Download The CHFR mitotic checkpoint protein delays cell cycle

Oncogene (2005) 24, 2589–2598
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ORIGINAL PAPERS
The CHFR mitotic checkpoint protein delays cell cycle progression by
excluding Cyclin B1 from the nucleus
Matthew K Summers1,2, John Bothos1,2 and Thanos D Halazonetis*,1,3
1
Molecular and Cellular Oncogenesis Program, The Wistar Institute, Philadelphia, PA 19104-4268, USA; 2Program in Cell and
Molecular Biology, Biomedical Graduate Studies, University of Pennsylvania, Philadelphia, PA 19104, USA; 3Department of
Pathology and Laboratory Medicine, University of Pennsylvania, Philadelphia, PA 19104, USA
CHFR, a novel checkpoint gene inactivated in human
cancer, delays chromosome condensation in cells treated
with microtubule poisons. To understand the molecular
mechanism for this delay, we characterized cells with
inactivated CHFR and stably transfected derivatives
expressing the wild-type gene. After exposure to microtubule poisons, the CHFR-expressing cells arrested
transiently in early prophase with a characteristic ruffled
morphology of the nuclear envelope and no signs of
chromosome condensation. Several markers suggested
that Cyclin A/Cdc2 had been activated, whereas Aurora-A and -B and Cyclin B1/Cdc2 were inactive. Further,
Cyclin B1 was excluded from the nucleus. Ectopic
expression of Cyclin B1 with a mutant nuclear export
sequence induced chromosome condensation, and thus
overcame the CHFR checkpoint. We conclude that the
mechanism by which CHFR delays chromosome condensation involves inhibition of accumulation of Cyclin B1
in the nucleus.
Oncogene (2005) 24, 2589–2598. doi:10.1038/sj.onc.1208428
Published online 24 January 2005
Keywords: CHFR; Cyclin B1; checkpoint; prophase;
mitosis
Introduction
The process by which eucaryotic cells segregate their
genetic material during cell division is referred to as
mitosis. Mitosis has been divided into distinct phases
based mostly on morphological criteria. In early
prophase, the centrosomes separate along the nuclear
periphery. In mid and late prophase, the chromosomes
condense. This is then followed by breakdown of the
nuclear envelope, which indicates entry into prometaphase. In metaphase, the chromosomes align on the
spindle equator. Separation of the sister chromatids
defines entry into anaphase and, finally, chomatid
*Correspondence: TD Halazonetis, Wistar Institute, 3601 Spruce
Street, Philadelphia, PA, USA;
E-mail: [email protected]
Received 30 August 2004; accepted 1 December 2004; published online
24 January 2005
decondensation occurs in telophase (McIntosh and
Koonce, 1989).
Since mitosis requires the ordered execution of several
biochemically distinct processes, eucaryotic cells have
developed checkpoint mechanisms to ensure the timely
execution of these events. The best-characterized mitotic
checkpoint is the spindle checkpoint, which inhibits
sister chromatid separation until all chromosomes have
aligned on the mitotic spindle (Amon, 1999; Shah and
Cleveland, 2000). This checkpoint is required for cells to
properly progress through mitosis. Thus, complete
inactivation of the spindle checkpoint leads to embryonic lethality and has not been described in cells that
retain the capacity to divide (Basu et al., 1999; Dobles
et al., 2000; Kalitsis et al., 2000). However, partial
inactivation of the spindle checkpoint, induced mostly
by heterozygous mutations of spindle checkpoint genes,
has been documented in some cancer cells, albeit
infrequently, and may contribute to the genetic instability that is characteristic of human cancer (Cahill
et al., 1998, 1999; Michel et al., 2001).
In addition to the spindle checkpoint, a checkpoint
that functions early in mitosis has also been described
(Jha et al., 1994; Rieder and Cole, 2000; Scolnick and
Halazonetis, 2000). The only gene known so far to
participate in this checkpoint is CHFR (Checkpoint with
FHA and RING domains). In CHFR-expressing cells,
exposure to microtubule poisons leads to a delay in
chromosome condensation (Scolnick and Halazonetis,
2000). Unlike the spindle checkpoint, the CHFR
checkpoint is not essential for cell proliferation. Perhaps
because of this, the CHFR gene can be inactivated in
human cancer, either by promoter methylation or in a
few cases by missense mutations. The overall frequency
of CHFR inactivation in cancer is quite high, ranging
from 15 to 50% in various tumor types (Mizuno et al.,
2002; Shibata et al., 2002; Bertholon et al., 2003; Corn
et al., 2003; Mariatos et al., 2003; Satoh et al., 2003;
Toyota et al., 2003; Erson and Petty, 2004), which is
much higher than the frequency of inactivation of all
spindle checkpoint genes combined.
Despite its potential importance in human cancer,
very little is known about the function of the CHFR
gene. The protein encoded by CHFR contains FHA and
RING domains and a cysteine-rich C-terminus. The
function of the C-terminal cysteine-rich region is
CHFR excludes Cyclin B1 from the nucleus
MK Summers et al
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a
Mitotic Index (%)
Morphology of CHFR-expressing cells exposed to
microtubule poisons
Characterizing CHFR-expressing cells whose cell cycle
progression is delayed in response to microtubule
poisons should help elucidate the function of CHFR at
the molecular level. Towards this goal, HCT116 cells,
which do not express endogenous CHFR protein, were
transfected with a bicistronic vector encoding hemaglutinin (HA)-tagged wild-type CHFR protein and a neoresistance marker or with the same vector expressing
only the neo-resistance marker. Pools of stably transfected CHFR and neo clones were then isolated. CHFR
protein was expressed in the CHFR clones at levels that
were modestly higher than the levels of endogenous
CHFR in SAOS2 cells (Figure 1a). Consistent with our
previous results (Scolnick and Halazonetis, 2000), 13 h
after exposure to nocodazole, two pools (#1 and #2) of
neo clones had a much higher fraction of cells with
condensed chromosomes compared to two pools of
CHFR-expressing clones. This was evident both by
monitoring chromosome condensation directly under
the microscope (Figure 1b) or by flow cytometric
analysis of cells staining positively for histone H3
phosphorylated on Ser10, which is a well-established
mitotic marker (Figure 1c). These results indicate that
the transfected CHFR gene was functioning in the stable
clones. Further, the two CHFR pools shown in Figure 1
Oncogene
R1
CHF
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CHFR
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Results
neo1
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SAO
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HCT
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H3 Ser10 P (%)
unclear; however, there has been some progress in
understanding the function of the FHA and RING
domains. Crystallographic evidence suggests that the
FHA domain of CHFR binds phosphorylated proteins
(Stavridi et al., 2002), as previously reported for other
FHA domains (Durocher et al., 1999). Biochemical
assays further suggest that the RING domain confers
ubiquitin ligase activity (Chaturvedi et al., 2002; Kang
et al., 2002; Bothos et al., 2003), consistent with previous
analyses of other proteins with RING domains (Joazeiro et al., 1999; Levkowitz et al., 1999; Lorick et al.,
1999). Many ubiquitin ligases target their substrates for
degradation in the proteasome. At least in Xenopus cellfree extracts, CHFR can mediate degradation of pololike kinase 1 (Plk1) leading to inhibition of Cdc25C and
Cdc2 (Kang et al., 2002). However, other in vitro studies
suggest that CHFR is a noncanonical ubiquitin ligase
(Bothos et al., 2003). Noncanonical ubiquitin ligases do
not target proteins for degradation, but rather utilize
their ubiquitin ligase activity for signal transduction
(Pickart, 2001).
Since the original publication on CHFR, very little
work has been carried out describing its effects on
human cells. Here, we studied cell lines that do not
express CHFR and stably transfected derivatives that
express wild-type CHFR, and characterize the CHFRinduced delay. Our findings suggest that CHFR delays
cells in early prophase by inhibiting entry of Cyclin B1
in the nucleus.
10
Nocodazole
13 h
Figure 1 Characterization of pools of stably transfected HCT116
neo- and CHFR-expressing clones. (a) CHFR protein expression in
HCT116 neo and CHFR stably transfected cells (pools #1).
Extracts of SAOS2 osteosarcoma cells were included to show the
levels of endogenous CHFR. (b) Mitotic index of stably transfected
HCT116 neo- and CHFR-expressing cells (pools #1 and #2) at
different times after exposure to nocodazole. DAPI-stained cells
were scored for chromosome condensation. Means and standard
errors of three independent experiments are shown. (c) Fraction of
cells scoring positive for histone H3 Ser10 phosphorylation. Stably
transfected HCT116 neo- and CHFR-expressing cells (pools #1 and
#2) were exposed to nocodazole, and H3 Ser10 phosphorylation
was monitored by flow cytometry
behaved identically to each other, as did the two neo
pools, prompting us to focus our characterization of the
effects of CHFR expression on one neo (#1) and one
CHFR (#1) pool.
The morphology of the CHFR and neo clones after
exposure to microtubule poisons was monitored by
DAPI staining and immunofluorescent staining of the
lamin cytoskeleton (Figure 2 and data not shown).
Many of the neo cells had condensed chromosomes and
diffuse lamin staining indicating nuclear envelope
breakdown, whereas the few cells that had not yet
entered metaphase showed a typical interphase morphology with intact lamin cytoskeleton. The majority of
the CHFR-expressing cells did not have condensed
chromosomes or nuclear envelope breakdown, and also
did not exhibit a classical interphase morphology;
instead, the outline of the nuclear envelope was ruffled
CHFR excludes Cyclin B1 from the nucleus
MK Summers et al
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Nocodazole
HCT116 − neo
Paclitaxel
HCT116
neo
HCT116
CHFR
IR
6h
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9h
HCT116 − CHFR
:20
:40
6h
7h
8h
9h
:20
:40
10h
:20
:40
11h
:20
:40
12h
:20
:40
13h
*
*
*
*
*
*
*
Figure 2 Deformation of the nuclear envelope of CHFR-expressing HCT116 cells delayed in the cell cycle after exposure to
microtubule poisons. Neo- and CHFR-expressing cells were
exposed to ionizing radiation (IR), nocodazole or paclitaxel. After
13 h, the cells were fixed and stained for Lamins A/C. The cells
marked with an asterisk are in prometaphase
(Figure 2). To determine whether this morphology was
specific to cell cycle delay induced in response to
microtubule poisons, we also examined cells 13 h after
exposure to ionizing radiation (IR). In response to IR,
both the neo- and CHFR-expressing cells had a smooth
nuclear envelope outline typical of cells arrested in G2
(Figure 2).
To better visualize the ruffled nuclear envelope
morphology of CHFR-expressing cells exposed to
microtubule poisons, we generated cells that, in addition
to CHFR or neo, also expressed histone H2B fused to
green fluorescent protein (GFP) (Kanda et al., 1998).
These cells were synchronized at the G1/S boundary
with a single thymidine block, released into S phase and
challenged with nocodazole 4 h later. Chromosome
condensation was monitored by live cell time-lapse
microscopy. The neo cells showed a smooth outline of
the nuclear envelope during S and G2 and then, about
9–10 h after release from the G1/S block, condensed
their chromosomes (Figure 3). By comparison, the
CHFR-expressing cells did not condense their chromosomes until approximately 12–13 h after release from the
G1/S block. For the first 9–10 h, these cells had a
smooth nuclear envelope outline, but then adopted a
ruffled outline, which persisted until the chromosomes
condensed (Figure 3).
In the absence of microtubule poisons, cells in G2
have a smooth outline of the nuclear envelope. However, as they progress through prophase, the nuclear
envelope shows symmetrical indentations, which are
thought to be initiated by microtubules penetrating a
partially depolymerized lamin cytoskeleton (Georgatos
et al., 1997; Beaudouin et al., 2002; Salina et al., 2002).
The ruffled nuclear envelope morphology of the CHFRexpressing cells exposed to microtubule poisons cannot
be due to penetrating microtubules, but could indicate
partial depolymerization of the lamin cytoskeleton. In
turn, this would indicate that these cells were past G2,
Figure 3 Deformation of the nuclear envelope of CHFR-expressing HCT116 cells delayed in the cell cycle after exposure to
microtubule poisons. Neo- and CHFR-expressing cells that also
express a histone H2B-GFP fusion protein were synchronized in
G1/S and then released into the cell cycle (0 h time point).
Nocodazole was added 4 h later. Images were captured every
20 min starting 6 h after release from the cell cycle block.
Representative cells are shown
yet not past mid prophase, since the chromosomes were
not condensed.
Molecular markers in CHFR-expressing cells exposed to
microtubule poisons
Analysis of molecular markers can be used to define the
phase of the cell cycle in which the CHFR-expressing
cells are delayed in response to microtubule poisons, as
well as to begin to elucidate the function of CHFR at the
molecular level. We first examined the Cyclin A/Cdc2
and Cyclin B1/Cdc2 complexes in the neo- and CHFRexpressing cells. Synchronized cells were exposed to
nocodazole and at various time points thereafter cell
extracts were prepared for immunoblotting analysis and
kinase activity assays or the cells were fixed for analysis
by immunofluorescence. Since CHFR induces a transient mitotic delay, whose duration can vary from cell to
cell, the population of synchronized cells we studied was
not homogeneous: some cells were being delayed in the
cell cycle with uncondensed chromosomes, while others
had overcome the CHFR checkpoint and had condensed
their chromosomes (late prophase and prometaphase
cells). Thus, before preparing extracts the two populations of cells were separated by shake-off and extracts
were prepared separately for each population. The neoexpressing cells were similarly separated into adherent
and nonadherent populations.
We first examined Cyclin B1/Cdc2 kinase activity
using histone H1 as a substrate (Figure 4a). In the
adherent CHFR-expressing cells, Cyclin B1/Cdc2 was
catalytically inactive, whereas in the nonadherent cells
Cyclin B1/Cdc2 activity was high. The low Cyclin B1/
Cdc2 activity in some adherent cell fractions could be
explained by low level contamination of these fractions
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CHFR excludes Cyclin B1 from the nucleus
MK Summers et al
2592
10 12 12
Ad
NA
c
HCT116−CHFR
11.5 h
Ad NA Ad NA
Ad
10
81 0 12 12 h
pTP
42 Kd
Cdc25C
42 Kd
M
Cdc2
pTP
32
Cdc2
NM
M
NM
M
pTP
MI 3
MI 11 15
NM
3
12
28 Kd
24 64
6
7
HCT116−CHFR
10
68
P−H1
8
NA
b
55 14 82 %
11.5 h
Ad
−
NA
Cdc2
Ad
HCT116−CHFR
NM
M
CycA
HCT116−neo
d
Cyclin B1
DAPI
HCT116
CHFR
a
16 67 %
Figure 4 Analysis of Cyclin A and Cyclin B1/Cdc2 in HCT116 neo- and CHFR-expressing cells exposed to nocodazole. The cells were
synchronized in G1/S, released into the cell cycle (0 h) and exposed to nocodazole (4 h later). Prior to preparing cell extracts, the G2/
early prophase (adherent, Ad) cells were separated from the prometaphase (nonadherent, NA) cells by mitotic shake-off. The efficiency
of cell separation was monitored by measuring the mitotic index (MI) of a small aliquot of DAPI-stained cells. (a,b) Cyclin B1/Cdc2
activity was measured directly by monitoring phosphorylation of histone H1 (32P-H1). Cdc2 and Cdc25C electrophoretic migration was
monitored by immunoblotting; M and NM indicate the predominant forms of Cdc2 and Cdc25C in mitotic and nonmitotic cells,
respectively. The presence of phosphorylated Cdc2 substrates was monitored by immunoblotting with an antibody that recognizes
phosphorylated threonine followed by a proline (pTP). (c) Electrophoretic migration of Cdc2 associated with Cyclin A in CHFRexpressing cells exposed to nocodazole. Cyclin A/Cdc2 complexes were immunoprecipitated with antibodies that recognize Cyclin A;
the immunoprecipitates were blotted for Cyclin A and Cdc2. (d) Intracellular localization of Cyclin B1 in CHFR-expressing HCT116
cells exposed for 13 h to nocodazole
with cells that had condensing or condensed chromosomes (as shown by the mitotic index (MI) in Figure 4a).
Thus, these findings suggest that in CHFR-expressing
cells, Cyclin B1/Cdc2 is inactive during the checkpointinduced cell cycle delay. This conclusion was further
supported by examining the electrophoretic migration of
Cdc2. A slowly migrating form of Cdc2, representing
inactive Cdc2 doubly phosphorylated on Thr14 and
Tyr15, was present in the adherent CHFR-expressing
cells, which had not yet initiated chromosome condensation, but was absent in the nonadherent (prometaphase/metaphase) cells (Figure 4a).
The low Cyclin B1/Cdc2 activity in CHFR-expressing
cells during the checkpoint-induced cell cycle delay was
somewhat inconsistent with the ruffled nuclear envelope
morphology of these cells, since lamin depolymerization
is thought to be mediated by Cdc2 (Ward and
Kirschner, 1990; Peter et al., 1991). Thus, to further
explore Cdc2 activity, we immunoblotted the cell
extracts with an antibody that recognizes phosphorylated threonine followed by a proline (Figure 4a). This
phosphospecific antibody reacts with Cdc2 substrates,
and also with substrates of Cdk2 and other kinases. In
the neo-expressing cells, at least two bands, corresponding to molecular weights of 28 and 42 kDa, respectively,
were detected specifically in the prometaphase/metaphase cells (nonadherent population, Figure 4a), suggesting that these bands are Cdc2 substrates.
Surprisingly, in the CHFR-expressing cells at the 12 h
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timepoint the same proteins were phosphorylated in
both the adherent and nonadherent cells (Figure 4a and
b). The phosphorylation of these proteins in the
adherent cells could not be attributed to gross contamination by prometaphase/metaphase cells, as indicated
by the low MI and the electrophoretic migration of
Cdc2 and Cdc25C (Figure 4a and b). Further, the
phosphorylation was unlikely to be mediated by Cyclin
B1/Cdc2, based on the kinase activity assays. This left us
with the possibility that the phosphorylation events
recognized by the phospho-Thr-Pro-specific antibody
were mediated by Cyclin A/Cdc2. We could not test
directly for Cyclin A/Cdc2 kinase activity by immunoprecipitating Cyclin A-associated kinases, because Cyclin A associates with both Cdc2 and Cdk2. However,
we could study the electrophoretic migration of Cdc2
that coimmunoprecipitates with Cyclin A as an indirect
indicator of Cyclin A/Cdc2 activity. At the 10 h time
point, when CHFR-expressing cells show little reactivity
with the phospho-Thr-Pro-specific antibody, the fraction of Cdc2 that coimmunoprecipitated with Cyclin A
migrated as a doublet with the higher band representing
Cdc2 phosphorylated on both Thr14 and Tyr15. At the
11.5 h time point, when the adherent CHFR-expressing
cells showed reactivity with the phospho-Thr-Prospecific antibody, the fraction of Cdc2 that coimmunoprecipitated with Cyclin A migrated mostly as a single
band representing Cdc2, in which Thr14 and Tyr15 were
dephosphorylated (Figure 4c). This result, which corre-
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MK Summers et al
2593
a
8
11.5
10
13 h
HCT116−
CHFR neo
Ad NA Ad NA Ad NA
Plk1
Plk1
γ − tubulin
DAPI
Plk1
HCT116
− neo
HCT116
− CHFR
b
c
HCT116 − neo
Ad
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NA
10 12 12
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NA
10 12 12 h
AurA
γ − tubulin
AurA
DAPI
HCT116
− neo
HCT116
− CHFR
d
e
γ − tubulin
AurA pT288
DAPI
*
HCT116
− CHFR
lates with the appearance of phospho-Thr-Pro-reactive
bands (Figure 4b), suggests that, unlike Cyclin B1/Cdc2,
Cyclin A/Cdc2 is active in CHFR-expressing cells during
the checkpoint-induced cell cycle delay. Interestingly, in
the nonadherent cell fraction (prometaphase/metaphase
cells), Cdc2 was not associated with Cyclin A
(Figure 4c), indicating that after chromosome condensation Cdc2 associates exclusively with Cyclin B1.
In addition to examining Cyclin/Cdc2 complexes in
cell extracts, we also monitored the intracellular
localization of Cyclins A and B1 by immunofluorescence. Cyclin A was predominantly nuclear in both the
neo- and CHFR-expressing cells (data not shown).
Cyclin B1, which is known to translocate from the
cytoplasm to the nucleus in mid prophase and whose
nuclear localization is required for chromosome condensation (Pines and Hunter, 1991), was cytoplasmic in
the CHFR-expressing cells during the checkpointinduced cell cycle delay (Figure 4d).
Entry of Cyclin B1 in the nucleus is regulated at least
in part by phosphorylation of residues surrounding its
nuclear export sequence. The polo-like kinase Plk1
phosphorylates some of these residues (Li et al., 1997;
Hagting et al., 1998, 1999; Toyoshima et al., 1998; Yang
et al., 1998, 2001; Toyoshima-Morimoto et al., 2001),
raising the possibility that CHFR might sequester Cyclin
B1 in the cytoplasm by inhibiting Plk1. In support of
this model, it is known that CHFR encodes a ubiquitin
protein ligase, which at least in Xenopus extracts can
target Plk1 for ubiquitin-dependent proteolysis (Kang
et al., 2002). We therefore examined whether Plk1
protein levels might decrease in CHFR-expressing cells
exposed to nocodazole. The cell extracts prepared from
the adherent and nonadherent populations of synchronized neo- and CHFR-expressing cells were immunoblotted with Plk1-specific antibodies. There was no
difference in Plk1 protein levels in response to CHFR
expression (Figure 5a), indicating that the CHFRmediated cell cycle delay was not associated with a
decrease in overall Plk1 protein levels.
We further considered the possibility that Plk1 might
be degraded at specific intracellular compartments, in
which case total Plk1 protein levels might not be
significantly affected. To explore this possibility, we
monitored by immunofluorescence the intracellular
localization of Plk1 in synchronized neo- and CHFRexpressing cells exposed to microtubule poisons. Again
we noted no difference between the two cell types. In the
CHFR-expressing cells that were arrested in early
prophase and in the neo-expressing cells that had not
yet entered prometaphase, Plk1 was present at the
centrosomes and in the nucleus in a kinetochore-like
pattern (Figure 5b).
A second kinase that regulates the intracellular
localization of Cyclin B1 is Aurora-A (Hirota et al.,
2003). We therefore monitored Aurora-A protein levels,
intracellular localization and phosphorylation state in
synchronized neo- and CHFR-expressing cells exposed
to microtubule poisons. In extracts prepared from the
adherent and nonadherent populations, there was no
difference in Aurora-A protein levels as a function of
Figure 5 Analysis of Plk1 and Aurora-A in HCT116 neo- and
CHFR-expressing cells exposed to nocodazole. The cells were
synchronized in G1/S, released into the cell cycle (0 h), exposed to
nocodazole (4 h) and examined 8–13 h after release from the cell
cycle block, as indicated. (a,c) Plk1 and Aurora-A (Aur A) protein
levels. G2/early prophase (adherent, Ad) cells were separated from
prometaphase (nonadherent, NA) cells by mitotic shake-off before
the extracts were prepared. (b,d,e) Immunofluorescence analysis for
Plk1, Aurora-A (AurA) and Aurora-A phosphorylated on Thr288
12 h after release from the G1/S block. The centrosomes (arrowheads) were marked by staining for g-tubulin. In (e),the cell marked
with an asterisk is in prometaphase
CHFR expression (Figure 5c). By immunofluorescence,
Aurora-A was present at the centrosomes of practically
every cell (the synchronized neo- and CHFR-expressing
cells were in late G2, prophase and prometaphase),
Oncogene
CHFR excludes Cyclin B1 from the nucleus
MK Summers et al
suggesting that CHFR does not inhibit accumulation of
Aurora-A at centrosomes (Figure 5d). We further
examined whether centrosomal Aurora-A was phosphorylated on Thr288, a residue that resides in the
activation loop and whose phosphorylation state
parallels Aurora-A activation (Hirota et al., 2003). In
the CHFR-expressing cells delayed prior to chromosome
condensation, centrosomal Aurora-A was not phosphorylated on Thr288; but phosphorylation was evident
in cells that had overcome the checkpoint and condensed their chromosomes (Figure 5e). Thus, CHFRinduced cell cycle delay is associated with the presence of
inactive Aurora-A at the centrosomes.
Aurora-B localizes to kinetochores in G2, prophase
and metaphase and phosphorylates histone H3 on Ser10
in mitosis (Hsu et al., 2000; Adams et al., 2001; Giet and
Glover, 2001). The absence of histone H3 phosphorylation on Ser10 in cells whose progression was delayed by
CHFR (Figure 1c) implies that Aurora-B is inactive in
these cells. By immunoblotting and immunofluorescence, we did not detect decreased Aurora-B protein
levels in cells delayed in the cell cycle by CHFR (data
not shown), suggesting that CHFR does not target
Aurora-B for protein degradation.
Forced expression of Cyclin B1 in the nucleus overcomes
the CHFR checkpoint
Inspection of more than a thousand synchronized
CHFR-expressing cells exposed to nocodazole failed to
identify any cells with cytoplasmic Cyclin B1 that were
initiating chromosome condensation or stained positive
for histone H3-Ser10 phosphorylation. However, we
were able to identify several cells that had Cyclin B1 in
the nucleus and were initiating chromosome condensation and/or exhibited phosphorylation of histone H3 on
Ser10 (Figure 6a). From this we infer that nuclear
accumulation of Cyclin B1 precedes chromosome
condensation and histone H3-Ser10 phosphorylation
and may be required for overcoming the cell cycle delay
induced by CHFR in response to microtubule poisons.
To test this hypothesis, we examined whether targeting
Cyclin B1 to the nucleus could overcome the CHFRmediated cell cycle delay.
The nuclear export sequence (NES) of Cyclin B1 can
be inactivated by substituting Phe146 with Ala (Hagting
et al., 1999). The Cyclin B1 A146 mutant or wild-type
Cyclin B1, as a control, were stably introduced in
SAOS2 cells, which have an intact CHFR checkpoint
(Scolnick and Halazonetis, 2000). The Cyclin B1 A146
mutant overcame the CHFR-induced cell cycle delay,
whereas expression of wild-type Cyclin B1 had a smaller
effect (Figure 6b), even though its level of expression
was similar to that of the A146 mutant (Figure 6c). A
dominant negative CHFR mutant that lacks the FHA
domain partially overcame the delay, consistent with
our previous findings (Scolnick and Halazonetis, 2000).
Since expression of the Cyclin B1 A146 mutant in other
cell systems is not sufficient to drive premature entry
into mitosis (Jin et al., 1998; Toyoshima et al., 1998;
Hagting et al., 1999; Yang et al., 2001), we propose that
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a
Cyclin B1
H3 Ser10 P
b
DAPI
1
2
H3 Ser10 P (%)
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20
15
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c
neo
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neo CHFR wt A146
∆FHA Cyclin B1
Cyclin B1 CHFR
wt A146 ∆FHA
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Cyclin
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Figure 6 Entry of Cyclin B1 in the nucleus overcomes the CHFRinduced delay. (a) Nuclear localization of Cyclin B1 is always
accompanied by histone H3 phosphorylation on Ser10. CHFRexpressing HCT116 cells were released from a G1/S cell cycle block
(0 h), exposed to nocodazole (4 h) and processed for immunofluorescence (12 h). The cells numbered ‘1’ and ‘3’ have nuclear Cyclin
B1 and stain positive for H3 phosphorylation on Ser10. DAPI
staining further shows that cell ‘3’ has initiated chromosome
condensation. (b,c) Expression of Cyclin B1 with a mutant nuclear
export sequence overcomes the CHFR-induced cell cycle delay. (b)
Stably transfected SAOS2 clones expressing a neo vector with no
other insert or with inserts expressing a dominant negative CHFR
mutant (CHFRDFHA), wild-type Cyclin B1 or Cyclin B1 A146, all
with an HA tag, were released from a G1/S cell cycle block (0 h),
exposed to nocodazole (8 h) and then examined for H3 Ser10
phosphorylation by flow cytometry (14 h). Each column represents
the mean and standard error from two independent pools of stably
transfected clones. (c) Levels of ectopically expressed HA-tagged
proteins in the stably transfected SAOS2 pools (two per insert)
assayed by immunoblotting with an antibody that recognizes the
HA tag
in our system the Cyclin B1 146 mutant overrides the
CHFR-induced delay.
Discussion
CHFR delays chromosome condensation in cells exposed to microtubule poisons (Scolnick and Halazonetis, 2000). The original findings were consistent with cell
cycle delay either in late G2 or in early prophase. In this
study, we suggest that the CHFR-induced delay occurs
within early prophase, rather than in late G2. This
conclusion stems from analysis of synchronized nocodazole-treated live cells expressing neo or CHFR, which
shows that at the time the neo-expressing cells condensed their chromosomes, the CHFR-expressing cells
deformed their nuclear envelope (Figure 3). In the
absence of microtubule poisons, deformation of the
nuclear envelope occurs in prophase, is associated with
penetrating microtubules from the maturing centrosomes and is quickly followed by nuclear envelope
breakdown (Georgatos et al., 1997; Schneider et al.,
1999; Salina et al., 2002). Under these conditions,
deformation of the nuclear envelope is thought to reflect
partial depolymerization of the lamin cytoskeleton,
which in turn is dependent on Cdc2 activity (Ward
and Kirschner, 1990; Peter et al., 1991). By analogy, we
interpret the deformation of the nuclear envelope in
CHFR-expressing cells exposed to nocodazole to
CHFR excludes Cyclin B1 from the nucleus
MK Summers et al
2595
indicate that the lamin cytoskeleton was partially
depolymerized and that the cells were delayed past G2
in prophase. Further, because the CHFR-delayed cells
had not initiated chromosome condensation, the delay
would have to be in early, rather than late, prophase.
A recent study has also characterized CHFR-delayed
cells and concluded that the cells were delayed in
antephase (Matsusaka and Pines, 2004). Antephase is a
rarely used term, originally introduced in 1951 (Bullough and Johnson, 1951), to define a phase of the cell
cycle in late G2 just before chromosomes begin to
condense, when normal mouse epidermal cells are
reversibly delayed when subjected to various stresses.
Those, who use the term antephase, consider prophase
synonymous with initiation of chromosome condensation. We have used the term early prophase to define
cells that are past G2, but have not yet initiated
chromosome condensation. Cells in early prophase can
be distinguished from cells in G2, because they have
initiated centrosome movement along the periphery of
the nuclear membrane, which is the earliest described
mitotic event visible under the microscope (McIntosh
and Koonce, 1989). From the definitions of antephase
and early prophase listed above, it follows that these two
terms may refer to the same phase of the cell cycle. We
will continue to use the term early prophase here, but
remain open to adjusting our terminology, as consensus
among investigators in the field of mitosis develops.
We believe that a key finding of our study is the
identification of active Cyclin A/Cdc2 in CHFR-delayed
cells (Figure 7). This finding was principally based on
the electrophoretic mobility of Cdc2 associated with
Cyclin A (Figure 4c). Even though Cdc2 electrophoretic
mobility is an indirect marker of kinase activity, several
other observations support the conclusion that Cyclin
A/Cdc2 is active in CHFR-delayed cells. Specifically, the
CHFR-delayed cells show deformation of the lamin
cytoskeleton, Plk1 localization at kinetochores and
phospho-Thr-Pro reactive proteins (Figure 7). The
former two features are known to require Cdc2 activity
(Ward and Kirschner, 1990; Peter et al., 1991; Elia et al.,
2003) and the same is likely to be true for the latter.
Since Cyclin B1/Cdc2 was inactive in CHFR-delayed
cells, these results are consistent with Cyclin A/Cdc2
being active.
Early Prophase
Mid Prophase
CHFR
Plk1 at
Kinetochores
Active
Cyclin A
/Cdc2
Deformation of
Lamin Cytoskeleton
pThrPro Antibody
Reactive Proteins
Active
Aurora B
H3 Ser10
Phosphorylation
Active
Aurora A
and
Plk1
Nuclear
Cyclin B1
Chromosome
Condensation
Active
Plk1
Active
Cdc25C
Active
Cyclin B1
/Cdc2
Nuclear
Envelope
Breakdown
Figure 7 Model for role of CHFR in regulating early mitotic
events. CHFR is shown to delay the transition from early to mid
prophase by keeping Aurora-A, Aurora-B, Plk1 and Cyclin B1/
Cdc2 inactive. However, the direct molecular target of the CHFR
protein remains unknown. See text for more details
Interestingly, another group recently concluded that
Cyclin A/Cdks are inactive in CHFR-delayed cells. This
group did not examine Cyclin A/Cdk2 or Cyclin A/Cdc2
activity or post-post-translational modifications, but
reached their conclusion by showing that injection of
Cyclin A in complex with a constitutively active Cdk2
mutant overcame the CHFR-induced delay (Matsusaka
and Pines, 2004). This result implies that even Cyclin A/
Cdk2 is inactive in CHFR-delayed cells. However, in our
opinion, this result cannot substitute for direct analysis
of endogenous Cyclin A/Cdks. While it would be
possible for Cyclin A/Cdc2 to be inactive in CHFRdelayed cells, it is particularly hard to imagine that
Cyclin A/Cdk2 would be inactive, given that this
complex is activated in S phase (Woo and Poon, 2003).
Unlike Cyclin A/Cdc2, Cyclin B1/Cdc2 was inactive
in CHFR-delayed cells (Figure 7). This became apparent
thanks to an important technical improvement from our
previous study (Scolnick and Halazonetis, 2000) that
allowed us to separate CHFR-delayed cells from cells
that had entered prometaphase. The molecular basis for
low Cyclin B1/Cdc2 activity in CHFR-delayed cells
might be related to cytoplasmic sequestration of Cyclin
B1; both in this study and in a previous study (Satoh
et al., 2003), Cyclin B1 was shown to be in the cytoplasm
of CHFR-expressing cells. We established a link between
cytoplasmic sequestration of Cyclin B1 and CHFRinduced delay by showing that ectopic expression of a
Cyclin B1 mutant that constitutively localizes to the
nucleus overcame the CHFR-induced delay. High levels
of Cyclin B1 in the nucleus could lead to activation of
Cyclin B1/Cdc2 via a positive feedback loop that
involves phosphorylation and activation of Cdc25C by
Cyclin B1/Cdc2 and subsequent dephosphorylation and
activation of Cyclin B1/Cdc2 by Cdc25C (Strausfeld
et al., 1994; Karlsson et al., 1999). We therefore propose
that the mechanism by which CHFR delays progression
through prophase involves regulation of the intracellular
localization of Cyclin B1.
By immunoblot analysis, we did not observe any
change in Cyclin B1 levels or electrophoretic migration
in CHFR-delayed cells that would indicate that CHFR
directly ubiquitinates Cyclin B1 (data not shown).
Therefore, CHFR is more likely to regulate Cyclin B1
indirectly. Cyclin B1 has both nuclear localization and
nuclear export sequences. Phosphorylation of serines
126, 128, 133 and 147 adjacent to the nuclear export
sequence leads to accumulation of Cyclin B1 in the
nucleus (Li et al., 1997; Hagting et al., 1998, 1999;
Toyoshima et al., 1998; Yang et al., 1998, 2001;
Toyoshima-Morimoto et al., 2001). The kinase or
kinases that phosphorylate Ser126 and Ser128 of Cyclin
B1 have not been identified, although Cdc2 appears to
be a good candidate. Interestingly, phosphorylation of
Ser147 and/or Ser133 is mediated by Plk1 (ToyoshimaMorimoto et al., 2001), which at least in vitro is a
substrate of CHFR (Kang et al., 2002). This raises the
possibility that CHFR inhibits accumulation of Cyclin
B1 in the nucleus by targeting Plk1 for degradation.
However, by immunoblotting, we did not observe a
significant change in protein levels of endogenous Plk1
Oncogene
CHFR excludes Cyclin B1 from the nucleus
MK Summers et al
2596
in the CHFR-delayed cells compared to the neo controls
(Figure 5a) and similar results were recently reported by
others (Erson and Petty, 2004; Matsusaka and Pines,
2004). CHFR expression also did not affect the Plk1
immunofluorescence signal at the centrosomes or kinetochores (Figure 5b), thus excluding the possibility that
CHFR promotes degradation of Plk1 at specific cellular
compartments. Aurora-A is another kinase that regulates the nuclear translocation of Cyclin B1 in mid
prophase (Hirota et al., 2003). The mechanism is unclear,
but requires activation of the pool of Aurora-A at the
centrosomes. CHFR expression did not affect the overall
protein levels of Aurora-A or its immunofluorescence
signal at the centrosomes (Figure 5). Thus, like Plk1,
Aurora-A was not targeted for degradation by CHFR.
CHFR is the most frequently inactivated mitotic
checkpoint gene in human cancer, but its molecular
function is poorly understood. Our study establishes that
CHFR delays cells in early prophase, provides morphological criteria to identify CHFR-delayed cells, and examines
the status of key regulators of mitosis, such as Cyclin A/
Cdc2, Cyclin B1/Cdc2, Plk1, Aurora-A and Aurora-B
(Figure 7). The identification of Cyclin B1 as an indirect
target of CHFR is likely to help dissect the molecular
function of this important mitotic checkpoint protein.
Materials and methods
Cell lines, antibodies and microtubule poisons
SAOS2 osteosarcoma and HCT116 colon carcinoma cells
express wild-type CHFR protein or no detectable CHFR
protein, respectively (Scolnick and Halazonetis, 2000). Antibodies to cell cycle markers were obtained from commercial
vendors, as follows: Plk1 and H3 phosphorylated on Ser 10
from Upstate Biothechnology; Cdc2, Aurora-A, Aurora-B,
Cyclin A and Cyclin B1 from Transduction Labs; Lamins A/C
and B from Santa Cruz; Aurora-A phosphorylated on Thr288
and phospho-Thr-Pro from Cell Signaling Technologies.
Mitotic stress was induced by adding nocodazole (1 mM) or
paclitaxel (5 mM) to the tissue culture media. DNA damage was
induced by exposing the cells to 5 Gy IR using a 137Cs
irradiator.
Stably transfected cell clones
HCT116 and SAOS2 cells were used to generate pools of
stably transfected clones using a pIRESN2 bicistronic plasmid
(Clontech Laboratories) containing various inserts, as indicated. Colonies of stably transfected cells were selected with
G418, expanded and screened by immunoblotting for expression of the protein encoded by the insert.
Cell cycle synchronization and mitotic shake-off
For cell cycle synchronization, HCT116 and SAOS2 cells were
incubated with 1.3 mM thymidine for 18 and 20 h, respectively,
then washed 2 with PBS (Life Technologies) and released
into media containing 250 mM thymidine and 250 mM cytidine.
Nocodazole was added 4–8 h later, as indicated. For separation
of G2/prophase from prometaphase cells, the tissue culture
plates were pulsed on a vortexer. The supernatant was collected
and combined with a PBS wash (prometaphase cells). The
attached cells (G2/prophase) were collected by trypsinization.
Oncogene
Live cell analysis
Pools of stably transfected HCT116 cells were generated, as
described above, except that a plasmid expressing histone H2B
fused to GFP was also cotransfected. The cells were
synchronized at the G1/S boundary; 4 h after release from
the cell cycle arrest, fresh media was added consisting of
DMEM without phenol red, 10% fetal calf serum (FCS),
25 mM HEPES (pH 7.4) and 1 mM nocodazole. Then, the cells
were placed on a microscope (Leica) equipped with water
immersion optics and a Delta-T4 heating system for the stage
and objective lens (Bioptechs). Images were acquired every
20 min starting 6 h after release from the cell cycle block using
an ORCA ER digital camera (Hamamatsu) and processed
using Imagevision tools (SGI).
Mitotic index
For determination of the mitotic index, the cells were fixed in
2% paraformaldehyde for 15 min at room temperature (RT),
stained with DAPI and mounted on glass slides using a
cytospin centrifuge (Shandon). The mitotic index, which
represents the fraction of cells with fully condensed chromosomes, was determined by inspecting per data point more than
200 cells.
Flow cytometry analysis for histone H3 Ser10 phosphorylation
For determination of the fraction of cells with histone H3
Ser10 phosphorylation, the cells were fixed in 0.1% paraformaldehyde for 15 min at RT, washed and resuspended in icecold 70% EtOH for at least 10 min. The cells were then
extracted on ice with 0.2% Triton X-100 for 5 min, washed
with PBS, blocked with 10% heat-inactivated FCS in PBS for
30 min at RT and incubated overnight at 41C with an antibody
that recognizes H3 phosphorylated on Ser10 diluted 1 : 200 in
1% BSA/PBS. The cells were then washed with PBS, incubated
for 30 min at RT with FITC-conjugated goat-anti-rabbit
antibody (Vector Laboratories) diluted 1 : 200 in 1% BSA/
PBS, washed again and resuspended in PBS supplemented with
0.1% Tween-20, 2% FCS, 5 ng/ml RNAse and 5 mg/ml
propidium iodide. After 1 h incubation at 371C, the cells were
analysed by flow cytometry (Becton Dickinson).
Immunofluorescence
The cells were seeded on 12-mm autoclaved glass coverslips
and allowed to adhere for at least 2 days. After the indicated
experimental treatments, the cells were washed once with PBS,
fixed in 3.7% paraformaldehyde/2% sucrose in PBS for 15 min
at RT or in methanol at 201C for 20 min and permeabilized
with 0.5% Triton X-100 in PBS on ice for 20 min. After two
washes in PBS, coverslips were blocked for 30 min at RT with
2.5% gelatin in PBS. Primary antibodies were diluted in 1%
BSA/PBS and incubated at RT for 2 h or overnight at 41C. The
coverslips were then washed with PBS supplemented with
0.1% Tween-20 and incubated for 30 min at RT with the
appropriate secondary antibody conjugated to Alexafluor 488
or 568 (Molecular Probes) at a dilution of 1 : 500 in 1% BSA/
PBS. After washing, the cells were counterstained with DAPI
and mounted with Fluorimount G (Southern Biotechnology
Associates).
Immunoblotting and immunoprecipitations
Cells were lysed with buffer consisting of 50 mM Tris (pH 8.0),
120 mM NaCl, 0.5% NP40, 1 mM DTT, 0.4 mg/ml Pefabloc SC,
2 mg/ml pepstatin, 1 mM Calyculin A, 1 mM okadaic acid, 15 mM
CHFR excludes Cyclin B1 from the nucleus
MK Summers et al
2597
NaF and 1 mM sodium vanadate. The lysates were cleared by
centrifugation, resolved by gel electrophoresis and immunoblotted with antibodies that recognize a panel of cell cycle
proteins. For immunoprecipitation experiments, whole-cell
lysates were diluted to 0.5 ml with lysis buffer and precleared
with 30 ml protein G beads (Pharmacia) for 30 min at 41C. The
supernatants were incubated with primary antibodies for 1 h at
41C and the immunocomplexes were captured on protein G
beads. After washing off the unbound proteins with lysis buffer,
the bound proteins were immunoblotted, as described above.
Cyclin B1/Cdc2 kinase assay
Cyclin B1/Cdc2 was immunoprecipitated from 50 mg wholecell lysate as described above using a monoclonal antibody
that recognizes Cyclin B1. The beads were washed 3x with lysis
buffer and then 2x with kinase buffer (50 mM HEPES (pH 7.4),
150 mM NaCl, 5 mM MgCl2, 1 mM DTT). The beads were then
resuspended in 30 ml kinase buffer supplemented with 5 mg
histone H1, 1 mCi 32P-g-ATP and 5 mM MnCl2. After incubation at 301C for 30 min, the reaction products were resolved by
SDS gel electrophoresis and kinase activity was monitored by
fluorography.
Acknowledgements
We thank G Wahl for the gift of plasmid expressing GFP fused
to histone H2B and Daniel Scolnick for helpful discussions.
Financial support was provided by the National Cancer
Institute (CA89630). MKS and JB were supported by the
National Cancer Institute Training Grants CA09677 and
CA09171, respectively.
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