating residues within
Scc1’s conserved N-terminal domain, which is
predicted to contain three helical sections and
to fold into a helix-turn-helix domain. Mutations in several residues within the third helical section abolish copurification of N- and
C-terminal Scc1 fragments after cleavage of
Scc1 with tobacco etch virus (TEV) protease,
implying that these mutations affect the binding
of Scc1’s N-terminal domain to Smc3 (3). The
same mutations abolish cohesin function, but in
at least one case (D92K) lethality is rescued by
fusing Scc1’s N terminus to the C terminus of
Smc3 but not by fusing its C terminus to the N
terminus of Smc1 (43). The dependency of the
interaction on prior binding of the Scc1’s C terminus to the Smc1 NBD ensures that separate
Scc1 molecules cannot bind to the two NBDs of
a single Smc1/3 heterodimer, which would preclude ring formation. The recent finding that
Eco1 regulates the establishment of cohesion
by acetylating of Smc3’s NBD (see section Establishment of Cohesion) emphasizes the importance of understanding how Scc1 binds to
Smc3 NBDs. It will also be important to ascertain whether or not the association is in any
way altered when Smc1 and Smc3 NBDs engage each other in the presence of ATP, as has
been suggested for the MukBEF complex (205).
The picture emerging from these biochemical
experiments, namely a gigantic tripartite ring
complex containing Smc1, Smc3, and Scc1, is
consistent with electron micrographs of cohesin
complexes (1).
Little is known about Scc3, whereas Pds5 is
predicted to be composed almost exclusively of
HEAT repeats (122, 128). The latter’s association with the tripartite ring is clearly weaker
and more salt sensitive than that of Scc3 (106,
166). Nevertheless, association of both proteins with the Smc1/3 subunits depends on
Scc1 (50, 106). Scc3 binds directly to amino
acid residues within Scc1’s C-terminal separase
cleavage fragment that are N-terminal to the
conserved C-terminal WHD. Scc3 also binds
to Rad61 (yeast Wapl), at least in vitro (143).
Whether Pds5 binds directly to Scc1 or only via
an intermediary like Rad61 is not known (143).
Pds5 forms a more stable subcomplex with
Rad61/Wapl (88, 143). Because Rad61 is not
essential for sister chromatid cohesion, whereas
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Pds5 is, the latter must presumably bind cohesin
independent of Rad61/Wapl. Strangely, in vivo
FRET failed to detect proximity of Pds5 to Scc1
(or Scc3) but did detect a weak signal with the
Smc1/3 hinge. However, it has not yet been
possible to measure a physical interaction between Pds5 and isolated hinge complexes (106).
The Ring Model
The discovery that Smc1, Smc3, and the
α-kleisin Scc1 form a ring has led to the proposal that cohesin associates with chromosomes
by trapping DNA/chromatin fibers. If so, cohesin could hold sister DNAs together by trapping them both inside the same ring (50). Ring
(or embrace) models of this nature come in
two flavors. The strong version holds that sister
DNAs are trapped inside a single monomeric
cohesin ring (Figure 2a), whereas weak versions (sometimes referred to as handcuff models) postulate that sister DNAs are held together
by interactions between two different rings, one
that has trapped one DNA and a second that has
trapped its sister (Figure 2b,c).
Both weak and strong ring models hold
that cohesin grasps chromatin using a topological principle rather than physically binding to
DNA or nucleosomes. Accordingly, both types
of model predict that breaking the ring at any
a
b
Strong ring model
point should trigger cohesin’s dissociation from
chromatin and loss of sister chromatid cohesion. This hypothesis has been tested through
use of TEV protease to cleave open cohesin
rings. Cleavage of α-kleisin at TEV sites either at a mutated separase site (187) or elsewhere within its central domain (44, 130) does
indeed have this effect. Importantly, if the TEV
sites are flanked by sequences encoding MP1
on one side and p14 on the other—two protein domains that associate with each other with
a low nanomolar KD and a low off rate—then
α-kleisin cleavage no longer affects sister chromatid cohesion (43). This finding implies that
it is not the generation of novel N or C termini that compromises cohesin’s ability to hold
sisters together but rather the disconnection of
N- and C-terminal domains of its α-kleisin subunit. Severing the coiled coil of Smc3 has been
achieved through insertion of TEV cleavage
sites within regions of low coiled-coil probability on both strands at positions that coincide
within its coiled-coil arm (44). Cleavage of only
one strand has little or no effect on cohesin’s
activity, but simultaneous cleavage triggers cohesin’s dissociation from chromosomes (44, 69).
Yet another way of opening cohesin rings that
have previously associated with chromatin is to
use mutations that weaken one of their three intersubunit interactions. The S525N mutation
c
Weak ring models
Figure 2
Sister chromatid cohesion by cohesin rings. (a) The strong version of the ring model envisages that sister
chromatids (displayed as 10-nm fibers, e.g., DNA wrapped around nucleosomes) are entrapped within a
single cohesin ring. (b) One version of a weak ring model (the handcuff model) envisages association of (by
binding to a single Scc3 subunit) two tripartite Smc1/3/Scc1 rings. (c) Another version of the weak ring model
proposes the topological interconnection of two cohesin rings, each with a single sister chromatid entrapped.
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within Scc1’s C-terminal WHD causes its interaction with Smc1’s NBD to be sensitive to
high temperature. If cells with this mutation
are allowed to undergo DNA replication at the
low temperature and are then arrested in M
phase, subsequent incubation at the restrictive
temperature destroys sister chromatid cohesion
(14). In addition, mutations disrupting either
hinge interface weaken Smc1/3 dimerization
and greatly reduce the stability of cohesin’s association with pericentric chromatin in yeast,
with lethal consequences for sister chromatid
cohesion (A. Mishra & K. Nasmyth, personal
communication).
Both weak and strong ring models predict
that once DNAs are trapped inside cohesin
rings it would be impossible to substitute
ring components after cohesion has been
established without DNA escaping and sister
chromatid cohesion being lost. Investigators
intially addressed this prediction by expressing
in postreplicative cells an α-kleisin subunit that
cannot be cleaved by separase. If the noncleavable subunit could exchange with the wild-type
protein, then it should hinder destruction of
sister chromatid cohesion when cells activate
separase at the onset of anaphase. This does not
occur. Noncleavable α-kleisin only blocks sister
chromatid disjunction when it is expressed prior
to DNA replication (51). Likewise, expression
of wild-type ring components postreplicatively
in cells with temperature-sensitive alleles cannot rescue their loss of cohesion upon transfer
to the restrictive temperature (93, 161, 163,
190), at least in the absence of DNA damage.
This is true for Scc1 and Smc1, which are core
ring components, but not for Scc3, which is
not part of the tripartite ring (M. Brunet & K.
Nasmyth, unpublished observations).
If association between cohesin and chromatin is primarily topological, then cohesin
rings might be free to slide along chromatin
fibers when these are not stably packaged into
higher order structures. Cohesin rings have a
diameter between 30 to 35 nm, which is considerably larger than an extended 10-nm nucleosomal chromatin fiber. This finding led to the important prediction that the association between
cohesin and a small circular minichromosome
should be broken by cleavage not only of the
cohesin ring but also of the DNA. Cohesin associated with purified minichromosomes can be
detected by immunoprecipitation of minichromosome DNA through use of antibodies specific for epitope tags attached to cohesin. As
predicted by both types of ring model, coprecipitation of minichromosome DNA with cohesin is abolished either by cleaving the cohesin
ring with TEV protease or by linearizing the
DNA with a restriction enzyme (69).
A key step has been the development of
methods to detect sister minichromosome cohesion in vitro (70). Extracts are first fractionated by differential sedimentation velocity on sucrose gradients and are subsequently
electrophoresed under native conditions in
agarose gels. G1 cells contain a single species of
minichromosome DNA, namely one that sediments slowly in gradients but migrates rapidly
in agarose gels. G2- or M-phase cells, however, contain an additional higher-molecularweight (dimeric) form that sediments more
rapidly and electrophoreses more slowly. Crucially, dimeric minichromosomes are converted
to the monomeric form by cleavage of Scc1
by TEV protease. As predicted by both strong
and weak ring models, linearization of cohesed dimeric minichromosomes converts their
electrophoretic mobility to that of linearized
monomers.
The notion that cohesin holds sister
minichromosomes together via a purely topological mechanism makes a strong prediction,
namely that if the three subunit interactions
that create the cohesin ring were cross-linked,
then sister DNAs would be trapped inside
a covalently circularized cohesin ring. Under these circumstances, they would migrate
as dimers during gel electrophoresis even after protein denaturation with sodium dodecyl
sulfate (SDS). Researchers have achieved chemical circularization by fusing Smc3’s C terminus to Scc1’s N terminus and by introducing at the other two ring interfaces pairs of
cysteine residues that can be cross-linked by
the thiol-reactive cross-linkers dibromobimane
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(bBBr) and bis-maleimidoethane (BMOE) (49).
Either bBBr or BMOE induces cross-linking
at both interfaces in ∼30% of cohesin rings,
which causes ∼30% of the DNAs from “native”
dimers to form SDS-resistant dimers that migrate slightly more slowly than do DNA-DNA
concatemers. These dimers are actually composed of monomeric DNAs that are held together by cohesin in a manner dependent on
all four cysteine insertions and cross-linking
reagents but that are sensitive to proteolytic
cleavage by TEV protease at a unique site
within the polypeptide interconnecting Smc3
and Scc1.
Given the specificity of the cross-linking by
bBBr and BMOE, there is no reason to suppose that putative interactions between different cohesin rings will have been cross-linked in
these experiments, and they therefore exclude
the possibility that the connection between sister DNAs is mediated by nontopological interactions between cohesin complexes associated with each sister (111). The efficiency of
SDS-resistant dimer formation and its resistance to site specific proteolysis when created
from diploids expressing cleavable and noncleavable cohesin rings are consistent with the
notion that sister DNAs are entrapped by a single monomeric ring (49). Double-ring models
involving either a pair of concatenated rings,
each holding a separate sister DNA (Figure 2c),
or double-sized rings (formed by interconnecting two subunits each of Smc1, Smc3, and Scc1)
are hard to reconcile with the finding that the
fraction of DNAs dimerized is almost identical
to the fraction of cohesin rings circularized and
not to the square of this fraction. Nevertheless, ultimate proof that a single cohesin ring
holds sister DNAs together may require direct
quantification of the number of cohesin rings
associated with dimeric minichromosomes.
If trapping of DNAs inside cohesin rings
forms the basis for sister chromatid cohesion of
real chromosomes as well as circular minichromosomes, then it is clearly essential to explain how this process occurs. There are three
possibilities: (a) Either the ring is assembled
de novo around DNAs, or (b) DNA itself is
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transiently broken and one arm of a preassembled ring passed through the break before it
is resealed, or (c) the ring itself must be transiently opened. Assembly of the ring around
DNA does not seem likely for a number of
reasons. First, cohesin complexes not associated with chromatin also exist as a ring (44).
These include soluble wild-type complexes as
well as complexes with mutations in Smc1 or
Smc3 Walker B motifs that compromise ATP
hydrolysis and cannot therefore associate stably
with chromosomes (2). Second, most cohesin in
plant, animal, and fungal cells other than yeast
dissociates from chromosomes during prophase
and exists as a soluble complex during mitosis until it reassociates with chromosomes during telophase (52, 98, 167). Third, the cohesin
subunits stored in eggs that are used to establish sister chromatid cohesion during cleavage
divisions are clearly present as preassembled cohesin complexes (98, 100) that have the appearance of monomeric rings under the electron microscope (1). The DNA-breakage model seems
inherently implausible. This leaves us with the
idea that one of the three interfaces between the
tripartite ring subunits is opened in a regulated
fashion and constitutes an entry gate.
Cohesin’s entry gate cannot be located at the
interface between Smc1/3 NBD domains and
α-kleisin, because fusion of α-kleisin’s N- or Cterminal domains to their cognate NBDs does
not prevent establishment of sister chromatid
cohesion (43). To test whether the entry gate
is instead situated at cohesin’s hinge, Smc1’s
and Smc3’s hinge domains were replaced by
p14 and MP1, respectively, which (like Smc
hinge domains) form tight pseudosymmetric
heterodimers and whose N and C termini are
close enough to each other to permit formation
of Smc coiled coils. Despite forming tripartite
rings capable of ATP hydrolysis, these hingesubstituted Smc1/3 heterodimers neither associate with chromatin nor build sister chromatid
cohesion (43). Cohesin’s hinge must therefore have an essential function in addition to
dimerization.
It has proved possible to insert at short loops
within Smc1’s and Smc3’s hinges amino acid
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sequences encoding the human proteins
FKBP12 and FRB, which bind to each other
with a low nanomolar KD only when FKBP12
binds rapamycin. Remarkably, cohesin complexes with FKBP12 inserted into Smc3’s hinge
and FRB inserted into Smc1’s hinge form sister chromatid cohesion in the absence but not
in the presence of rapamycin (43). Rapamycin
blocks establishment of cohesion when added
before S phase but has no effect when added
immediately afterward (43). These findings are
consistent with the notion that the hinge must
be opened transiently during cohesion establishment. However, this hypothesis is by no
means proven, as clamping together of Smc1
and Smc3 hinge domains may abolish cohesion
establishment for reasons other than preventing hinge dissociation.
Smc1 and Smc3 hinge domains associate
with a low nanomolar KD , and ATP binding
and/or hydrolysis may be required to dissociate them. The conundrum is that these NBDs
are separated from the hinge by a 50-nm-long
coiled coil. This finding has led to the suggestion that the hinge is brought into direct proximity to the NBDs by folding the coiled coil
separating them.
Although passage of DNA double helices
through opened hinges may seem the obvious
route, it is not the only mechanism capable
of trapping DNA. DNA trapping could also
arise by passage through the hinge of a section of the cohesin ring itself (A. Leung, personal communication). Consider, for example,
the following scenario. If the DNAs destined
to be trapped were to lie orthogonally across
the parallel coiled-coil arms of an extended cohesin ring and the coiled coils were now folded,
bringing the hinge into contact with the NBDs,
the DNAs would be trapped between sections
of cohesin’s coiled coils, albeit in a temporary embrace. If the hinge were to open, the
NBDs were to disengage, and a central section
of Scc1 α-kleisin polypeptide connecting the
NBDs were to pass through the opened hinge,
then subsequent hinge closure would create a
twisted cohesin ring with the DNA trapped inside the ring. Untwisting the ring, which would
be possible because the Scc1 linker is a single
polypeptide, would transfer the change in linking number from the cohesin ring to the DNA,
which would be wound around the ring. If both
sister DNAs were to lie side by side on cohesin’s
coiled-coil arms at the initial step, then both
could be trapped inside the ring by this process.
Arguments Against the Ring Model
A criticism of models postulating that cohesin’s
association with chromatin is primarily topological is that they cannot explain the observation that cohesin is localized at specific
sequences along the genome (65, 111). Although it is certainly true that DNA entrapment does not per se provide an explanation
for cohesin’s localization, entrapment does not
preclude sequence-specific localization. For example, rings that have trapped DNA may diffuse laterally along chromatin fibers until they
meet site-specific DNA-binding proteins (such
as CTCF; see section Does Cohesin Mediate Nonsister Connections Important for Transcriptional Control?) that bind to them with low
affinity. Alternatively, localization could stem
from the positioning of cohesin’s Scc2/4 loading factor at precise loci.
Some criticism has been directed specifically
at the strong ring model. There are a number
of circumstances in which cohesin persists on
chromosomes even when sister chromatid cohesion has been lost, and this is thought to be
inconsistent with the notion that cohesin holds
sister DNAs together by entrapping them inside a single ring. One example is cohesin’s
behavior in eco1 mutants, where cohesin still
loads onto chromosomes but where sister chromatid cohesion is not established during DNA
replication (112, 155, 180). A second example
is the observation that sister chromatid cohesion of circular silent-mating type (HMR) loci
looped out from yeast chromosomes by sitespecific recombination is lost when silencing
factors are inactivated, and yet cohesin persists
on the chromatin circles (12). These phenomena undoubtedly require explanations, but they
are not necessarily inconsistent with the ring
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model. For example, Eco1 may be required to
entrap both sister DNAs inside a single ring but
not for entrapment per se.
Other Models: The Handcuff Model
It has been postulated that cohesion is mediated by interactions among separate cohesin rings associated with each sister DNA
(111). Weak ring models fall into this category
(Figure 1b). Is there any evidence for intercohesin complex interactions? In yeast, where
it is easy to make diploid strains heterozygous
for differently tagged cohesin subunit alleles
(each identically expressed at physiological levels from native promoters), little if any coprecipitation is observed either with soluble cohesin complexes or with complexes released
from chromatin by nuclease digestion (50). Indeed, the former and latter have been shown to
sediment similarly with velocities expected for
monomeric complexes (199). Likewise, there
appears to be little or no FRET between YFPand CFP-tagged versions of the same Smc subunits in vivo in yeast (106). However, if cohesin complexes that actually are engaged in
holding sisters together were rare, then they
may have been missed by the above studies.
More pertinent, therefore, is the finding that
chemical/translational cross-linking of the cohesin ring’s three interfaces creates circularized rings that hold sister minichromosomes
together even after denaturation in 1% SDS at
high temperatures (49). This result should not
be possible if nontopological interactions between different cohesin rings were an integral
part of the mechanism by which sister DNAs are
held together, because such interactions should
be destroyed by SDS treatment. It appears that
the only way of salvaging nontopological weak
ring models would be to propose that minichromosome cohesion uses an atypical mechanism,
which does not seem to be the case (143).
Soluble vertebrate cohesin complexes also
sediment with velocities expected of monomers
(52, 100, 166). In contrast, it has recently been
reported that differently tagged α-kleisin subunits coprecipitate, as do differently tagged
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Smc1 or Smc3 subunits (212). A fundamental problem with these experiments is that the
proteins are overexpressed, which is known to
facilitate interactions between separate Smc1/3
heterodimers. For example, separate yeast
Smc1/3 heterodimers can be interconnected by
Scc1 when overexpressed in insect cells but not
when expressed at physiological levels in yeast
cells (50). Conclusions based on overexpression studies should therefore be treated with
great caution. Is there any evidence for interactions between identical cohesin subunits expressed at physiological levels? Interestingly,
mammalian tissue culture cells provide a natural way of testing this hypothesis because they
express two variants of the Scc3 subunit, known
as SA1 and SA2. There is a consensus that these
two proteins cannot be coimmunoprecipitated
with each other (52, 100, 166, 167, 212). Does
this mean that cohesin does not form multimers? Not necessarily: It has been suggested
that the α-kleisin subunits of two different tripartite cohesin rings are connected by a single
Scc3 molecule (Figure 2b) (212).
Do α-kleisin subunits associate with each
other without overproduction? In extracts from
stable cell lines expressing moderate levels of
the Scc1 α-kleisin subunit tagged at its C terminus with myc epitopes, immunoprecipitation of
the myc-tagged protein does not coprecipitate
wild-type protein (194, 212). The argument
that the C-terminal myc tag interferes with
an all important kleisin-kleisin interaction is
hard to reconcile with the finding that the very
same protein can sustain mouse development
(K. Tachibana & K. Nasmyth, personal communication). The claim that different results
are obtained in extracts from stable cell lines expressing an N-terminally tagged α-kleisin subunit (212) also do not withstand careful scrutiny.
Although it is expressed at a lower level than the
endogenous protein, N-terminally tagged αkleisin coprecipitates only very modest amounts
of wild-type protein—indeed, no more than is
immunoprecipitated from extracts from cells
not expressing a myc-tagged form (212). In conclusion, there is currently little if any firm evidence that multimeric cohesin rings are formed
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under physiological conditions, which does not
necessarily mean they do not exist.
Other Models: Bracelets
Another suggestion is that cohesin rings are
merely a storage form for soluble cohesin
(65) and that sister chromatids are in fact
held together by cohesin complexes that have
oligomerized. It is argued that the known interactions between cohesin ring subunits could be
used to form not only rings but also oligomeric
filaments or “bracelets” that somehow wind
around sister chromatids (65, 121). This model
postulates oligomeric cohesin complexes that
have hitherto defied detection and cannot
explain the chemical cross-linking experiments
with minichromosomes (49). Despite this
drawback, the notion that Smc proteins may
form and function as multimeric filamentous
structures has received a boost from the
suggestion that MukB NBDs cannot engage
one another while both are associated with
MukF C-terminal domains (205). It has been
proposed that NBD engagement causes release
of one end of symmetrical MukE4 F2 hexamers
from one of the two NBDs of MukB dimers
and that the freed C-terminal MukF domain
can then interact with another MukB dimer in
a process that could be reiterated to form filamentous structures. Whether this really occurs
under physiological circumstances is unclear.
THE COHESIN RING CYCLE
Loading onto Chromatin
In animal and probably most fungal and plant
cells, 90% or more of the cohesin bound
to chromosomes dissociates during prophase
(131). This process generates a large soluble
pool of mitotic cohesin complexes that are
spared separase cleavage and reassociate with
chromatin during telophase. Loading of cohesin onto chromosomes is not confined to
postmitotic or G1 cells. This is true not only
in yeast, where α-kleisin expressed after S
phase associates with chromosomes (126, 186),
but also in HeLa cells, where cell lines stably expressing green fluorescent protein (GFP)tagged cohesin subunits have enabled measurement of chromosome residence times through
use of inverse fluorescence recovery after photobleaching (iFRAP). Such studies have revealed that most chromosomal cohesin has a
mean residence time of less than 25 min in
both G1 and G2 cells (35). G2 but not G1 cells
possess another population of cohesin (corresponding to one-third of the total), whose residence time is much longer and which may
correspond to the cohesin pool actually engaged in holding sister chromatids together.
Significantly, a large fraction of chromosomal
cohesin cycles on and off chromatin several
times during the cell cycle. Cohesin engaged
in holding sister centromeres together during mitosis turns over slowly if at all in yeast
(209), consistent with the findings that noncleavable α-kleisin subunits cannot form cohesion when expressed after DNA replication (51)
and that wild-type Smc1/3 or α-kleisin subunits
expressed in G2 cannot substitute temperature
sensitive variants (93, 163, 182).
Loading requires ATPase activity associated
with Smc NBDs. Mutant Smc1 or Smc3
proteins that can bind but not hydrolyze ATP
due to Walker B mutations form cohesin rings,
but these fail to associate stably with chromosomes (2, 3). The finding that engagement of
MukB’s NBDs is incompatible with both of
them binding to the kleisin-like MukF protein
(205) raises questions as to whether NBD
engagement causes displacement of either the
N- or the C-terminal domain of cohesin’s
α-kleisin subunit and, if so, whether this
process is essential for stable association with
chromatin. Pertinent to this issue is the fact that
cohesin complexes defective in hydrolyzing
ATP associated with either Smc1’s or Smc3’s
NBD can be isolated as stable rings, implying
that neither end of the α-kleisin subunit has
disengaged from the NBDs. In addition, full
disconnection of either NBD from its α-kleisin
partner is not necessary for establishing sister
chromatid cohesion, as yeast cells remain viable
when Smc3’s C terminus is fused to α-kleisin’s
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N terminus or when α-kleisin’s C terminus is
fused to Smc1’s N terminus (43). It is therefore
possible but not proven that cohesin’s NBDs
retain their association with both N- and
C-terminal α-kleisin domains throughout the
process of loading onto chromatin. Loading
also involves cohesin’s hinge domain, which
either acts as an entry gate (43) or conceivably enables initial contact with chromatin as
envisioned for Bacillus subtilis Smc proteins (58).
Importantly, cohesin’s association with
chromosomes requires several factors, some required at all positions on the genome and others
only at specific loci. The most important factor
is the Scc2/4 complex. Association of cohesin
with chromatin at all sites within the genome
depends on this complex in Saccharomyces cerevisiae (14, 180), in Schizosaccharomyces pombe (7,
178), in Xenopus extracts (36, 171), and in mammalian cells (150, 197). As might therefore be
expected, mutations in Scc2 (110) or Scc4 (14)
cause major defects in sister chromatid cohesion
(110). It has been suggested that Mis4, Scc2’s
S. pombe ortholog, may be part of an independent cohesion apparatus known as adherin
(32). This possibility is unlikely because neither Scc2 nor Scc4 is required to maintain sister
chromatid cohesion in postreplicative cells (7,
14). Nevertheless, loading cohesin onto chromosomes may not be the sole function of the
Scc2/4 complex, as yeast scc2 mutants also have
partial defects in the loading onto chromosomes of Smc5/6 complexes (94) and condensin
(17)—defects that cannot be attributable to
lack of cohesin loading. Scc2 is a large protein containing multiple HEAT repeats within
its C-terminal domain (122, 128) and an Scc4binding N-terminal extension whose length
varies enormously between species. Meanwhile,
Scc4 is composed of TPR repeats (197). The
complex is highly conserved and is sometimes
referred to as the NIPBL/Mau2 complex in humans (150).
In light of the cohesin-ring model, the
main function of the Scc2/4 complex may be
to promote entrapment of chromatin fibers by
cohesin. How then might Scc2/4 perform this?
It may act directly in the entrapment process,
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helping to open the ring and/or bringing cohesin into close proximity to chromatin fibers,
or it may act only indirectly, modifying chromatin fibers in a way that subsequently enables
cohesin to entrap them. If Scc2/4 acts on cohesin directly, the complexes must at some stage
interact physically with each other. Is there any
indication that this is the case? Scc2 and Scc4
form a tight stoichiometric complex that for
the most part does not copurify with cohesin
subunits (7, 14, 197). However, small amounts
of cohesin have been identified via mass spectrometry in association with purified Scc2/4
complexes (2), and Western blotting of cohesin
in Scc2/4 immunoprecipitates or vice versa is
consistent with this finding (7, 170, 180).
There is accumulating evidence that cohesin’s association with chromosomes requires
other factors in addition to the Scc2/4 complex. The best example has been discovered in
extracts from Xenopus eggs, where association
with chromatin of the Scc2/4 complex as well
as cohesin is dependent on the formation of
prereplication complexes (pre-RCs) (36, 171).
Most Scc2/4 in these extracts is associated with
the Cdc7/Drf1 kinase (DDK), which is an essential component of pre-RCs (170). Considerable amounts of cohesin are also associated
with Scc2/4. DDK and other pre-RC components such as Cdt1 are essential for loading both
Scc2/4 and cohesin onto chromatin in these
extracts. The association between DDK and
Scc2/4 requires both the Cdc7 kinase subunit itself and its regulatory Drf1 subunit and involves
Scc2’s N-terminal domain bound by Scc4, but
it does not require the C-terminal HEAT repeat domain. Though not required for association with DDK, the latter is nevertheless essential for cohesin’s loading onto chromatin. These
observations raise the possibility that DDK recruits Scc2/4 to pre-RCs prior to the initiation
of DNA replication and that Scc2/4 in turn recruits cohesin. One suspects that once cohesin
has been recruited to the pre-RC by Scc2/4,
the latter then facilitates chromatin fiber trapping both before and after the initiation of
replication from the pre-RC. If this scenario
is correct, then Scc2/4 has at least two key
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functions: (a) to recruit cohesin to the vicinity
of chromatin and (b) to catalyze chromatin fiber
trapping.
Strangely, pre-RCs do not seem to be required to recruit cohesin to yeast chromosomes
(27, 186), and Scc2/4 complexes have not been
found to be associated with pre-RCs. In addition, expression of α-kleisin in G2 cells, which
no longer possess pre-RCs, leads to cohesin’s
association with chromosomes with a pattern
that is similar if not identical to that of α-kleisin
loaded during S phase (93, 126). The finding
that a large fraction of cohesin in G2 HeLa
cells has a residence time of less than 30 min
(35) also suggests that cohesin must frequently
associate with chromatin de novo long after preRCs have been disassembled. The linkage between pre-RCs, recruitment of the Scc2/4 complex, and loading of cohesin onto chromatin
may therefore be specific to a short stage of the
vertebrate cell cycle.
If, as the Xenopus studies suggest, Scc2/4
interacts directly with cohesin prior to the
latter’s loading onto chromatin, the two complexes may not always maintain this connection
after loading is completed. Chromosome
spreads suggest that cohesin and Scc2/4 do
not colocalize on yeast chromosomes (14).
This notion has also been addressed through
microarray analysis of DNA associated with
cohesin or Scc2/4 immunoprecipitates following formaldehyde cross-linking (ChIP-chip).
Cohesin is enriched in a ∼40-kbp region
around centromeres and at specific sites along
chromosome arms that frequently correspond
to sites of convergent transcription (30, 39, 92).
The distribution of Scc2/4 appears very different and resembles that of condensin and transcription factor IIIC associated with transfer
RNA (tRNA) genes (18). It has been suggested
that cohesin first associates with loci where
Scc2/4 is present and only then moves to sites of
convergent transcription, having possibly been
swept there by RNA polymerase. Interestingly,
a tRNA gene flanking the HMR locus has
been implicated in the establishment of sister
chromatid cohesion at silent mating-type genes
at this locus, raising the possibility that cohesin
rings holding mating-type genes together may
be loaded at the neighboring tRNA gene (26).
A striking example of a specific locus involved in cohesin recruitment is the finding
that enrichment of cohesin for ∼40 kbp around
S. cerevisiae centromeres depends on (a) the 150bp CEN sequence (198), (b) specific proteins
that associate with this sequence, such as the
centromere-specific histone H3, and (c) central kinetochore proteins such as Mtw1 (27).
Remarkably, transfer of a CEN to a chromosomal arm is sufficient to enhance cohesin’s association with sequences in a 20–50-kbp interval surrounding the ectopic kinetochore (198).
Cohesin’s concentration around centromeres
has been visualized in live cells by imaging
GFP-tagged Smc1 or Smc3 proteins (209). The
clearest images are those of metaphase cells in
which Smc3-GFP forms a barrel-like structure
along the spindle axis, with kinetochores that
have bioriented and been split by spindle forces
situated at each end of the barrel and pole-topole microtubules running through the middle of the barrel. Whether some of this cohesin is associated with DNA sequences that
have been pulled apart by the spindle and are
no longer cohesed (C loops) or whether most
of it is associated with neighboring pericentric
DNAs that are still held together by cohesin
is unclear. It has been suggested that cohesin
on C loops may participate in intrachromatid
cohesion, but there is no evidence for this interesting idea. How yeast kinetochore proteins
promote cohesin’s recruitment to pericentric
chromatin is currently mysterious. One explanation for the ability of kinetochores to enhance cohesin’s recruitment to a broad region
around centromeres is that (together with the
Scc2/4 complex) they catalyze entrapment of
sequences close to the core centromere by cohesin rings that subsequently diffuse (or even
are actively moved) to sequences 20 kbp or further away.
An instance of a factor implicated in cohesin
recruitment in S. pombe is the Swi6 HP1-like
factor, which binds to trimethylated K9 residues
on histone H3 around centromeres and silent
mating-type genes. Swi6 mutants have reduced
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amounts of pericentric cohesin and defective
centromeric sister chromatid cohesion (8, 125).
It is uncertain whether HP1 orthologs have a
similar function in mammalian cells, where neither their depletion through RNA interference
(151) nor deletion of a pair of genes encoding
the histone H3 K9 methylase (82) appears to
affect centromeric cohesion adversely.
In summary, it is striking that work on
Xenopus extracts and budding yeast suggests
that cohesin recruitment requires complex
chromatin-bound machines in addition to the
Scc2/4 complex, namely pre-RCs, kinetochores, and the tRNA transcription apparatus.
Moreover, these factors may be only the tip of
the iceberg. This possibility suggests that reproducing the recruitment process in vitro with
defined components, a step that will clearly be
essential for any deep mechanistic understanding, will be a major challenge. Key questions
for the future are whether chromatin fiber entrapment inside cohesin rings is the mechanism
by which they associate stably with chromatin,
whether entrapment involves transient hinge
dissociation, whether and how the Scc2/4 complex participates in this process, and finally how
chromosome locus–specific factors contribute.
The invariable dependence of the loading process on these factors suggests that cohesin (and
all other Smc complexes for that matter) shows
little or no promiscuity in its liaisons with chromatin. One can only assume that such promiscuity is avoided for good reasons, possibly to
prevent establishing potentially damaging connections between chromosomal loci or jeopardizing efficient sister chromatid cohesion at
crucial regions such as around centromeres.
Establishment of Cohesion
Cohesin associates with chromosomes before
DNA replication. What happens to cohesin
rings associated with chromatin fibers ahead of
the replication fork during and after its passage?
Are they transferred in cis to sister chromatids
immediately after the fork in a manner that
sometimes leads to sister chromatid cohesion,
or are they ejected from the chromosome?
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According to one particular version of the
strong ring model, coentrapment of sister
DNAs may be generated from passage of replication forks through rings that had previously
entrapped the unreplicated ancestral fiber (50).
The finding that temperature-sensitive (ts)
scc2-4 yeast cells can establish sister chromatid
cohesion (at the URA3 locus) at the restrictive
temperature after release from arrest in early
S phase by hydroxyurea at the permissive temperature has led to the conclusion that “sister
chromatid cohesion can be built exclusively
with cohesin that was already bound to chromosomes before arrival of the replication fork”
(93, p. 789). The problem is that considerable
replication takes place in cells arrested for long
periods in hydroxyurea, and it is possible that
the cohesion measured had been produced
during the arrest. More rigorous experiments
will be required to settle this key issue. Moreover, the recent finding that cohesion can be
established on fully replicated chromosomes in
the apparent absence of replication forks (161,
163, 190) implies that replication through
cohesin rings cannot be the only mechanism
(if at all) by which cohesion is generated.
What is much clearer is that when expressed
in G2 or M phase yeast cells, components
of the ring complex—namely Smc1/3 and αkleisin subunits—associate with chromosomes
but, under normal circumstances, fail to establish cohesion even in the presence of preexisting sister chromatid cohesion (51, 93, 161, 163,
190). A corollary is that there can be little or
no turnover of these cohesin subunits within
the cohesive structures holding sisters together.
What then enables cohesin present during S
phase to establish cohesion but not that present
in G2 or M phase? A clue lies in genetic studies
that have revealed genes required for establishing cohesion but not for loading it onto chromosomes or for maintaining cohesion already
established. Important but not essential are a
variety of proteins thought to travel with replication forks (73, 207) that are either implicated
in DNA replication, such as Ctf4 (33), or in
the loading or unloading of proliferating cell
nuclear antigen (PCNA) onto chromosomes
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such as the RFCCtf18/Dcc1/Ctf8 complex (104,
105).
More crucial is an acetyl transferase known
as Eco1, whose acetylation of a pair of lysines
(K112 and K113) within Smc3’s NBD, close to
but not immediately adjacent to its ATP binding pocket (Figure 1c), is essential for cohesion
establishment (5, 143, 189, 211). Orthologs of
Eco1 are encoded by most but not all eukaryotic
genomes (121), and mutations invariably result
in cohesion defects (176, 201). Humans contain two such proteins, Esco1 and Esco2 (64).
Mutations in the gene encoding Esco2 are responsible for the developmental defects associated with Roberts syndrome and SCphocomelia
(149, 192), and cells from these patients are defective in centromeric cohesion. Furthermore,
the Smc3 residues corresponding to K112 and
K113 are highly conserved among eukaryotes
and are acetylated in human tissue culture cells
(211). In yeast, Smc3 acetylation increases during S phase in a manner that appears to depend
(a) on the Scc2/4 complex (189), suggesting that
it only occurs if cohesin can associate with chromatin, and (b) at least partly on the prereplication protein Cdc6 (5), implying at least partial
dependency on replication forks. Smc3 acetylation remains high for much of the G2 and
M phases, decreasing around the time cells undergo anaphase (5). This finding suggests that
deacetylation may be triggered by cohesin’s dissociation from chromatin.
What effect does acetylation of K112 and
K113 have on cohesin rings that enables them
to establish connections between sister DNAs?
The discovery that inactivation of Pds5 in
S. pombe (a nonessential cohesin subunit in this
organism) suppresses lethality caused by deletion of Eco1’s ortholog Eso1 (172) raised the
possibility that Eco1’s modification of cohesin
counteracts aspects of its function that otherwise interfere with cohesion establishment
during S phase. Without this “antiestablishment” activity, acetylation of Smc3 is at least
partially redundant. Subsequent genetic studies in budding yeast have revealed that null
alleles of Rad61/Wapl and point mutations
within specific domains of essential cohesin
subunits—namely Smc3, Pds5, and Scc3—have
a similar effect, enabling proliferation without
Eco1 (5, 143, 169). It has been suggested that
although Eco1 itself acts during DNA replication, Smc3 acetylation might be required
postreplication to counteract a tendency of
Rad61 to remove cohesin from chromosomes.
Contrary to the predictions of this model,
Rad61 does not reduce cohesin’s association
with chromosomes in S. cerevisiae, but rather
promotes it (143, 169, 195). Moreover, there is
little or no evidence that cohesin’s association
with chromatin in postreplicative cells is greatly
destabilized by a lack of Smc3 acetylation (143).
Despite the very different phenotypes
caused by inactivating Rad61 in yeast and
Wapl in HeLa cells (see section Dissociation
During Prophase and Anaphase, below), it
is nevertheless likely that these orthologs
operate using similar mechanisms. Both
form stable stoichiometric complexes with
Pds5 (34, 88, 143). Rad61 (at least) also
binds Scc3, whose phosphorylation during mitosis in mammalian cells promotes
cohesin’s dissociation from chromosomes
during prophase (52). Whether these proteins
promote or reduce cohesin’s association with
chromosomes may depend on their state of
modification and the stage of the cell cycle
as well as any evolutionary divergence producing different properties. The Smc3 NBD
appears to be the target of antiestablishment
activities and its acetylation presumably serves
to counteract these during S phase. However,
the process normally mediated by acetylation
of K112 and K113 can also be achieved,
albeit less efficiently, by a variety of mutations
close to Smc3’s ATP binding pocket (5, 143),
including those that do not mimic acetylation.
In summary, these studies indicate that cohesin establishment requires a reorganization
of Smc3’s NBD that can be hindered by
Rad61/Wapl, Pds5, and Scc3 proteins and that
part of the function of Smc3’s acetylation by
Eco1 is to counteract this inhibition.
The finding that Smc3 acetylation is at least
partly dependent on DNA replication (5) suggests that a lack of Smc3 acetylation could
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explain why cohesin subunits produced in G2
or M phase cannot generate sister chromatid
cohesion despite forming cohesin rings that associate stably with chromatin. Consistent with
this notion is the finding that overexpression of
Eco1 enables (some) postreplicative cohesion
establishment (190). Remarkably, DNA damage also enables cohesin to establish sister chromatid cohesion during M phase, even on chromosomes that have not been damaged (161,
190), raising the possibility that signals stemming from local DNA damage and involving
phosphorylation of cohesin’s α-kleisin subunit
by the Chk1 kinase (54) somehow stimulate
more widespread acetylation of cohesin subunits outside of S phase (55).
How might the act of DNA replication or
signals stemming from replication forks facilitate Smc3 acetylation by Eco1? A clue is the
finding that Eco1’s N terminus binds PCNA,
at least in vitro, and that point mutations in
a QXXL/I motif abolish binding and cause
inviability (114). The notion that Eco1 may be
recruited to replication forks through binding
to PCNA would explain why mutations in
the RFCCtf18/Dcc1/Ctf8 complex compromise
cohesion more than DNA replication if this alternative polymerase clamp loader is especially
important in loading those PCNA clamps that
recruit Eco1 to replication forks. Whether the
plethora of genes thought to regulate DNA
replication and to promote sister chromatid
cohesion, e.g., Ctf4, Tof1/Csm3, or Chl1,
facilitate cohesion establishment by helping to
recruit Eco1 to replication forks should be easy
to test. However, whether Eco1 is indeed recruited to replication forks (93) is far from clear.
An important feature of cohesion established in postreplicative cells is that it depends
not only on de novo Eco1 activity but also on
preexisting cohesion (161, 163, 190); it cannot
be generated if the sisters are not already paired.
This is presumably why cohesion must be established during DNA replication (186), even
if it can in principle be repaired or reinforced
by new cohesion produced in G2- or M-phase
cells. The ability to generate new cohesin linkages after S phase may be particularly important
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in cells that spend very long periods in G2, for
example oocytes where linkages produced during S phase may decay with age. Does cohesion
regeneration actually occur under normal circumstances? The observation that the meiosisspecific Smc1 isoform (Smc1B) is synthesized
after S phase and yet contributes to sister chromatid cohesion, albeit so far only in cells artificially induced to enter M phase, suggests so
(138).
Dissociation During Prophase
and Anaphase
Although the mechanics of cohesin’s removal
from chromosomes may be simpler than the
establishment of cohesion in the first place,
the process must nevertheless be very tightly
regulated. Chromosome biorientation during
mitosis is impossible without sister chromatid
cohesion, and its premature loss would be
disastrous. In most eukaryotic cells, cohesin’s
dissociation takes place during two phases of
mitosis (98, 131, 166, 194). The first (known
as the prophase pathway) takes place during
prophase and prometaphase, when most but
not all cohesin dissociates from chromosome
arms but not from centromeres. The second takes place shortly before the onset of
anaphase, when all remaining cohesin (mainly
at centromeres but also on arms) dissociates
due to cleavage of its α-kleisin subunit by
separase (53, 89, 118, 185, 187, 202). These
two processes have different mechanisms in
addition to different temporal regulation, as the
prophase pathway does not involve cleavage by
separase (166). The prophase pathway may not
be an essential aspect of mitotic chromosome
segregation and is largely if not completely
absent during yeast mitosis. Moreover, little or
no cohesin dissociates from chromosome arms
in prometaphase during meiosis I, when sister
chromatid cohesion along chromosome arms
is essential for holding bivalent chromosomes
together. Bivalents are eventually converted
to dyad chromosomes exclusively (possibly)
due to cleavage of arm cohesin complexes by
separase (10, 79, 86, 87, 175).
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How cohesin is removed from chromosome
arms by the prophase pathway is not known.
The ring model predicts that it must involve
ring opening, but whether (and if so how) this
takes place is not known. Key questions are
whether exit and entry gates are the same and
whether they both involve cohesin’s hinge.
The process is facilitated by the Aurora B
and polo-like kinases (PLK) (91, 99, 167),
by PLK-mediated phosphorylation of the
C-terminal domain of cohesin’s SA1/2 Scc3
subunits (52), and by condensin I (60). Aurora
B’s role may be quite indirect, either ensuring
that Sgo1, a protein that protects cohesin
from the prophase pathway, is directed to
centromeres, and/or promoting recruitment
of condensin I to chromosome arms (95).
However, these effectors/processes have quite
modest roles, as most cohesin still dissociates
from chromosome arms in their absence. The
Wapl protein is more crucial. Its depletion
from mammalian tissue culture cells, either
by RNA interference or by gene inactivation
( J.-M. Peters, personal communication),
causes most cohesin to remain on chromosome
arms (34, 88). Wapl is found associated with
cohesin, forms a particularly stable complex
with its Pds5 subunit, is hyperphosphorylated
during mitosis (as are α-kleisin, Pds5, and Scc3
SA1/2), and contributes to turnover of cohesin
on interphase chromosomes. Importantly,
Wapl’s function is not merely to promote
hyperphosphorylation of SA2. The Scc2/4
complex also dissociates from chromosomes
during prophase, which may also contribute to
the removal of arm cohesin (197).
What is the function of the prophase pathway? Unlike the APC/C-separase pathway, it
may not be a universal feature of chromosome segregation. There is no intrinsic reason why α-kleisin cleavage could not deal with
all cohesin (88). Moreover, the prophase pathway has not yet been implicated in any obvious regulatory function. Nevertheless, removal of cohesin from chromosome arms
may facilitate their deconcatenation by Topo
II during prometaphase and metaphase and
may thereby contribute to timely disjunction
at anaphase. Another potential function is
raised by the observation that the large soluble
pool of cohesin created by the prophase pathway is not cleaved by separase at the metaphaseto-anaphase transition (168, 194). A large fraction of this pool reassociates with chromosomes
during telophase and very possibly has important functions regulating transcription and the
structure of chromatin during interphase (131).
If most cohesin were removed from chromosomes by separase, then reassociation would
have to await resynthesis of its α-kleisin subunit, as occurs in budding yeast, with potentially
grave consequences for transcriptional control.
The prophase pathway must not remove
all cohesin from chromosomes, as doing so
would compromise their biorientation on mitotic spindles. Indeed, centromeric cohesin is
protected from the prophase pathway due to
the centromere-specific Sgo1 protein (77, 108,
146). Sgo1 is a member of a class of proteins known as shugoshins, whose founding
member is the Mei-S332 protein in Drosophila
melanogaster (75). Shugoshins possess a conserved coiled-coil domain that binds the ABC
PP2A holoenzyme (139, 174), which is also
localized to centromeres in mitotic cells and
may therefore counteract phosphorylation of
cohesin subunits, Wapl, and Scc2/4. Whether
shugoshins protect centromeric cohesin in mitosis by recruiting PP2A is still unclear, and
there are conflicting reports as to whether
PP2A’s recruitment to centromeres requires
shugoshins (140) or vice versa (80, 174).
Cleavage by Separase
The metaphase-to-anaphase transition is the
point of no return. There is no going back
once cells destroy all sister chromatid cohesion (186). There is overwhelming evidence in
yeast that sister chromatid disjunction is triggered by cleavage of cohesin’s α-kleisin subunit
Scc1 by separase (178, 185, 187). Likewise, resolution of chiasmata, which converts bivalent
chromosomes to dyads at the first meiotic division and requires loss of cohesion along chromosome arms, is achieved by cleavage of the
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meiosis-specific α-kleisin Rec8 (10, 79). Separase activity is also required in Caenorhabditis elegans (153), in Drosophila melanogaster (71),
in mammals (87, 89, 202), and in Arabidopsis
(96). Whether α-kleisin cleavage is either necessary or sufficient in eukaryotic cells other than
yeast is less clear cut. A noncleavable version of
Scc1 interferes with sister chromatid disjunction in HeLa cells, although it may not completely block loss of centromeric cohesion (53),
whereas a supposedly noncleavable version of
Rec8 merely slows down the process of chiasmata resolution (86). The latter causes the
first meiotic division to be aborted and causes
sterility in spermatocytes, but not in oocytes.
The finding that inactivation of separase protease activity completely blocks Rec8’s removal
from bivalents (87), whereas mutation of cleavage sites (detected in vitro) has only a partial effect (86), implies that the failure to arrest completely sister chromatid disjunction is caused by
cryptic cleavage sites that are not detected by
in vitro assays and are still present in supposedly noncleavable variants. There is therefore
no overwhelming reason to doubt that α-kleisin
cleavage is both necessary and sufficient for sister chromatid disjunction in most if not all eukaryotic organisms, even if rigorous proof for
this assertion is still lacking. Thus, there is no
justification at present for concluding that “the
cohesin cleavage model does not ring true” (45).
Separase activity must be highly regulated
and is inhibited for much of the cell cycle
through its association with a specific inhibitory
chaperone known as securin (187). During mitosis, phosphorylation of separase (on serine
1121) by Cdk1 (159) induces stable binding
of cyclin B/Cdk1 (42). This also inactivates
the protease, a process especially important in
postmigratory primordial germ cells (67) and
preimplantation mouse embryos (66), in which
securin levels may be lower than in other cells.
Separase is only (fully) activated when all chromosomes have bioriented, which liberates the
APC/C’s Cdc20 activator protein from sequestration by its Mad2 inhibitor (116, 119). This
permits Cdc20 to recruit securin and cyclin B
to the APC/C, leading to their ubiquitinylation
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and subsequent proteolysis. Upon liberation from securin and cyclin, active separase
cleaves the Scc1 subunits of cohesin (53, 185)
still associated with chromosomes (168, 194),
which induces cohesin’s dissociation from chromatin and triggers sister chromatid disjunction.
Though separase normally only removes cohesin that has resisted the prophase pathway
from chromosomes, it appears capable of removing larger amounts at the metaphase-toanaphase transition, for example when Wapl
depletion inactivates the prophase pathway
(88). Crucially, it is the APC/C-separase pathway, and not the prophase pathway, that is regulated by the SAC (107). Due to its inhibition by two different APC/C substrates (securin and cyclin B) whose ubiquitinylation is
blocked by Mad2, separase can only be fully
activated once the SAC has been turned off.
Some suggest that separase may be partially active during prometaphase, at least when cells
are arrested for prolonged periods in a mitotic state by spindle poisons such as nocodazole (118). How separase selectively cleaves
only α-kleisins associated with chromatin is not
understood (168). Separase is a huge protein
(with over 2000 residues in humans) with an
N-terminal domain containing 26 Armadillo
(ARM) repeats separated from a pair of caspaselike protease domains (only one of which is active) by an unstructured central region. Securin
is thought to bind to the N-terminal ARM repeats, but it may also interact with the protease domain (62). It is conceivable that separase’s ARM-repeat-containing domain binds
chromatin and thereby juxtaposes the protease
with its α-kleisin substrate (193).
One of the more remarkable phenomena
concerning sister chromatid disjunction is the
mechanism that protects Rec8 α-kleisin in the
vicinity of centromeres from separase cleavage at the first meiotic division (134). Cleavage along chromosome arms is essential for the
resolution of chiasmata at meiosis I, but it must
not occur at centromeres because their cohesion is essential for biorientation of dyad chromosomes at meiosis II. It has long been known
that the persistence of centromeric cohesion
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after meiosis I in D. melanogaster depends on a
centromeric protein termed Mei-S332 (40, 75).
Identification of fungal orthologs (78, 102, 136)
revealed a family of conserved proteins (now
known as shugoshins, described above) that
are associated with AB’C PP2A holoenzymes
during meiosis (139). Humans possess two
members of this family, Sgo1, which protects
cohesin from the prophase pathway (108, 146),
and Sgo2, which protects centromeric cohesin
from separase during meiosis I (97). Shugoshins
contain an N-terminal homodimeric parallel
coiled coil whose docking onto PP2A’s C and B
subunits recruits it to centromeres and protects
Rec8 from separase (208). How does recruitment of PP2A to centromeres block α-kleisin
cleavage at this location? If phosphorylation,
either of separase or of cohesin, were necessary for Rec8 cleavage, then PP2A might act
by removing the requisite phosphate groups.
The finding that Rec8’s mitotic counterpart
Scc1 cannot be protected by shugoshin/PP2A
complexes suggests that PP2A’s target is Rec8
itself (181). Rec8 is indeed phosphorylated at
multiple serines and threonines during meiosis
I (9). Crucially, substitution of these residues
by alanine greatly reduces cleavage and hinders chiasmata resolution, whereas substitution with aspartate, mimicking phosphorylation, causes precocious loss of sister centromere
cohesion (V. Katis & K. Nasmyth, unpublished results). The notion that shugoshins
protect centromeric cohesion by dephosphorylating Rec8 is consistent with the finding that
Rec8 phosphorylation is necessary for its efficient cleavage by separase in vitro (86).
A Case Against Cohesin?
In the light of the above discussion, it is curious that doubts have recently been raised as to
whether cohesin really holds sister centromeres
together up until the onset of anaphase and
whether this linkage is destroyed by the
APC/C-separase pathway (15, 23, 38, 45). It
has been suggested that neither APC/C (38) nor
separase (37, 45) is required for the disjunction
of centromeres at anaphase, that depletion of
cohesin does not prevent chromosome biorientation (23), that the fraction of Scc1 molecules
cleaved by separase is insufficient to explain sister chromatid disjunction at anaphase, and that
expression of noncleavable Scc1 in mammalian
cells affects disjunction of sister chromatid
arms more severely than that of centromeres
(23, 127).
The first two of these claims are based
largely on the results of knocking down APC/C
subunits or separase by RNA interference.
Both sets of proteins are notoriously difficult
to deplete efficiently and rapidly using this
technique. More importantly, these claims are
clearly inconsistent with a wealth of data on
the effect of mutations in a wide variety of
micro-organisms and invertebrates. One would
have to argue that sister chromatid disjunction
is mediated by a very different mechanism in
vertebrate cells. However, even this claim is
not borne out by the facts. Complete depletion
of the APC/C using gene deletion in hepatocytes induced to proliferate by partial hepatectomy causes them to arrest in metaphase (203),
whereas mouse embryonic fibroblasts lacking
separase, again due to gene deletion, clearly fail
to disjoin sister chromatids upon activation of
the APC/C. In addition, depletion of separase
in oocytes, via gene deletion, prevents the conversion of bivalent chromosomes into dyads at
meiosis I, and this failure is accompanied by
the persistence of cohesin along their interchromatid axes (87).
The claim that at least some degree of
chromosome biorientation can take place in
mitotic cells lacking cohesin is less controversial. Some studies report considerable biorientation in cohesin-depleted cells (74, 130, 157),
whereas others describe major defects (173,
183). This is an area in which the limitations
of RNA interference are fully exposed. It is difficult to exclude the possibility that apparently
normal biorientation is caused by incomplete
protein depletion, and it can be argued that
more severe defects may be due to off-target
effects. In contrast, there is a consensus that cohesin inactivation via ts mutations, Scc1 cleavage, or gene repression invariably triggers the
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SAC and causes sister chromatid disjunction in
the presence of high levels of cyclin B (130, 183)
or securin (110). What about the argument that
insufficient Scc1 is cleaved to explain anaphase
(23, 45)? The concentration of cleavage fragments is indeed low compared to intact protein, but there is a very good reason for this: the
rapid degradation of the cleavage fragments by
the N-end rule Ubr1 ubiquitin protein ligase
(137).
The last argument is that despite major defects in disjoining sister chromatids at anaphase
(53), cells expressing noncleavable Scc1 protein
appear to separate sister centromeres (23). It
is difficult to ascertain the exact importance of
cleavage without observing mitoses in live cells
that had expressed physiological levels of noncleavable protein prior to the preceding S phase.
This has not yet proved possible. It is moreover
difficult to be certain that the noncleavable variants used hitherto are truly noncleavable and
do not contain cryptic cleavage sites. Despite
this drawback, a case can be made that even
if the noncleavable proteins are not fully noncleavable and even if they are not expressed at
physiological levels, they should surely disrupt
disjunction of sister centromeres as severely
if not more severely as that of chromosome
arms, which appears not to be the case. This
is not a strong argument, but it does suggest
that the role of cohesin cleavage merits further
investigation.
Despite the the fact that the case against
cohesin is weak, might “Topo II and cohesin
contribute equally to regulate sister chromatid
association” (183, p. 2301)? There is no question that decatenation of sister DNAs by Topo
II is required for sister chromatid disjunction,
and it is certainly possible that sister DNA
intertwining may help cells lacking cohesin
to biorient chromosomes for a limited period
during the early phases of mitosis. However,
such cells do not exist in nature as far as we
know. Moreover, there is no evidence that Topo
II is ever inhibited sufficiently during mitosis to ensure that DNA concatenation can resist spindle forces. It therefore makes little
sense to regard the DNA concatenation/Topo
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II system as a backup to the cohesin/separase
system.
There is one clear, albeit peculiar, example
of cohesion between sister chromatid DNAs
that is both cell-cycle regulated and independent of cohesin, namely sister chromatid cohesion between tandem rDNA repeats on chromosome XII in yeast. Possibly due to their high
rates of transcription, sister rDNA loci in yeast
are entangled via a poorly understood cohesinindependent mechanism (11, 19, 165). Artificial
cleavage of cohesin’s Scc1 subunit by TEV protease therefore triggers disjunction of most sister chromatids but not of sister rDNA repeats.
A different mechanism, possibly DNA decatenation, must trigger sister rDNA disjunction.
rDNA loci do not disjoin particularly slowly in
animal cells, and it is unclear whether the process described in yeast also operates in animal
cells.
ROLES BEYOND COHESION
Cohesin and Double-Strand-Break
Repair
Cohesin plays key roles in the repair of DNA
double-strand breaks in mitotic (17, 154) and
meiotic (29, 81, 191) cells. In mitotic cells, two
different populations of cohesin contribute to
the repair process: cohesin engaged in holding
sisters together at the time of the break and cohesin subsequently recruited to chromatin surrounding the break itself (162, 188). Cohesin is
particularly important in meiotic cells, where
programmed double-strand-break repair creates the reciprocal recombination events (chiasmata) that hold bivalent chromosomes together (134). One of the remarkable features of
the meiotic process is that double-strand breaks
are repaired preferentially using nonsister chromatids, a property possibly facilitated by replacing mitotic α-kleisin subunits (Scc1/Rad21) by
meiosis-specific versions (Rec8) (4, 206). Mammals also express meiosis-specific versions of
Smc1 (Smc1B) (138) and Scc3 (STAG3) (135)
and there are therefore a large variety of cohesin complexes during spermatogenesis and
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oogenesis. Cohesin forms the backbone of the
synaptonemal complex (28, 81) and presumably
plays fundamental roles in orchestrating the
repair of meiotic double-strand breaks (160).
Whether cohesin is actively recruited to the
sites of double-strand breaks during meiosis as
it is in mitotic cells is not known. Because of its
crucial role during recombination, loss of function usually leads to a checkpoint-induced arrest at the pachytene stage (4, 138), which hampers analysis of its roles and regulation during
subsequent chromosome segregation (9).
Cohesin and Mono-orientation
During Meiosis I
Work on S. pombe and Arabidopsis indicates that
cohesin is involved in another meiosis-specific
function, namely mono-orientation of sister
kinetochores during meiosis I. During mitosis, microtubules pull sister kinetochores in
opposite directions (biorientation), but during
meiosis I they instead pull maternal and paternal kinetochore pairs in opposite directions.
Electron micrographs show that sister kinetochores are seamlessly bound together in a single
structure at this stage (41). Meiosis-specific
“monopolin” proteins confer this property
(103, 133, 181, 210), but how do they function?
In both S. pombe and Arabidopsis, mutations in
the meiosis-specific α-kleisin Rec8 lead to biorientation of sister kinetochores (13, 196) as well
as to sister chromatid cohesion defects, raising
the possibility that cohesin may connect sister
kinetochores during meiosis I in a way that precludes their biorientation. It appears that Rec8,
but not its mitotic equivalent Rad21, manages
to create sufficient cohesion between sister
kinetochores to preclude their biorientation
(145), a process that only occurs in meiotic cells
and is regulated by monopolins. If such a process is also responsible for mono-orientation in
S. cerevisiae, which is not known, then it is not
an exclusive property of Rec8-containing cohesin complexes, as cells expressing the mitotic
Scc1 instead of Rec8 can also mono-orient
their meiosis I kinetochores (181).
Does Cohesin Mediate Nonsister
Connections Important for
Transcriptional Control?
The first indication that proteins involved in
sister chromatid cohesion may have functions in
addition to holding sister chromatids together
was the description of Roberts syndrome (141)
and SCphocomelia (56) in humans. These are
rare autosomal recessive conditions associated
with limb and growth deficiencies, craniofacial
anomalies, and mild to severe mental deficiency.
Mitotic chromosomes from these patients are
characterized by heterochromatin repulsion,
in particular at centromeres, indicating premature (albeit partial) loss of sister chromatid
cohesion (31, 72, 176). Due possibly to their
defective sister chromatid cohesion, cultured
Roberts syndrome cells have mitotic defects
(177), but it is unclear whether these defects
alone account for either the severe (in the case
of Roberts) or the mild (in the case of SCphocomelia) developmental defects. The presence
of severely and mildly affected individuals in
the same sibship (children from the same set
of parents) indicated that the two disorders
are allelic, and it is now clear that both are
caused by mutations in ESCO2 that abolish its
function (149, 192). What is unclear is whether
the variable developmental defects are caused
by variations between individuals in how their
embryonic cells react to partial loss of sister
chromatid cohesion. Affected cells may differ
in the extent of their arrest by the SAC or in
their propensity to undergo apopotosis. However, given the superficial similarity of the limb
defects with those caused by known regulators
of gene expression, for example HOX genes, it
is not inconceivable that the highly pleiotropic
developmental defects arise due to defects in
the expression of genes regulated by Esco2
through its acetylation of cohesin subunits.
SCphocomelia birth defects are similar to those
caused by thalidomide, which conceivably may
act by interfering with processes regulated by
cohesin.
A stronger clue that cohesin may have nonmitotic functions stems from the discovery that
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half or more patients with Cornelia de Lange
syndrome (CdLS) (20) have mutations in only
one allele of the gene encoding human Scc2,
known also as Nipbl due to its homology with
the fly ortholog Nipped-B (85, 179). CdLS
is a dominantly inherited multisystem developmental disorder characterized by growth
and cognitive retardation; abnormalities of the
upper limbs; gastroesophageal dysfunction;
cardiac, ophthalmologic, and genitourinary
anomalies; hirsutism; and characteristic facial
features. These and other defects are caused
by only modest (30%) reductions in the level
of Scc2/Nipbl protein, which does not appear
to be accompanied by major defects in sister
chromatid cohesion. The homology with Scc2
raised the possibility that CdLS defects may
be caused by reduced cohesin loading, a notion
consistent with the subsequent discovery that
milder forms of CdLS are caused by Smc1A
and Smc3 mutations (21, 117). Consistent with
the notion that CdLS is caused by misregulated gene expression, the D. melanogaster Scc2
ortholog, Nipped-B, facilitates long-range
enhancer-promotor interactions, at least for
certain genes whose regulatory sequences
have been mutated (25, 142). Furthermore,
mutations in mau-2, the C. elegans Scc4 ortholog, cause defects in axon guidance (6) and
two cohesin subunits, Scc1/Rad21 and Smc3
and have been implicated in expression of
the hematopoietic transcription factor Runx1
in zebrafish (63). Despite these findings, the
possibility that developmental cohesinopathies
are caused by knock-on effects of compromising sister chromatid cohesion cannot be fully
excluded.
The discovery that cohesin resides within
the nuclei of most postmitotic cells, including neurons (200), and that cohesin inactivation causes pruning defects in postmitotic
mushroom-body γ-neurons in Drosophila, due
in part to a lack of expression of the ecdysone
receptor, demonstrate unequivocally that cohesin does indeed have nonmitotic functions,
including the regulation of transcription (130,
148). In yeast, such functions include restricting
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the spread of silencing at silent mating-type
loci (24) and regulating transcriptional termination (47). It is striking that in mammalian
cells cohesin frequently colocalizes with the
site-specific zinc finger DNA binding protein
CTCF, which has been proposed to define
boundaries between active and inactive chromatin domains (129, 144, 158, 200). CTCF
appears necessary for cohesin’s localization to
CTCF sites but not vice versa, suggesting that
CTCF recruits cohesin and not the other way
around (144, 200). Recruitment in this sense
does not necessarily mean that CTCF (together
with the Scc2/4 complex) actively promotes
cohesin’s loading onto chromosomes. Indeed,
the amount of cohesin associated with chromatin is little changed in cells depleted for
CTCF (200). If cohesin’s loading onto chromatin invariably involves entrapment of DNA
by cohesin rings that can subsequently diffuse laterally, then CTCF may merely determine where on chromosomes entrapped rings
accumulate.
At the imprinted H19/IGF2 locus where
CTCF and cohesin are recruited in an allele
specific manner to a site between the two genes,
both proteins have a role in preventing activation of the maternal IGF2 allele by an enhancer at the 3 end of the H19 gene (200).
The ability of cohesin rings to entrap sister
chromatin fibers (49) raises the possibility that
they may also be capable of entrapping distant chromatin segments (within a single ring),
thereby creating loops. Cohesin may therefore be a key regulator of long range interactions between enhancers and promoters. Crosslinking studies indicate that such loops form
in a cohesin-dependent fashion at the IFNG
locus in T cells. If so, they do not seem crucial for transcriptional induction, as this is not
greatly affected by cohesin depletion (48). The
finding that cohesin is recruited to the CTCF
sites of V and J segments of immunoglobulin
genes in pro B cells raises the possibility that
cohesin may contribute to contraction of the
locus thought to facilitate gene rearrangement
(22).
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It therefore seems very likely that the
developmental defects caused by a modest drop
in the level of the Scc2/4 (Nipbl) complex are
due to changes in the loading of cohesin onto
chromosomes and thereby to changes in gene
expression. To explain the devastating effect of
such a small change in a single factor, it may be
necessary to postulate that different genes/loci
are in constant competition for a limiting
amount of Scc2/4 and that only a small drop
in its abundance causes major changes in the
amount of cohesin recruited to particular loci.
Microarray studies suggest that cohesin frequently colocalizes with RNA polymerase in
Drosophila tissue culture cells (113). This raises
the possibility that cohesin may be a “hitchhiking device” to which other factors can bind
while it (cohesin) moves along chromosomes
with RNA polymerase.
A Function at Centrosomes?
Two studies have recently detected cohesin subunits at spindle poles (204) or centrosomes (83).
Whether or not this population of cohesin has
an important role at these locations is less clear.
Cohesin depletion does indeed cause spindle
pole defects, but it is difficult at this stage to exclude the possibility that these are indirect consequences of prior mitotic defects caused by the
lack of sister chromatid cohesion. Nevertheless,
these are intriguing findings in the light of the
suggestion that separase may have a function in
centriole disengagement (184).
FUTURE ISSUES
1. Does cohesin’s stable association with chromatin involve trapping of chromatin fibers?
Are sister DNAs held together due to their entrapment inside monomeric cohesin rings?
2. Is transient dissociation of cohesin’s hinge required for DNA entrapment? Does the
Scc2/4 complex facilitate DNA trapping?
3. What are the mechanistic consequences of Smc3 acetylation?
4. What are the structural consequences of ATP binding and hydrolysis by Smc proteins?
5. Is postreplicative cohesion establishment on undamaged chromosomes of physiological
importance?
6. Does cohesin regulate transcription by generating loops between distant DNAs and do
Esco1/2 modulate cohesin activity for the purposes of transcriptional control?
7. To answer these questions, it will be important to (a) determine atomic structures of
the Smc3 NBD-kleisin interface, the non-Smc proteins Scc3, Scc2/4, Wapl, and Pds5;
(b) develop in vitro systems that recapitulate key steps in cohesin’s association with chromatin fibers; and (c) develop assays to measure cohesion and/or DNA loop formation in
vitro in systems other than budding yeast.
DISCLOSURE STATEMENT
The authors are not aware of any affiliations, memberships, funding, or financial holdings that
might be perceived as affecting the objectivity of this review.
ACKNOWLEDGMENTS
We thank Arthur C. Leung for pointing out the possibility that Scc1 passage through disengaged
Smc1/3 hinge domains can be used to entrap DNA within cohesin rings, Sara Cuylen for help
with figures, and all our colleagues for suggestions and discussions.
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