Research Article
3493
SA1 binds directly to DNA through its unique AT-hook
to promote sister chromatid cohesion at telomeres
Kamlesh K. Bisht, Zharko Daniloski and Susan Smith*
Kimmel Center for Biology and Medicine at the Skirball Institute, Department of Pathology, New York University School of Medicine, New York,
NY 10016, USA
*Author for correspondence ([email protected])
Journal of Cell Science
Accepted 7 May 2013
Journal of Cell Science 126, 3493–3503
ß 2013. Published by The Company of Biologists Ltd
doi: 10.1242/jcs.130872
Summary
Sister chromatid cohesion relies on cohesin, a complex comprising a tri-partite ring and a peripheral subunit Scc3, which is found as two
related isoforms SA1 and SA2 in vertebrates. There is a division of labor between the vertebrate cohesin complexes; SA1-cohesin is
required at telomeres and SA2-cohesin at centromeres. Depletion of SA1 has dramatic consequences for telomere function and genome
integrity, but the mechanism by which SA1-cohesin mediates cohesion at telomeres is not well understood. Here we dissect the
individual contribution of SA1 and the ring subunits to telomere cohesion and show that telomeres rely heavily on SA1 and to a lesser
extent on the ring for cohesion. Using chromatin immunoprecipitation we show that SA1 is highly enriched at telomeres, is decreased at
mitosis when cohesion is resolved, and is increased when cohesion persists. Overexpression of SA1 alone was sufficient to induce
cohesion at telomeres, independent of the cohesin ring and dependent on its unique (not found in SA2) N-terminal domain, which we
show binds to telomeric DNA through an AT-hook motif. We suggest that a specialized cohesion mechanism may be required to
accommodate the high level of DNA replication-associated repair at telomeres.
Key words: SA1, Cohesion, Telomeres
Introduction
Sister chromatid cohesion ensures the faithful distribution of sister
chromatids in mitosis and provides a template for homology
directed repair and recombination in late S and G2 phase of the cell
cycle. Cohesion is mediated by cohesin, a ring complex comprising
Smc1, Smc3, and Scc1(Anderson et al., 2002; Haering et al.,
2002), and a peripheral subunit Scc3, that in vertebrates exists as
two closely related isoforms SA1 and SA2 (Losada et al., 2000;
Sumara et al., 2000). A cohesin complex contains either SA1 or
SA2. In somatic cells SA2 is found in great excess (12- to 15-fold)
over SA1 (Holzmann et al., 2011). Precisely how the ring works in
vivo to cohere sisters is not known, but models range from the ring
encircling the sisters to hold them together (Gruber et al., 2003) to
rings on each sister that are held together by a protein bridge
(Chang et al., 2005; Zhang et al., 2008).
The best-understood and most fundamental role of cohesin is at
centromeres, where it concentrates to hold sister chromatids
together against the spindle forces until the metaphase to anaphase
transition (Tanaka et al., 2000). Cohesin is also distributed along
chromosome arms, albeit at a much lower frequency; the average
distance between two cohesin sites is ,340 kb in human cells
(Wendt et al., 2008). Cohesin binds to the same sites as the
CCCTC-binding factor CTCF (Parelho et al., 2008; Rubio et al.,
2008; Stedman et al., 2008; Wendt et al., 2008) a zinc finger DNAbinding protein that acts as a transcriptional activator/repressor and
insulator-binding protein (Wallace and Felsenfeld, 2007),
indicating a role for cohesin in the organization of interphase
chromatin. Cohesin is also required for DNA repair; it is recruited
to sites of double strand breaks to hold the sisters in close
proximity for repair (Ström et al., 2004; Ünal et al., 2004). The
observation that cohesin needs to be recruited to sites of damage,
suggests that normally there is insufficient cohesion to hold the
sisters in the close proximity needed for DNA repair.
Telomeres are unique heterochromatic structures that contain
TTAGGG repeats and shelterin, a six-subunit complex
comprising TRF1, TRF2, TIN2, RAP1, TPP1 and POT1 (de
Lange, 2005). Shelterin function is regulated by transiently
associating proteins, such as tankyrase 1, a poly(ADP-ribose)
polymerase that PARsylates TRF1 (Smith et al., 1998) and can
influence association of TRF1 and its shelterin binding partner
TIN2 with telomeres (Houghtaling et al., 2004; Smith and de
Lange, 2000). Telomeres rely on specialized mechanisms for
their replication (Gilson and Géli, 2007; Stewart et al., 2012),
protection (Palm and de Lange, 2008), and cohesion. The first
indication for a distinct cohesion mechanism came with the
observation that sister telomeres remained cohered at mitosis in
normal human cells approaching senescence (Ofir et al., 2002;
Yalon et al., 2004). A subsequent study in HeLa cells showed that
depletion of tankyrase 1 led to persistent telomere (but not arm or
centromere) cohesion at mitosis (Dynek and Smith, 2004).
Subsequently, the shelterin subunits TIN2 and TRF1, were found
to be associated with SA1-cohesin, but not SA2-cohesin
(Canudas et al., 2007). In cells depleted of TIN2 or SA1, but
not SA2, telomere cohesion was not established in S phase
(Canudas and Smith, 2009). Conversely, depletion of SA2, but
not SA1 or TIN2, led to a defect in centromere cohesion
(Canudas and Smith, 2009). Additional studies showed that the
heterochromatin protein HP1c associated with TIN2 and was
required for cohesion at telomeres, but not at centromeres
(Canudas et al., 2011).
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The studies described above indicate a unique mode of
telomere cohesion in human cells that relies exclusively upon
SA1-cohesin. Similar results were obtained in mouse cells
deleted for SA1 (Remeseiro et al., 2012a). The need for
specialized cohesion at telomeres is likely due to the repetitive
G-rich structure, which poses problems for replication fork
progression, leading to nicks and gaps that must be repaired prior
to mitosis. Consistent with this notion, depletion of SA1 led to
the inability to repair breaks in G2 and sister telomere loss in
human cells (Canudas and Smith, 2009), and to fragile telomeres
in mouse cells (Remeseiro et al., 2012a). Despite its importance
in genome integrity, the mechanism by which SA1-cohesin
mediates sister telomere cohesion is not well understood.
Here we dissect the individual contribution of SA1 and the ring
subunits to telomere cohesion. Surprisingly, we find that while
depletion of the ring subunits dramatically affects centromere
cohesion, it only minimally affects cohesion at telomeres. Instead,
we show that SA1, through its unique AT-hook, is the major
driving force for cohesion at telomeres. Hence, telomeres employ
(in addition to the canonical ring-mediated cohesion that occurs
along chromosome arms) a specialized SA1-mediated mechanism
for cohesion. We suggest that the difficulty to replicate G-rich
DNA at telomeres demands a specialized (more intimate)
association between sister chromatids to allow continual
surveillance and repair during and after DNA replication.
Results
Resolution of telomere cohesion occurs in G2/M and is
dependent on PARP-active tankyrase 1
Telomere cohesion (unlike centromere cohesion) cannot be
analyzed by metaphase-spread analysis; telomere cohesion is
sensitive to the hypotonic swelling conditions used during
metaphase-spread preparation (Canudas et al., 2007; Dynek and
Smith, 2004). Hence, for all the experiments described herein, in
order to preserve the protein-protein interactions that mediate
sister telomere cohesion, live cells are fixed immediately
(without pretreatment) in methanol-acetic acid and subjected
directly to fluorescent in situ hybridization (FISH).
We showed previously (using chromatin immunoprecipitation
(ChIP) analysis across the cell cycle) that tankyrase 1 localized to
telomeres in G2/M (Bisht et al., 2012). To determine if this
localization coincided temporally with resolution of telomere
cohesion, we performed FISH analysis with a chromosomespecific subtelomere probe across the cell cycle. Cells were
treated with TNKS1 siRNA or GFP as a control, synchronized by
a double thymidine block, released for 4, 6, 8 and 10 hours and
analyzed by FISH with a 16ptelo probe (Fig. 1A,B). In control
cells in S phase (4–6 hours after release) the majority of
telomeres appeared as singlets indicating that they were
cohered. As cells entered G2/M (8–0 hours after release),
telomere cohesion was resolved (Fig. 1A, upper panel and
Fig. 1B, left side), coinciding precisely with the time when
tankyrase 1 localizes to telomeres (Bisht et al., 2012). In cells
depleted of tankyrase 1, telomeres remained as singlets throughout
the cell cycle (Fig. 1A, bottom panel and Fig. 1B, right side). To
determine if resolution of telomeres and arms was coincident
across the cell cycle, double FISH was performed with probes
against the telomere and arm of the same chromosome (13qtelo
13qarm). In control cells resolution of telomere and arm cohesion
occurred simultaneously across the cell cycle (Fig. 1C, upper
panel and Fig. 1D, left side). By contrast, in tankyrase-1-depleted
cells arm, but not telomere, cohesion was resolved in G2/M
(Fig. 1C, lower panel and Fig. 1D, right side).
To demonstrate the specificity of resolution of telomere
cohesion, we asked if it required the catalytic activity of
tankyrase 1. Cells were treated with TNKS1 siRNA and cotransfected with siRNA resistant plasmids encoding wild type or
PARP-dead tankyrase 1. Cells were synchronized by a double
thymidine block, released for 10 hours, isolated by mitotic shakeoff, and analyzed by immunoblot (Fig. 1E) and by FISH with a
16ptelo probe (Fig. 1F,G). Wild type tankyrase 1, but not PARPdead, rescued the persistent sister telomere cohesion. Consistent
with the notion that the catalytic activity of tankyrase 1 was
required for resolution of telomere cohesion, treatment of cells
with the tankyrase-specific PARP inhibitor XAV939 (Huang
et al., 2009) led to persistent sister telomere cohesion (Fig. 1H,I).
Thus, the PARP activity of tankyrase 1 is required to disrupt
cohesion at telomeres.
We showed previously that persistent telomere cohesion
induced by tankyrase 1 depletion could be rescued by depletion
of SA1, but not SA2 (Canudas et al., 2007) and see Fig. 1J–L. We
now sought to determine if depletion of the cohesin ring would
similarly rescue the tankyrase 1-induced persistent cohesion. Cells
were subjected to double siRNA treatment with TNKS1 and
individual cohesin subunits, analyzed by immunoblot (Fig. 1J),
and by FISH with a 16ptelo probe following mitotic shake-off
(Fig. 1K,L). Depletion of SA1 led to a 6-fold reduction in cells
with singlets, rescuing cohesion to nearly wild type levels. By
contrast, depletion of SA2 led to only a 1.3-fold reduction in cells
with singlets (Fig. 1K,L). Surprisingly, depletion of the ring
subunit Scc1 had only a minimal effect (1.3-fold; similar to SA2),
indicating that persistent telomere cohesion in tankyrase-1depleted cells is mediated largely by SA1, with only minimal
contribution by SA2 and the cohesin ring.
A limited requirement for cohesin ring subunits in
telomere cohesion
We next asked if the cohesin ring was required to maintain
cohesion at telomeres irrespective of tankyrase 1. Following
treatment with siRNA for each cohesin subunit, cells were
analyzed by immunoblot (Fig. 2A) and by FISH with a 16ptelo
probe following mitotic shake-off (Fig. 2B–F,G). In HeLaI.2.11
cells treated with GFP siRNA sister telomeres appeared as
closely associated doublets separated by an average distance of
0.76 mm (Fig. 2B). Depletion of SA1 led to a dramatic increase
(average distance 2.14 mm) in the distance between most sister
telomeres, whereas in SA2-depleted cells most sister telomeres
remained close, although a subset showed increased distance
[Fig. 2C,D, and shown previously (Canudas and Smith, 2009)].
In cells depleted of the ring subunits Scc1 or Smc3, most sister
telomeres remained closely associated as in control cells,
although a subset showed increased distance (Fig. 2E,F).
Together these data show that in cells depleted of SA2 or the
ring (but not SA1), the majority of sister telomeres remain closely
associated, suggesting that SA2 and the ring contributed only
minimally to telomere cohesion.
To validate that cohesin ring function was impaired in the
Scc1- and Smc3-depleted cells, we measured centromere
cohesion using FISH with a chromosome-specific centromere
probe 6cen (Fig. 2H–L,M). In HeLaI.2.11 cells treated with GFP
siRNA centromeres were cohered, appearing as closely
associated doublets with an average diameter of 0.62 mm
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SA1 promotes telomere cohesion through its AT-hook
Fig. 1. See next page for legend.
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(Fig. 2H). Depletion of SA1 led to only a minor effect (1.2-fold
increase) in the distance between sister centromeres, whereas in
SA2-depleted cells the majority of sister centromeres showed an
increased distance [Fig. 2I,J, and shown previously (Canudas and
Smith, 2009)]. In cells depleted of the ring subunits Scc1 or
Smc3, the distance between most sister centromeres was
increased dramatically (Fig. 2K,L). Together these data show
that in cells depleted of SA2 or the ring, the majority of sister
centromeres have lost cohesion (Fig. 2M), whereas the majority
of sister telomeres retain their close association (Fig. 2G).
Indeed, double FISH analysis of Scc1- or Smc3-depleted cells
showed that within the same cell telomere cohesion remained
intact despite a dramatic loss in centromere cohesion (Fig. 2N).
We were surprised that depletion of the ring had only a limited
effect on telomere cohesion measured at mitosis. To probe the
issue further we queried the impact of ring depletion on
establishment of cohesion at telomeres in S phase. siRNAtreated HeLa.I.2.11 cells were synchronized with a double
thymidine block, harvested 4 hours after release (mid S phase),
and processed for FACS analysis (Fig. 3A) and chromosomespecific telomere (Fig. 3B,D) or centromere (Fig. 3C,E) FISH.
FACS analysis indicated that the siRNA treated cells were in S
phase (Fig. 3A). In cells treated with GFP siRNA the majority
of telomeres and centromeres appeared as singlets, consistent
with the view that in mid-S-phase chromosomes are either
unreplicated or replicated, but still cohered. Cells depleted of
SA1 showed a dramatic (5.5-fold) increase in telomere doublets,
whereas SA2-depleted cells showed only a 1.8-fold increase in
telomere doublets [Fig. 3B,D, and shown previously (Canudas
and Smith, 2009)]. In cells depleted of the ring subunits Scc1 or
Fig. 1. Resolution of telomere cohesion occurs in G2/M and is dependent
on PARP-active tankyrase 1. (A–D) Cell cycle analysis of cohesion.
HeLa.I.2.11 cells were synchronized by a double thymidine block, treated
with control (GFP) or TNKS1 siRNA, released at 4, 6, 8 and 10 hours, and
analyzed by FISH with (A) a 16ptelo (green) or (C) a 13qtelo (green) and
13qarm (red) probe. (B,D) Graphical representation of the frequency of cells
with singlet FISH signals from (A; n5250 cells or more each) and (C; n5150
cells each), respectively. (E–I) Resolution of sister telomere cohesion requires
the PARP activity of tankyrase 1. (E–G) Supertelomerase HeLa cells were
synchronized by a double thymidine block, co-transfected with tankyrase 1
siRNA and a control vector (V) or a plasmid containing tankyrase 1 wild type
(WT) or PARP-Dead (PD), released for 10 hours, isolated by mitotic shakeoff, and analyzed by (E) immunoblot or by (F) FISH with a 16ptelo probe.
(G) Graphical representation of the frequency of mitotic cells with singlet
FISH signals derived from two independent experiments. Values are means 6
s.e.m. (n520 cells or more each). (H,I) HeLa1.2.11 cells were synchronized
by a double thymidine block, released for 10 hours in the absence (2) or
presence (+) of the tankyrase PARP inhibitor XAV939 (1 mm), isolated by
mitotic shake-off and analyzed by (H) FISH with a 16ptelo probe.
(I) Graphical representation of the frequency of mitotic cells with singlet
FISH signals. Values are means 6 s.d. from three independent experiments
(n599 cells or more each). (J–L) Tankyrase 1-induced persistent telomere
cohesion is rescued by depletion of SA1, but not by depletion of SA2 or the
cohesin ring. HeLaI.2.11 cells were transfected without (2) or with (+)
TNKS1 siRNA along with a second siRNA against GFP, SA1, SA2 or Scc1
for 48 hours and analyzed by (J) immunoblot (protein levels relative to atubulin and normalized to the GFP siRNA control are indicated next to the
blots) or isolated by mitotic shake-off, and analyzed by (K) FISH using a
16ptelo probe (green). (L) Graphical representation of the frequency of
mitotic cells with unseparated telomeres in mitosis. Values are means 6
s.e.m. from two independent experiments (n5120 cells or more each). In A,
C, F, H, and K DNA was stained with DAPI (blue). Scale bars: 5 mm.
Smc3, most telomeres remained as singlets, although there was
a (2-fold) increase in doublets (Fig. 3B,D,F). Analysis of
centromere cohesion under the same conditions revealed the
opposite; depletion of SA1 had no effect, whereas depletion of
SA2, Scc1 or Smc3 resulted in at least a 4-fold increase in
doublets (Fig. 3C,E,F). Together these data support the notion
that SA1 drives telomere cohesion, whereas, SA2 along with the
cohesin ring drives centromere cohesion.
SA1 is enriched at telomeres
The studies described above raised the possibility that SA1 may be
acting in a unique or independent way at telomeres. We thus asked
if we could detect accumulation of SA1 at human telomeres. The
genomic distribution of SA1 was measured previously using ChIPChIP on the non-repetitive ENCODE regions representing 1% of
the human genome (Rubio et al., 2008; Wendt et al., 2008).
However, these studies did not address the association with
telomeres. To address this question, we performed a specialized
telomeric ChIP assay that uses a dot blot approach to hybridize the
immunoprecipitated DNA with a telomere repeat probe (Telo).
Alu repeats are used as a negative control. ChIP analysis of
HeLaI.2.11 cells using anti-TRF1 antibody revealed a robust
telomere-specific signal (Fig. 4A,B), as expected for a telomericDNA-binding protein. ChIP with anti-SA1 antibody revealed a
weaker, but specific (and reproducible) telomeric association
(Fig. 4A,B). To validate that the increased SA1 signal observed at
telomeres was due to SA1 protein, ChIP was performed on SA1depleted cells. Immunoblot analysis of cells treated with GFP or
SA1 siRNA showed efficient knockdown of SA1 (Fig. 4C) and a
corresponding knockdown of the SA1-ChIP signal (Fig. 4D,E).
Telomere cohesion is released at mitosis in a tankyrase 1
dependent manner (described above; Fig. 1A–D). Accordingly,
we asked if SA1 was released from telomeres at mitosis.
HeLaI.2.11 cells were synchronized by a double thymidine block,
released, and collected at 0 and 10 hours. FACS analysis
indicated that the cells were synchronized in G1/S (0 hours)
and G2/M (10 hours; Fig. 4F). ChIP analysis of the cell cycle
staged extracts indicated a greater than 2-fold reduction of SA1 at
telomeres in mitosis (Fig. 4G,H). To determine if removal of
SA1 relied on tankyrase 1, HeLaI.2.11 cells were treated with
GFP or TNKS1 siRNA and mitotic cells were isolated by mitotic
shake-off. Immunoblot analysis showed efficient knockdown of
TNKS1 (Fig. 4I). ChIP analysis revealed a threefold increase in
SA1 at telomeres in TNKS1-depleted mitotic cells (Fig. 4J,K).
Together the data are consistent with a role for SA1 in telomere
cohesion; SA1 is enriched at telomeres, is reduced at telomeres in
mitosis when cohesion is removed, and is increased at telomeres
in TNKS1-depleted mitotic cells when cohesion persists.
SA1 overexpression promotes sister telomere cohesion
Our studies thus far indicated that SA1 was enriched at telomeres
where it was required (to a greater extent than SA2 and the cohesin
ring) to establish and maintain sister telomere cohesion. SA1 and
SA2 are highly conserved along their length, where they bind to
the cohesin ring through association with Scc1, but are
distinguished by unique sequences in their N- and C-termini
(Fig. 5A). To dissect the telomeric specificity of SA1 versus SA2,
HeLaI.2.11 cells were transiently transfected with epitope-tagged
alleles, FlagSA1 or FlagSA2. Transfected cells were analyzed
by immunoblot (Fig. 5B) and isolated by mitotic shake-off for
FISH analysis with a 16ptelo probe. As shown in Fig. 5C,D,
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SA1 promotes telomere cohesion through its AT-hook
Fig. 2. Telomere cohesion is maintained in mitosis in cohesin-ring-depleted cells. (A) Immunoblot analysis of extracts from HeLaI.2.11 cells transfected with
siRNA to GFP, SA1, SA2, Scc1 or Smc3 for 48 hours and probed with the indicated antibodies. Protein levels relative to a-tubulin and normalized to the
GFP siRNA control are indicated below the blots. (B–F) Telomere and (H–L) centromere FISH analysis of HeLaI.2.11 cells isolated by mitotic shake-off
following 48 hours transfection with GFP (B,H), SA1 (C,I), SA2 (D,J), Scc1 (E,K) or Smc3 (F,L) siRNA with a 16ptelo (green) or 6cen (red) probe. The cen locus
is trisomic. DNA was stained with DAPI (blue). Scale bars: 5 mm. Histograms showing the distance between FISH signals (n598–225) are on the right
with the average (Avg) distance (6 s.e.m.) indicated. (G,M) Graphical representation of the average distance (6 s.e.m.) between sister telomeres (G) or
centromeres (M). (N) Combined telomere and centromere FISH analysis. HeLaI.2.11 cells were transfected with GFP, Scc1 or Smc3 siRNA for 48 hours and
probed with 16ptelo (green) and 10cen (red). The cen locus is trisomic. DNA was stained with DAPI (blue). Scale bar: 5 mm.
overexpression of SA1 induced a fivefold increase in persistent
telomere cohesion at mitosis, whereas overexpression of SA2 had
a less than twofold effect.
Our previous observation that the 72-amino-acid domain of
SA1 (but not the corresponding 69-amino-acid domain in SA2)
bound to the shelterin subunit TRF1 (Canudas et al., 2007),
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Fig. 3. Telomere cohesion is established in S phase in cohesinring-depleted cells. HeLa.I.2.11 cells were synchronized by a
double thymidine block, treated with siRNA against GFP, SA1,
SA2, Scc1 or Smc3 and analyzed by (A) FACS, (B) telomere FISH
with a telomere 16ptelo (green), or (C) centromere FISH with a
6cen (red) probe 4 hours after release from the second thymidine
block. DNA was stained with DAPI (blue). Scale bar: 5 mm.
(D,E) Graphical representation of the frequency of telomere (from
B; n5514 cells or more each) and centromere (from C; n5441
cells or more each) doublets in S phase, respectively. Values are
means 6 s.e.m., derived from two independent experiments.
(F) Graphical representation of the fold increase in telomere (from
D) and centromere (from E) doublets relative to the GFP siRNA
control.
suggested that this domain could be important for telomere
cohesion. Hence, we generated a chimeric protein replacing the
N-terminal domain of SA2 with that of SA1 (SA1/SA2; Fig. 5A).
FISH analysis of HeLaI.2.11 cells transiently transfected with an
epitope-tagged allele of the chimera (FlagSA1/SA2) induced
persistent telomere cohesion similar to SA1 (Fig. 5C,D),
consistent with the notion that the unique N-terminal domain of
SA1 imparts telomere specificity to SA1-mediated cohesion.
To further dissect the telomere specificity of SA1 in cohesion,
HTC75 cell lines stably overexpressing FlagSA1 or an Nterminally deleted allele of SA1 (FlagSA1D72) were generated
and analyzed by immunoblot (Fig. 5E) and isolated by mitotic
shake-off for FISH analysis with a 16ptelo probe. As shown in
Fig. 5F,H, the SA1-HTC75 cell line, but not the SA1D72-HTC75
line, showed persistent sister telomere cohesion in mitosis,
consistent with a role for the 72-amino-acid N-terminal domain
of SA1 in telomere cohesion. To determine if the effect of SA1
on cohesion was specific to telomeres, double FISH was
performed with probes against the telomere and arm of the
same chromosome. As shown in Fig. 5G,H, FISH analysis with
the 13qtelo probe (like the 16ptelo probe) showed persistent
telomere cohesion dependent on the N-terminal domain of SA1.
By contrast, FISH analysis with an arm probe from the same
chromosome (13qarm) showed only a slight increase in cohesion
and this was independent of the N-terminal domain of SA1.
Finally, to determine the protein requirements for the persistent
sister telomere cohesion induced by overexpression of SA1, the
SA1-HTC75 cell line was treated with siRNA against GFP
control, the cohesin ring subunit Scc1 or the shelterin subunit
TIN2 and analyzed by immunoblot (Fig. 5I) and isolated by
mitotic shake-off for FISH analysis with a 16ptelo probe. As
shown in Fig. 5J,K, depletion of Scc1 did not rescue the
persistent cohesion induced by SA1. By contrast, depletion of
TIN2, shown previously to be required for sister telomere
cohesion (Canudas et al., 2007; Canudas and Smith, 2009),
partially rescued the SA1-induced persistent cohesion at
telomeres. Together these data indicate that SA1 induces
persistent cohesion (specifically at telomeres) that is dependent
SA1 promotes telomere cohesion through its AT-hook
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Fig. 4. SA1 associates with telomeres in vivo. (A,B) Telomeric
ChIP analysis of TRF1 and SA1. (A) Autoradiograph showing
telomeric DNA ChIP analysis of HeLaI.2.11 cells using the
indicated antibodies. Dot blots with the immunoprecipitated
DNA were analyzed by Southern blotting with 32P-labelled
telomeric (Telo) or Alu repeat (Alu) probes. (B) Graphical
representation of the percentage of immunoprecipitated Telo or
Alu DNA relative to total input DNA (as in A), derived from four
independent experiments. Values are means 6 s.d.
(C–E) Ablation of the SA1 signal at telomeres by SA1 siRNA.
HelaI.2.11 cells were transfected with siRNA to GFP or SA1 for
48 hours and analyzed by (C) immunoblotting with antibodies
against SA1 or a-tubulin and (D) telomeric ChIP with antibodies
against TRF1 or SA1. (E) Graphical representation of the
percentage of SA1-immunoprecipitated Telo or Alu DNA
relative to total input DNA (as in D). (F–H) SA1 is reduced at
telomeres in mitosis. HeLaI.2.11 cells were synchronized by a
double thymidine block, released, and collected by trypsinization
at 0 hours (G1/S) or 10 hours (G2/M) and analyzed by (F) FACS
(y-axis, cell numbers; x-axis, relative DNA content based on
propidium iodide staining) and (G) telomeric ChIP with
antibodies against TRF1 or SA1. (H) Graphical representation of
the percentage of SA1-immunoprecipitated Telo or Alu DNA
relative to total input DNA (as in G). Values are means 6 s.e.m.,
derived from two independent experiments. (I–K) SA1 is
increased at telomeres in TNKS1-depleted mitotic cells.
HelaI.2.11 cells were transfected with siRNA to GFP or TNKS1
for 48 hours, isolated by mitotic shake-off, and analyzed by
(I) immunoblotting with antibodies against TNKS1, SA1 or
a-tubulin and (J) telomeric ChIP with antibodies against TRF1 or
SA1. (K) Graphical representation of the percentage of SA1immunoprecipitated Telo or Alu DNA relative to total input
DNA (as in J). Values are means 6 s.e.m., derived from two
independent experiments.
on its unique N-terminal TRF1-binding domain and is
independent of the cohesin ring.
SA1 binds directly to DNA through a N-terminal AT-hook
motif
The studies described above indicate a crucial role for the unique
N-terminal domain of SA1 in mediating telomere cohesion.
Inspection of the amino acid sequence revealed the presence of
an AT-hook motif at amino acid position 34 [previously identified
in a screen for all detectable AT-hooks in the protein sequence
database (Aravind and Landsman, 1998)]. The AT-hook motif, first
described in the high mobility group proteins (HMGA1/2), binds to
AT-rich DNA sequences in the minor groove of DNA in a nonsequence-specific manner (Reeves, 2001). The AT-hook motif in
SA1 (KRKRGRP) is highly conserved, but is not found in SA2
(Fig. 6A). To determine if the AT-hook containing N-terminal
domain of SA1 binds to DNA, we generated recombinant proteins
(Fig. 6B) and performed gel shift assays with a 32P-labeled duplex
(TTAGGG)12 probe and poly(dG-dC) as non-specific competitor
(Fig. 6C). As a positive control we used TRF1 (Fig. 6B, lane 6),
which binds specifically to telomere repeats (Chong et al., 1995)
(shown in Fig. 6C, lanes 2 and 3). For SA1, the 72-amino-acid Nterminal domain was expressed as a Sumo fusion protein and
purified from E. coli (Fig. 6B, lane 2). Sumo-SA1.72 (like TRF1)
bound to telomeric DNA (Fig. 6C, lanes 4 and 5).
To determine if the binding was due to the AT-hook motif, we
generated a double point mutation, converting the arginines
[predicted to participate in DNA–protein interactions (Huth et al.,
1997)] at position 37 and 39 to alanines (Sumo-SA1.72RA, see
Fig. 6A). As shown in Fig. 6C, lane 6, Sumo-SA1.72RA did not
bind telomeric DNA, indicating a requirement for the AT-hook
motif in DNA binding. As additional controls, we show that an
unrelated Sumo protein (Sumo-GMD, Fig. 6B, lane 5) (Bisht
et al., 2012) did not bind DNA (Fig. 6C, lane 7) and that the
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Journal of Cell Science 126 (15)
Journal of Cell Science
Fig. 5. SA1 induces persistent sister
chromatid cohesion at telomeres.
(A) Schematic representation of FLAGtagged SA1, SA2 and mutant alleles SA1/SA2
and SA1D72. (B–D) Overexpression of SA1
but not SA2, induced persistent telomere
cohesion. HeLaI.2.11 cells were transfected
with vector, SA1, SA2 or SA1/SA2 for
20 hours and analyzed by (B) immunoblot
and (C) telomere FISH using a 16ptelo probe
(green) following mitotic shake-off.
(D) Graphical representation of the frequency
of mitotic cells with unseparated telomeres.
Values are means 6 s.e.m., derived from two
independent experiments (n568 cells or more
each). (E–H) SA1 induced persistent
cohesion specifically at telomeres, dependent
on its N-terminal domain. Stable HTC75 cell
lines overexpressing SA1 or SA1D72 or
control cells (Con) were analyzed by
(E) immunoblot, (F) telomere FISH using a
16ptelo probe or (G) by double FISH with a
13qtelo (green) and 13qarm (red) probe
following mitotic shake-off. (H) Graphical
representation of the frequency of mitotic
cells with unseparated telomeres or arms
(n5102 cells or more each). (I–K) SA1
induced persistent telomere cohesion
independent of Scc1, but dependent on TIN2.
Stable SA1-HTC75 or control cells were
treated without (2) or with siRNA against
GFP, Scc1 or TIN2 and analyzed by
(I) immunoblot (protein levels relative to atubulin and normalized to the GFP siRNA
control are indicated next to the blots) and
(J) telomere FISH using a 16ptelo probe
(green) following mitotic shake-off.
(K) Graphical representation of the frequency
of mitotic cells with unseparated telomeres.
Values are means 6 s.e.m., derived from two
independent experiments (n529 cells or more
each). In C, F, G and J DNA was stained with
DAPI (blue). Scale bars: 5 mm.
SA1-72 protein alone (cleaved and purified away from Sumo,
Fig. 6B, lane 4) bound to the (TTAGGG)12 probe (Fig. 6C, lanes
8 and 9), like Sumo-SA1-72.
To determine if SA1 showed the same specificity for telomeric
DNA as TRF1, we asked if E. coli DNA could compete with binding
of SA1 to telomeric DNA. As shown in Fig. 6D, under conditions
where the TRF1-(TTAGGG)12 complex was resistant to competition
(Fig. 6D, lanes 2 and 3), the SA1-(TTAGGG)12 complex was
competed by E. coli DNA (Fig. 6D, lanes 4 to 6), consistent with the
notion that AT-hooks do not bind in a sequence-specific fashion to
DNA. Together these data indicate the while SA1 is not a sequencespecific DNA-binding protein like TRF1, it does binds to telomeric
(and likely other AT-rich) DNA through its AT-hook motif.
Finally, to determine if the AT-hook was required for telomere
cohesion in vivo, we created the same (RA) double point
mutation in the full-length FLAG-tagged SA1 allele to generate
FlagSA1RA. HeLaI.2.11 cells were transiently transfected with
FlagSA1 or FlagSA1RA, analyzed by immunoblot (Fig. 6E), and
isolated by mitotic shake-off for FISH analysis with a 16ptelo
probe. As shown in Fig. 6F, G, mutation of the AT-hook abrogated
the ability of SA1 to induce persistent cohesion at telomeres.
Discussion
Telomeres are unique heterochromatic structures that rely on
specialized mechanisms for their replication and protection.
Perhaps it is not surprising then that they utilize a novel
mechanism for their cohesion. The repetitive, G rich nature of
telomeres makes them highly susceptible to replication damage
and hence more reliant on homology directed repair with the sister
chromatid to restore genome integrity following DNA replication
in late S and G2 phases of the cell cycle. Our study indicates that
SA1, through its unique AT-hook, drives cohesion at telomeres.
SA1 promotes telomere cohesion through its AT-hook
3501
Journal of Cell Science
Fig. 6. SA1 binds to telomeric DNA in
vitro and promotes persistent telomere
cohesion in vivo through its AT-hook.
(A) Alignment of the SA1 domain
containing the AT-hook. Amino acids
identical to human SA1 are in bold. The
AT-hook RA mutation is indicated on the
right. (B–D) SA1 binds telomeric DNA.
(B) Purified recombinant proteins (500 ng
each) were fractionated by SDS-PAGE and
visualized by staining with Coomassie Blue.
(C,D) Autoradiographs of the telomere
repeat binding assay with the indicated
recombinant proteins and a 32P-labeled
(TTAGGG)12 probe. SA1 binds to telomeric
DNA dependent on the AT-hook (C).
Addition of E. coli competitor DNA
competes for SA1, but not TRF1, binding to
telomeric DNA (D). (E–G) SA1 promotes
persistent telomere cohesion dependent on
its AT-hook. HeLaI.2.11 cells were
transfected with a vector control, SA1 and
SA1RA for 20 hours and analyzed by
(E) immunoblot and (F) telomere FISH
using a 16ptelo probe (green) following
mitotic shake-off. Scale bar: 5 mm.
(G) Graphical representation of the
frequency of mitotic cells with unseparated
telomeres. Values are means 6 s.e.m.,
derived from two independent experiments
(n5102 cells or more each). (H) Model of
how SA1 promotes cohesion independent of
the cohesin ring by associating with
telomeric DNA and proteins.
We suggest that SA1, along with the shelterin subunits TIN2 and
TRF1 and heterochromatin protein HP1c, promote an intimate
association between sister telomeres that allows continual
surveillance during and after DNA replication that cannot be
achieved by the cohesin ring (see model in Fig. 6H).
Our studies indicate that depletion of the cohesin ring, despite
having a robust effect at centromeres, only minimally affects
cohesion at telomeres. This is perhaps not surprising since cohesin
is highly enriched at centromeres where it plays an essential role
holding sister centromeres together against the forces of the mitotic
spindle. In our study depletion of the ring subunits (Smc3 and
Scc1) phenocopied depletion of SA2 (Figs 2, 3), consistent with
the notion that these subunits (SA2, Smc3 and Scc1) act in concert.
However, depletion of the ring did not phenocopy depletion of
SA1, suggesting that these subunits (SA1, Smc3 and Scc1) do not
act in concert. Depletion of SA2, Smc3 or Scc1 had some impact
on telomere cohesion, but it was significantly less than the impact
of SA1 depletion, suggesting a distinct role for SA1 at telomeres.
Indeed, we detected enrichment of SA1 at telomeres by ChIP
(Fig. 4), but we did not detect SA2, Scc1 or Smc3 at telomeres
(data not shown). While this could be due to differences between
antibodies used for SA1 versus the other subunits, given the sparse
distribution of cohesin rings along chromosome arms [one every
340 kb (Wendt et al., 2008)], we favor the interpretation that
telomeres, which are 20–25 kb in HeLa1.2.11cells, may be devoid
of rings. We suggest that the small cohesion defects observed at
telomeres upon depletion of SA2 and the ring, are due to loss of
cohesion at the level of the whole chromosome, from the
neighboring arm or centromere. Together our studies suggest
that the robust effect of SA1 depletion or overexpression on
telomere cohesion is due to intrinsic properties of the SA1 subunit.
SA1 contains an AT-hook in its N-terminal domain that is
highly conserved in SA1 homologs from other species and is not
found in SA2 (Fig. 6A). We show that SA1 binds directly to
telomeric DNA in vitro (Fig. 6C). However, this binding does not
have the specificity of the sequence-specific telomere-repeatbinding protein TRF1, since in the case of SA1, we can compete
the binding using E. coli competitor DNA (Fig. 6D), consistent
with the idea that AT-hooks bind in a non-sequence-specific
manner (Reeves, 2001). We suggest that the AT-hook endows
SA1 with the capacity to associate with telomeres where it then
binds to TRF1 and other proteins to promote cohesion.
Our finding that SA1 contains a functional AT-hook may shed
light on additional roles of SA1 in vivo, revealed recently by
work in mice. Knockout of SA1 in mouse resulted in embryonic
lethality (Remeseiro et al., 2012a; Remeseiro et al., 2012b).
While analysis of SA1-null mouse cells indicated defects in
telomere cohesion and telomere replication (Remeseiro et al.,
2012a) [consistent with studies in human cells (Canudas and
Smith, 2009)], additional observations indicated a role for SA1 in
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Journal of Cell Science 126 (15)
different processes. Analysis of the distribution of SA1-cohesin
versus SA2-cohesin in the non-repetitive parts of the mouse
genome by ChIP-sequencing indicated striking differences in
their genomic distribution; SA1 was enriched at promoters and
SA1 null cells showed dramatic changes in gene expression
(Remeseiro et al., 2012b). Considering that AT-hook containing
proteins have been shown to function in chromatin remodeling
and transcription, it is tempting to speculate that SA1 may
directly impact transcription through its unique AT-hook.
The initial finding that the Scc3 subunit of cohesin existed as
two isoforms in vertebrate cells was surprising. However, given
the strong homology between the two (and their association with
the ring) it seemed likely that it was simply a gene duplication
event that might not lead to dramatic difference in function.
However, studies in human and mouse cells show that SA1 is
required for sister chromatid cohesion at telomeres and its
depletion has dramatic consequences on cell and organismal
function despite the presence of SA2. Our work here, identifying
a functional AT-hook motif that is unique to SA1, provides a
molecular mechanism and sheds slight on the role of SA1 at
telomeres and potentially throughout the genome.
Journal of Cell Science
Materials and Methods
Plasmids
The SA1 cDNA (kindly provided by Jose Luis Barbero) containing amino acids 1–
1258 and the SA2 cDNA containing amino acids 1–1231 were cloned into the vector
p3XFLAG-CMV-10 (Sigma) to generate 3XFlagSA1 and 3XFlagSA2. The SA1/SA2
chimera was generated by replacing amino acids 1–69 of FlagSA2 with SA1 amino
acids 1–70; the resulting hybrid contained three alanines at the junction site. SumoSA1.72 contains amino acids 1–72 of SA1 cloned into the BamHI and XhoI sites of the
pET28b-Sumo-6xHis vector (Chen et al., 2008). The SA1RA mutation was created
by substituting the arginine (R) residues at position 37 and 39, with alanine (A)
residues, by site-directed mutagenesis of Sumo-SA1.72 and 3XFlagSA1 using the
oligonucleotide 59-CAGAGGTCAAAGGAAAAAGAAAAGCGGGTGCTCCTGGCCGGCC-39. Mutagenesis was performed using the Stratagene QuikChange sitedirected mutagenesis kit according to the manufacturer’s instructions. For tankyrase 1
constructs, the tankyrase1 cDNA under the CMV promoter, derived from plasmid
TT20 (Smith et al., 1998), was inserted into a modified pLKO.1ps vector (a kind gift
from Bill Hahn, Dana-Farber Cancer Institute, Boston, MA). The siRNA resistant
(Dynek and Smith, 2004) and tankyrase 1 PARP-dead (Cook et al., 2002) plasmids
were generated as previously described.
Chromatin lmmunoprecipitation
HeLaI.2.11 cells were processed for ChIP as described previously (Bisht et al.,
2012). Following preparation of cell lysates, 25% was saved as input.
Immunoprecipitation was performed by addition of the following antibodies: 5 ml
of rabbit TRF1 415 (Cook et al., 2002) crude sera, 5 mg goat anti-SA1 BL143G
(Bethyl Laboratories, Inc.); or 5 mg goat IgG. Hybridization with a 32P-TTAGGG or
Alu probe was performed in Church buffer [0.5 M sodium phosphate buffer (pH 7.2),
1% BSA, 1 mM EDTA, 7% SDS] as described previously (de Lange, 1992). Washed
membranes were exposed to Kodak XAR film and ImageJ software was used to
quantify the percentage of precipitated DNA relative to the input DNA.
medium, and transfected with siRNA. After 10 hours the medium was replaced with
medium containing 2 mM thymidine and the cells were incubated for 16 hours,
washed three times with PBS, and released into fresh medium. Cells were then
harvested by trypsinization at 4, 6, 8 and 10 hours and analyzed by FISH.
For the tankyrase 1 rescue experiment supertelomerase HeLa cells (Cristofari
and Lingner, 2006) were synchronized and transfected with siRNA as describe
above, but were in a addition, transfected with tankyrase 1 plasmids prior to the
second thymidine block. After release for 10 hours, cells were harvested by mitotic
shake-off and analyzed by immunoblot and FISH.
For the XAV939 inhibition, HeLaI.2.11 cells were synchronized by a double
thymidine block as describe above, released for 10 hours in 0.5% serum with or
without 1 mM XAV939, harvested by mitotic shake-off, and analyzed by FISH.
Stable cell lines
The SA1 cDNA (amino acids 1–1258) (Carramolino et al., 1997; Losada et al.,
2000) was obtained from The I.M.A.G.E. Consortium. The clone (accession no.
AAH64699) contained an internal deletion from amino acids 1150–1186. SA1 and
SA1D72 (lacking the first 72 amino acids) were cloned into the vector p3XFLAGCMV-10 (Sigma) to generate 3XFlagSA1 and 3XFlagSA1D72, described
previously (Canudas et al., 2007), and then inserted into the retroviral vector
pLPCX (Clontech). To create stable cell lines, amphotropic retroviruses were
generated by transfecting FlagSA1 and FlagSA1D72 into phoenix amphotropic
cells (ATCC) using calcium phosphate precipitation. HTC75 cells [an HT1080derived clonal cell line (van Steensel and de Lange, 1997)] were infected and
selected in 2 mg/ml puromycin as described (Houghtaling et al., 2004).
Cell extracts
Cells were resuspended in 4 volumes of TNE buffer [10 mM Tris (pH 7.8), 1%
Nonidet P-40, 0.15 M NaCl, 1 mM EDTA, and 2.5% protease inhibitor cocktail
(PIC) (Sigma)] and incubated for 1 hour on ice. Suspensions were pelleted at 8000
g for 15 minutes. Twenty-five micrograms (determined by Bio-Rad protein assay)
of supernatant proteins were fractionated by SDS-PAGE and analyzed by
immunoblotting.
Immunoblot analysis
Immunoblots were incubated separately with the following primary antibodies:
mouse anti-a-tubulin ascites (1:10,000; Sigma); goat anti-SA1 BL143G (1 mg/ml;
Bethyl Laboratories, Inc.); goat anti-SA2 BL146G (1 mg/ml; Bethyl Laboratories,
Inc.); rabbit anti-Scc1 (2 mg/ml; Bethyl Laboratories, Inc.); rabbit anti-Smc3 (1 mg/
ml; Abcam); rabbit anti-TIN2 701 (0.5 mg/ml) (Houghtaling et al., 2004); or rabbit
anti-tankyrase 1 762 (1.8 mg/ml) (Scherthan et al., 2000), followed by horseradish
peroxidase-conjugated donkey anti-rabbit or anti-mouse IgG (1:2500; Amersham).
Bound antibody was detected by Super Signal West Pico (Thermo Scientific).
siRNA transfection
siRNA transfections were performed with in HeLaI.2.11 cells, a HeLa-derived
clonal cell line (van Steensel et al., 1998) or HTC75 cell lines with Oligofectamine
(Invitrogen) according to the manufacturer’s protocol. The final concentration of
siRNA was 100 nM. The following siRNAs (synthesized by Dharmacon Research
Inc.) were used: TNKS1 (59-AACAAUUCACCGUCGUCCUCU-39) described
previously (Dynek and Smith, 2004); SA1.a (59-GUGAUGCCUUCCUAAAUGA39); SA2.a (59-GUACGGCAAUGUCAAUAUA-39); and TIN2.a (59-AACGCCUUUGUAUGGGCCUAA-39) described previously (Canudas et al., 2007); Scc1
(59-GGUGAAAAUGGCAUUACGGUU-39) described previously (Watrin et al.,
2006); Smc3 (59-AUCGAUAAAGAGGAAGUUUUU-39) described previously
(Kueng et al., 2006); and GFP Duplex I. Following 48 hours transfection, cells
were isolated by mitotic shake-off and processed for chromosome-specific FISH.
Plasmid transfections
Chromosome-specific FISH
Cells were fixed and processed as described previously (Dynek and Smith, 2004).
Briefly, cells were fixed twice in methanol:acetic acid (3:1) for 15 minutes,
cytospun (Shandon Cytospin) at 2000 rpm for 2 minutes onto slides, rehydrated in
26 SSC at 37 ˚C for 2 minutes, and dehydrated in an ethanol series of 70%, 80%
and 95% for 2 minutes each. Cells were denatured at 75 ˚C for 2 minutes and
hybridized overnight at 37 ˚C with the following probes from Cytocell: a
subtelomeric FITC-conjugated probe (16ptelo); a chromosome-6-specific alphasatellite TRITC-conjugated centromere probe (6cen); a TRITC-conjugated
chromosome 10 centromere probe (10cen); or a dual probe comprising a
TRITC-conjugated 13qarm and a FITC-conjugated 13q subtelomere probe. Cells
were washed in 0.46SSC at 72 ˚C for 2 minutes, and in 26SSC with 0.05% Tween
20 at RT for 30 seconds. DNA was stained 0.2 mg/ml DAPI. The distance between
FISH signals was measured using OpenLab software (Perkin Elmer).
Cell synchronization
For FISH analysis across the cell cycle, HelaI.2.11 cells were grown in the presence
of 2 mM thymidine for 16 hours, washed three times with PBS, released into fresh
HeLa1.2.11 cells were transfected with Lipofectamine 2000 (Invitrogen) according
to the manufacturer’s protocol. Following 20 hours transfection, cells were
isolated by mitotic shake-off and processed for chromosome-specific FISH.
Purified recombinant proteins and gel-shift assays
TRF1 was purified from baculovirus as described previously (Bianchi et al., 1997).
Recombinant Sumo-SA1.72, Sumo-SA1.72RA, and Sumo-GMD (Bisht et al., 2012)
were expressed and purified from E. coli BL21 cells according to standard protocols.
Digestion and removal of the SUMO protein tag to generate SA1.72 was performed
with SUMO Protease 1 according to the manufacturer’s instructions (Life Sensors).
Gel-shift assays were performed with a 32P-end-labeled XbaI fragment from plasmid
pTH12 (a kind gift from Titia de Lange, The Rockefeller University, New York,
NY) containing 12 tandem TTAGGG repeats. Purified recombinant protein (200–
300 ng) was incubated for 25 minutes at room temperature in a 20 ml reaction
containing 20 mM Hepes-KOH (pH 7.5), 100 mM KCl, 5% glycerol, 1 mM EDTA,
0.1 mM MgCl2, 0.5 mM DTT, 100 ng casein, 5 ng polydG-dC, and 3 ng labeled
probe. Competition experiments were performed by adding 5–25 ng E. coli
DNA. Samples were fractionated on 4% polyacrylamide native gels run in 0.1%
SA1 promotes telomere cohesion through its AT-hook
TBE (Tris-Borate-EDTA) at 150 V for 55 minutes in the cold. Prior to loading
samples, gels were prerun for 30 minutes at 80 V in the cold. Gels were dried onto
Whatman DE81 paper and autoradiograph.
FACS analysis
siRNA transfected, trypsinized cells were washed twice with PBS containing 2 mM
EDTA, fixed in cold 70% ethanol and stained with propidium iodide (50 mg/ml) and
analyzed using a Becton-Dickenson FACSAN and FlowJo 8.8.6 software.
Image acquisition
Images were acquired using a microscope (Axioplan 2; Carl Zeiss, Inc.) with a
Plan Apochrome 636 NA 1.4 oil immersion lens (Carl Zeiss, Inc.) and a digital
camera (C4742-95; Hamamatsu Photonics). Images were acquired and processed
using Openlab software (Perkin Elmer).
Acknowledgements
We thank members of the Smith lab and Tom Meier for comments
on the manuscript and helpful discussion.
Author contributions
K.K.B. and Z.D. designed and performed experiments and
interpreted data. S.S. designed experiments, interpreted data, and
wrote the manuscript.
Journal of Cell Science
Funding
This work was supported by the National Cancer Institute of the
National Institutes of Health [grant number R01CA116352 to S.S.].
Deposited in PMC for release after 12 month.
References
Anderson, D. E., Losada, A., Erickson, H. P. and Hirano, T. (2002). Condensin and
cohesin display different arm conformations with characteristic hinge angles. J. Cell
Biol. 156, 419-424.
Aravind, L. and Landsman, D. (1998). AT-hook motifs identified in a wide variety of
DNA-binding proteins. Nucleic Acids Res. 26, 4413-4421.
Bianchi, A., Smith, S., Chong, L., Elias, P. and de Lange, T. (1997). TRF1 is a dimer
and bends telomeric DNA. EMBO J. 16, 1785-1794.
Bisht, K. K., Dudognon, C., Chang, W. G., Sokol, E. S., Ramirez, A. and Smith,
S. (2012). GDP-mannose-4,6-dehydratase is a cytosolic partner of tankyrase 1 that
inhibits its poly(ADP-ribose) polymerase activity. Mol. Cell. Biol. 32, 3044-3053.
Canudas, S. and Smith, S. (2009). Differential regulation of telomere and centromere
cohesion by the Scc3 homologues SA1 and SA2, respectively, in human cells. J. Cell
Biol. 187, 165-173.
Canudas, S., Houghtaling, B. R., Kim, J. Y., Dynek, J. N., Chang, W. G. and Smith,
S. (2007). Protein requirements for sister telomere association in human cells. EMBO
J. 26, 4867-4878.
Canudas, S., Houghtaling, B. R., Bhanot, M., Sasa, G., Savage, S. A., Bertuch,
A. A. and Smith, S. (2011). A role for heterochromatin protein 1c at human
telomeres. Genes Dev. 25, 1807-1819.
Carramolino, L., Lee, B. C., Zaballos, A., Peled, A., Barthelemy, I., Shav-Tal, Y.,
Prieto, I., Carmi, P., Gothelf, Y., González de Buitrago, G. et al. (1997). SA-1, a
nuclear protein encoded by one member of a novel gene family: molecular cloning
and detection in hemopoietic organs. Gene 195, 151-159.
Chang, C. R., Wu, C. S., Hom, Y. and Gartenberg, M. R. (2005). Targeting of
cohesin by transcriptionally silent chromatin. Genes Dev. 19, 3031-3042.
Chen, Y., Yang, Y., van Overbeek, M., Donigian, J. R., Baciu, P., de Lange, T. and
Lei, M. (2008). A shared docking motif in TRF1 and TRF2 used for differential
recruitment of telomeric proteins. Science 319, 1092-1096.
Chong, L., van Steensel, B., Broccoli, D., Erdjument-Bromage, H., Hanish, J., Tempst,
P. and de Lange, T. (1995). A human telomeric protein. Science 270, 1663-1667.
Cook, B. D., Dynek, J. N., Chang, W., Shostak, G. and Smith, S. (2002). Role for the
related poly(ADP-Ribose) polymerases tankyrase 1 and 2 at human telomeres. Mol.
Cell. Biol. 22, 332-342.
Cristofari, G. and Lingner, J. (2006). Telomere length homeostasis requires that
telomerase levels are limiting. EMBO J. 25, 565-574.
de Lange, T. (1992). Human telomeres are attached to the nuclear matrix. EMBO J. 11,
717-724.
de Lange, T. (2005). Shelterin: the protein complex that shapes and safeguards human
telomeres. Genes Dev. 19, 2100-2110.
Dynek, J. N. and Smith, S. (2004). Resolution of sister telomere association is required
for progression through mitosis. Science 304, 97-100.
Gilson, E. and Géli, V. (2007). How telomeres are replicated. Nat. Rev. Mol. Cell Biol.
8, 825-838.
Gruber, S., Haering, C. H. and Nasmyth, K. (2003). Chromosomal cohesin forms a
ring. Cell 112, 765-777.
Haering, C. H., Löwe, J., Hochwagen, A. and Nasmyth, K. (2002). Molecular
architecture of SMC proteins and the yeast cohesin complex. Mol. Cell 9, 773-788.
3503
Holzmann, J., Fuchs, J., Pichler, P., Peters, J. M. and Mechtler, K. (2011). Lesson
from the stoichiometry determination of the cohesin complex: a short protease
mediated elution increases the recovery from cross-linked antibody-conjugated beads.
J. Proteome Res. 10, 780-789.
Houghtaling, B. R., Cuttonaro, L., Chang, W. and Smith, S. (2004). A dynamic
molecular link between the telomere length regulator TRF1 and the chromosome end
protector TRF2. Curr. Biol. 14, 1621-1631.
Huang, S. M., Mishina, Y. M., Liu, S., Cheung, A., Stegmeier, F., Michaud, G. A.,
Charlat, O., Wiellette, E., Zhang, Y., Wiessner, S. et al. (2009). Tankyrase
inhibition stabilizes axin and antagonizes Wnt signalling. Nature 461, 614-620.
Huth, J. R., Bewley, C. A., Nissen, M. S., Evans, J. N., Reeves, R., Gronenborn, A. M. and
Clore, G. M. (1997). The solution structure of an HMG-I(Y)-DNA complex defines
a new architectural minor groove binding motif. Nat. Struct. Biol. 4, 657-665.
Kueng, S., Hegemann, B., Peters, B. H., Lipp, J. J., Schleiffer, A., Mechtler, K. and
Peters, J. M. (2006). Wapl controls the dynamic association of cohesin with
chromatin. Cell 127, 955-967.
Losada, A., Yokochi, T., Kobayashi, R. and Hirano, T. (2000). Identification and
characterization of SA/Scc3p subunits in the Xenopus and human cohesin complexes.
J. Cell Biol. 150, 405-416.
Ofir, R., Yalon-Hacohen, M., Segev, Y., Schultz, A., Skorecki, K. L. and Selig,
S. (2002). Replication and/or separation of some human telomeres is delayed beyond
S-phase in pre-senescent cells. Chromosoma 111, 147-155.
Palm, W. and de Lange, T. (2008). How shelterin protects mammalian telomeres.
Annu. Rev. Genet. 42, 301-334.
Parelho, V., Hadjur, S., Spivakov, M., Leleu, M., Sauer, S., Gregson, H. C., Jarmuz,
A., Canzonetta, C., Webster, Z., Nesterova, T. et al. (2008). Cohesins functionally
associate with CTCF on mammalian chromosome arms. Cell 132, 422-433.
Reeves, R. (2001). Molecular biology of HMGA proteins: hubs of nuclear function.
Gene 277, 63-81.
Remeseiro, S., Cuadrado, A., Carretero, M., Martı́nez, P., Drosopoulos, W. C.,
Cañamero, M., Schildkraut, C. L., Blasco, M. A. and Losada, A. (2012a).
Cohesin-SA1 deficiency drives aneuploidy and tumourigenesis in mice due to
impaired replication of telomeres. EMBO J. 31, 2076-2089.
Remeseiro, S., Cuadrado, A., Gómez-López, G., Pisano, D. G. and Losada,
A. (2012b). A unique role of cohesin-SA1 in gene regulation and development.
EMBO J. 31, 2090-2102.
Rubio, E. D., Reiss, D. J., Welcsh, P. L., Disteche, C. M., Filippova, G. N., Baliga,
N. S., Aebersold, R., Ranish, J. A. and Krumm, A. (2008). CTCF physically links
cohesin to chromatin. Proc. Natl. Acad. Sci. USA 105, 8309-8314.
Scherthan, H., Jerratsch, M., Li, B., Smith, S., Hultén, M., Lock, T. and de Lange,
T. (2000). Mammalian meiotic telomeres: protein composition and redistribution in
relation to nuclear pores. Mol. Biol. Cell 11, 4189-4203.
Smith, S. and de Lange, T. (2000). Tankyrase promotes telomere elongation in human
cells. Curr. Biol. 10, 1299-1302.
Smith, S., Giriat, I., Schmitt, A. and de Lange, T. (1998). Tankyrase, a poly(ADPribose) polymerase at human telomeres. Science 282, 1484-1487.
Stedman, W., Kang, H., Lin, S., Kissil, J. L., Bartolomei, M. S. and Lieberman,
P. M. (2008). Cohesins localize with CTCF at the KSHV latency control region and at
cellular c-myc and H19/Igf2 insulators. EMBO J. 27, 654-666.
Stewart, J. A., Chaiken, M. F., Wang, F. and Price, C. M. (2012). Maintaining the
end: roles of telomere proteins in end-protection, telomere replication and length
regulation. Mutat. Res. 730, 12-19.
Ström, L., Lindroos, H. B., Shirahige, K. and Sjögren, C. (2004). Postreplicative
recruitment of cohesin to double-strand breaks is required for DNA repair. Mol. Cell
16, 1003-1015.
Sumara, I., Vorlaufer, E., Gieffers, C., Peters, B. H. and Peters, J. M. (2000).
Characterization of vertebrate cohesin complexes and their regulation in prophase.
J. Cell Biol. 151, 749-762.
Tanaka, T., Fuchs, J., Loidl, J. and Nasmyth, K. (2000). Cohesin ensures bipolar
attachment of microtubules to sister centromeres and resists their precocious
separation. Nat. Cell Biol. 2, 492-499.
Ünal, E., Arbel-Eden, A., Sattler, U., Shroff, R., Lichten, M., Haber, J. E. and
Koshland, D. (2004). DNA damage response pathway uses histone modification to
assemble a double-strand break-specific cohesin domain. Mol. Cell 16, 991-1002.
van Steensel, B. and de Lange, T. (1997). Control of telomere length by the human
telomeric protein TRF1. Nature 385, 740-743.
van Steensel, B., Smogorzewska, A. and de Lange, T. (1998). TRF2 protects human
telomeres from end-to-end fusions. Cell 92, 401-413.
Wallace, J. A. and Felsenfeld, G. (2007). We gather together: insulators and genome
organization. Curr. Opin. Genet. Dev. 17, 400-407.
Watrin, E., Schleiffer, A., Tanaka, K., Eisenhaber, F., Nasmyth, K. and Peters,
J. M. (2006). Human Scc4 is required for cohesin binding to chromatin, sisterchromatid cohesion, and mitotic progression. Curr. Biol. 16, 863-874.
Wendt, K. S., Yoshida, K., Itoh, T., Bando, M., Koch, B., Schirghuber, E., Tsutsumi,
S., Nagae, G., Ishihara, K., Mishiro, T. et al. (2008). Cohesin mediates
transcriptional insulation by CCCTC-binding factor. Nature 451, 796-801.
Yalon, M., Gal, S., Segev, Y., Selig, S. and Skorecki, K. L. (2004). Sister chromatid
separation at human telomeric regions. J. Cell Sci. 117, 1961-1970.
Zhang, N., Kuznetsov, S. G., Sharan, S. K., Li, K., Rao, P. H. and Pati, D. (2008). A
handcuff model for the cohesin complex. J. Cell Biol. 183, 1019-1031.