Download Loss of p38 MAPK induces pleiotropic mitotic defects and massive

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Research Article
Loss of p38 MAPK induces pleiotropic mitotic defects
and massive cell death
Anu Kukkonen-Macchi1,2, Oana Sicora1, Katarzyna Kaczynska1, Christina Oetken-Lindholm1,
Jeroen Pouwels1,2, Leena Laine2 and Marko J. Kallio1,2,*
1
Turku Centre for Biotechnology, University of Turku, 20521 Turku, Finland
VTT Technical Research Centre of Finland, Medical Biotechnology, 20521 Turku, Finland
2
*Author for correspondence ([email protected])
Accepted 12 September 2010
Journal of Cell Science
Journal of Cell Science 124, 216-227
© 2011. Published by The Company of Biologists Ltd
doi:10.1242/jcs.068254
Summary
The p38 mitogen-activated protein kinase (p38 MAPK) family, which is comprised of four protein isoforms, p38, p38, p38 and
p38, forms one of the key MAPK pathways. The p38 MAPKs are implicated in many cellular processes including inflammation,
differentiation, cell growth, cell cycle and cell death. The function of p38 MAPKs in mitotic entry has been well established, but their
role in mitotic progression has remained controversial. We identify p38 MAPK as a modulator of mitotic progression and mitotic cell
death. In HeLa cells, loss of p38 results in multipolar spindle formation and chromosome misalignment, which induce a transient M
phase arrest. The majority of p38-depleted cells die at mitotic arrest or soon after abnormal exit from M-phase. We show that p38
MAPKs are activated at the kinetochores and spindle poles throughout mitosis by kinase(s) that are stably bound to these structures.
Finally, p38 is required for the normal kinetochore localization of polo-like kinase 1 (Plk1), and this contributes to the activity of the
p38 MAPK pathway. Our data suggest a link between mitotic regulation and the p38 MAPK pathway, in which p38 prevents
chromosomal instability and supports mitotic cell viability.
Key words: p38 MAPK, p38, Mitosis, Cell death, Kinetochore
Introduction
The mitogen-activated protein kinases (MAPKs) work in
fundamental cellular processes and participate in signaling
networks composing four distinct pathways: the extracellular signalregulated kinases (ERK) pathway, the Jun N-terminal or stressactivated protein kinase (JNK–SAPK) pathway, the ERK5–BMK1
(big MAP kinase 1) pathway, and the p38 MAPK pathway. The
p38 MAPK cascade is involved in several crucial cellular signaling
events, including inflammatory and stress responses, cell growth
and differentiation, and regulation of cell cycle and cell death
(Aouadi et al., 2006; Thornton and Rincon, 2009). Four p38
MAPKs, namely p38 (MAPK14), p38 (MAPK11), p38
(MAPK12/ERK6/SAPK3) and p38 (MAPK13/SAPK4) have been
characterized in mammals (Martín-Blanco, 2000).
The role of the p38 MAPK pathway in the regulation of mitotic
entry has been well established in nonstress conditions (Cha et al.,
2007) and in response to various genotoxic stresses such as DNA
damage (Bulavin et al., 2002; Mikhailov et al., 2005). However,
evidence for direct roles of the p38 MAPKs in mitotic progression
after the G2–M transition has remained controversial and is very
limited. Takenaka and co-workers (Takenaka et al., 1998) presented
evidence that the p38 MAPK cascade is activated in NIH3T3 cells
in response to spindle damage induced by the microtubule drug,
nocodazole, but not during unperturbed mitosis. Moreover, the
authors reported that the nocodazole-induced mitotic block was
alleviated by SB203580, a putative inhibitor of p38 MAPK. In
contrast to these results, p38 MAPK pathway activation was found
to be essential for unperturbed mitosis in HeLa and NIH3T3 cells
(Tang et al., 2008) and in rat retinal cells (Campos et al., 2002).
Moreover, treatment with SB203580 led to mitotic arrest in both
studies. Last, p38 but not p38 or p38 promoted the G2–M
transition in Xenopus oocytes by activating XCdc25C (Perdiguero
et al., 2003). To get new insights into p38 MAPK signaling during
mitosis we have individually silenced each of the four p38 MAPK
isoforms in HeLa cells and monitored the cell progression through
mitosis. We find that depletion of p38 results in severe mitotic
defects including multipolar spindle formation, chromosome
misalignment and massive cell death from mitosis. This paper
represents the first comprehensive analysis of p38 function during
mitosis and identifies a new link between the p38 MAPK pathway
and the mitotic signaling network.
Results
p38-depleted cells exhibit a transient M phase arrest
followed by massive cell death
To address how the lack of p38 MAPK affects mitotic progression
in cells, we depleted individual p38 MAPK isoforms from human
HeLa cells using RNAi and followed their progression through the
cell cycle. First, the relative amounts of mRNA encoding the
individual p38 MAPK isoforms were determined using RT-PCR;
mRNAs encoding p38 and p38 were much more abundant in
HeLa cells compared with those encoding p38 and p38 (Fig.
1A,B). Transfection of 100 nM siRNA targeting p38 or p38
reduced the mRNA levels by 89.4±12.6% and 82.5±10.4%,
respectively, compared with mock-transfected cells (Fig. 1B). The
silencing of p38 (66.2±23.0% reduction) and p38 (27±14.0%
reduction) was not as successful. The results were confirmed with
quantitative PCR, which indicated reduction of mRNA encoding
p38 and p38 by 87.4±6.3% and 89.5±4.1%, whereas p38 and
p38 mRNAs were down by 59.3±9.1% and 76.9±5.2%,
respectively (Fig. 1C). We also tested two other siRNAs targeting
p38 and p38. The second p38-targeting siRNA (-siRNA2)
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p38 MAPK in mitosis
Fig. 1. p38 MAPK silencing efficacy and siRNA specificity in HeLa cells.
Quantification of p38 MAPK mRNAs from control (mock transfection) and
isoform-specific siRNA transfected HeLa cell populations 48 hours after
transfection determined using RT-PCR (A,B) and Q-PCR (C). (D)p38 MAPK
isoform mRNA levels 48 hours after silencing of p38 or p38. All data are
from three separate assays.
did not markedly reduce the p38 mRNA levels, whereas the
second p38-targeting siRNA (-siRNA2) resulted in a 58.8±4.1%
reduction of p38 mRNA (Fig. 1C). Therefore, the silencing of p38
(-siRNA1) and p38 (-siRNA1) was effective and these isotypes
were selected for further analyses. The specificity of the siRNAs
selected for further analysis were confirmed by Q-PCR analysis,
demonstrating that siRNAs targeting p38 or p38 did not affect
the quantity of mRNA encoding the other p38 MAPKs (Fig. 1D).
Next, we monitored the cell cycle progression in HeLa cells
transfected with the potent p38 siRNA (-siRNA1) or with the
two individual p38 siRNAs (-siRNA1 and -siRNA2) and
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compared results with those from cells transfected with
nontargeting control siRNA (Fig. 2A,B). Both p38-targeting
siRNAs produced the same phenotype described below. However,
the more potent one (-siRNA1) was selected for further analyses.
Live cell imaging (Fig. 2A–C) and FACS (Fig. 2E) analyses
showed normal proliferation rates (mitotic indices 4.1±1.8% and
6.2±2.3%), cell cycle progression and low cell death indices
(4.8% and 4.0% at 72 hours) after transfections with control or
p38-targeted siRNA, respectively. By sharp contrast, p38
silencing resulted in a significant accumulation of cells in mitosis
(27.4±8.3%, P<0.001) and an increase in the cell death rate
(31.6% at 72 hours, Fig. 2E). Transfection of HeLa cells with the
p38-targeting -siRNA1 resulted in a notable reduction (39%) in
the p38 protein levels 30 hours after transfection (Fig. 2D),
which most likely is an underestimation due to the induced cell
death. Since FACS might underestimate cell death because of
loss of fragmented nuclei, we confirmed p38 siRNA-induced
cell death using transmitted light microscopy. Various cell
populations were filmed using time-lapse imaging for 72 hours
starting immediately after transfection. The cell death index was
determined by classification of cells into living and dead cell
categories using the criteria defined by Häcker (Häcker, 2000) by
scoring a total of 500 cells per sample and time point indicated.
Significantly more of the p38-depleted cells (69.0±13.0%) died
by 72 hours after transfection compared with p38-depleted cells
(19±3%) and cells transfected with control siRNA (16.4±1.0%)
(Fig. 2F, P<0.001). Apo-ONE® assay indicated rapid and
statistically significant elevation of caspase-3 and caspase-7
activity in the p38-depleted cells and in the All Death® control
cells at 48 hours and 72 hours compared with control and p38depleted cells (Fig. 2G, P<0.01 at 48 hours and P<0.001 at 72
hours after transfection for p38-depleted cells). In addition, 20
M pan-caspase inhibitor, zVAD.fmk, was used to inhibit caspase
activity in the p38-depleted cell population to examine the mode
of cell death. Inhibitor was added 40 hours after siRNA
transfection and cells were followed by time-lapse imaging. The
cell death index was determined as described above. Significantly
fewer p38-depleted cells died in zVAD.fmk treated population
compared with controls; 26±7% vs 44±9% at 24 hours and 67±5%
vs 79±4% at 48 hours after drug addition, respectively (P<0.05,
supplementary material Fig. S1). These data suggest that depletion
of p38 induces caspase-dependent apoptotic cell death. During
the course of the time-lapse filming, it became evident that the
majority of p38-depleted cells die at M phase. Therefore, we
examined the fate of mitotic cells using separate sets of siRNAtransfected cells that were subjected to time-lapse filming starting
48 hours after transfection. The mitotic cells were classified into
living and dead cell categories using the same morphological
criteria as above. The viability of mitotic cells in the p38depleted population declined dramatically during the siRNAinduced M phase arrest. An average of 50% of the cells (n100)
were dead 7 hours after they had entered M phase and the
maximum death index of mitotic cells (96%) was achieved 12
hours after entry into M phase (Fig. 2H). This is significantly
different (P<0.001) from p38 and control siRNA populations,
in which fewer than 5% of the mitotic cells died during the
filming session (Fig. 2H). The majority of mitotic cells (72%,
n100) analyzed in the p38-depleted population died without
clear signs of exit from mitosis whereas 24% of the cells exited
M-phase with cytokinesis, but all daughter cells died within 4
hours of the mitotic exit.
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Fig. 2. See next page for legend.
p38 MAPK in mitosis
Journal of Cell Science
p38-depleted cells fail to maintain metaphase
chromosome alignment and show anaphase defects
To determine the effects of p38 silencing on chromosome
segregation in more detail, we used HeLa cells stably expressing
histone H2B–GFP. Populations of HeLa H2B–GFP cells were
transfected with p38, p38 or nontargeting control siRNA, and
their mitotic progression was monitored with live cell imaging
(Fig. 3, supplementary material Movies 1 and 2). As before, loss
of p38 resulted in a transient mitotic arrest that was followed by
cell death or occasionally abnormal exit from M phase (Fig. 3A,B).
The average duration of mitosis in the p38-depleted population
was significantly longer (275±130 minutes, range 49–910 minutes,
P<0.001) compared with p38-depleted cells (54±28 minutes) and
control cell population (59±19 minutes, Fig. 3C, n30–50 cells per
group in three separate assays). The p38-depleted cells typically
exhibited many misaligned chromosomes. In the majority of the
cells, chromosomes failed to move to the spindle equator, whereas
in some cells, the chromosomes first congressed to the metaphase
plate but failed to maintain their alignment (Fig. 3B). Quantification
of mitotic phases confirmed that loss of p38 led to a significant
accumulation of prometaphase cells and a decrease in the proportion
of anaphase cells (P<0.001) compared with p38-depleted and
control cells (Fig. 3B). Many of the p38-depleted cells that evaded
cell death and succeeded to enter anaphase exhibited unaligned
chromosomes or chromatid bridges (Fig. 3C). To investigate
whether silencing of p38 leads to an override of the mitotic arrest
Fig. 2. Transient mitotic arrest and loss of cell viability after p38 but not
p38 siRNA transfection. (A)Still images from time-lapse films of cell
populations treated with control, p38 or p38 siRNA. Depletion of p38
greatly suppresses cell population growth compared with control and p38
siRNA treatments. The first time point (0 h) is immediately after the
transfection. Scale bar: 50m. (B)Cell population growth curves after
transfections with control, p38, p38 or All Death siRNA. The two siRNA
duplexes targeting p38 inhibit cell growth starting ~24 hours after
transfection, whereas control and p38 siRNAs have no effect. The All Death
siRNA induces cell killing starting ~20 hours after transfection. The data are
from three separate experiments. (C)High-resolution images from time-lapse
films showing induction of a transient M-phase arrest and cell death from
mitosis after depletion of p38 whereas the cell populations transfected with
control and p38 siRNA divided normally. The first time point is ~36 hours
after transfection. The arrows indicate G2–M phase cells that either divide
normally (asterisks indicate daughter cells) or die (arrowheads). Scale bar:
10m. (D)Western blotting with an anti-p38 antibody from HeLa cell
populations depleted with control, p38 or p38 siRNAs. Cells were harvested
and prepared for western blotting 30 hours after transfection. The graph shows
the normalized p38 protein levels in the different cell populations that were
notably reduced (by 39%) in p38. (E) FACS analysis of HeLa cells
transfected with control, p38 or p38 siRNA. Loss of p38 leads to the
accumulation of cells at G2–M phase and elevated cell death (sub-G1
population). Quantification of cell cycle effects as percentages of gated sub-G1
(apo), G1 and G2–M phase cell populations. (F)The percentage of cell death
determined from control, p38 and p38 siRNA treated samples using light
microscope (data from three separate assays). The increased cell death in the
control and p38 siRNA samples at the 72 hour time point was due to
overgrowth of the populations. (G)Caspase3/7 activities as relative
fluorescence units (RFU) in control, p38 or p38 siRNA treated cell
populations determined with Apo-ONE assay kit 24, 48 and 72 hours after
transfection. Caspase-mediated apoptosis is significantly increased in p38 and
All Death siRNA cell populations compared with control and p38 siRNA
cells. (H)The viability of mitotic cells in control, p38 or p38 siRNA cell
populations determined from time-lapse films starting 48 hours after
transfection. **P<0.01, ***P<0.001. Data are means ± s.e.m.
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induced by microtubule drugs, we filmed p38-depleted and control
HeLa-H2B cells that entered mitosis in the presence of Taxol (50
cells per group were analyzed). No notable differences were
observed in the length of M phase arrest or in the number of cells
escaping the taxol block. Only 2.5% and 4.5% of the taxol-treated
p38-depleted cells and control cells, respectively, exited mitosis
within 10 hours of entering mitosis.
Lack of p38 leads to the formation of extra-microtubuleemanating foci and reduced interkinetochore tension
To examine spindle morphology in p38-depleted and control cells,
we processed the cells for immunolabeling with anti-pericentrin
and anti-tubulin antibodies 48 hours after transfection. Multipolarity
was significantly increased in p38-depleted cells (65.5±4%,
n100, P<0.001) compared with controls (3±3%), whereas no
monopolarity was observed in either population (Fig. 4A).
Typically, loss of p38 resulted in the formation of one or more
extra-microtubule-emanating foci near the main spindle poles and
elongation of the spindle apparatus without disturbance to the
general spindle architecture (Fig. 4A). The average pole-to-pole
distance at prometaphase was significantly increased in p38depleted cells to 16.3±1.82 m from the 12.7±0.68 m of control
cells (P<0.01, n25 spindles per group). Loss of p38 also reduced
the interkinetochore distance of metaphase chromosomes (Fig. 4B)
to 0.92±0.09 m (range 0.77–1.20 m, n5 sister kinetochore
pairs in each of five cells analyzed), which was almost identical to
the distance in nocodazole-treated cells with no tension (0.83±0.08
m, range 0.65–0.96 m, n5 sister kinetochore pairs in each of
five cells analyzed). These values were both significantly different
(P<0.001) from the control (1.59±0.14 m, range 1.42–2.0 m,
n5 sister kinetochore pairs in each of five cells analyzed) (Fig.
4B). We also tested whether the loss of p38 affected the stability
of kinetochore–microtubule attachments by analyzing the resistance
of kinetochore–microtubule fibers (k-fibers) to treatment with cold
Ca2+ buffer, which removes unstable microtubules (Yao et al.,
2000). This treatment abolished astral microtubules in both control
and p38 -depleted cells, but did not have any major effect on the
k-fibers (Fig. 4C).
Eg5 is required for multipolar spindle formation in p38depleted cells
To test whether p38-depleted cells exhibit mono-, bi- or multipolar
spindles upon reactivation of Eg5, a mitotic kinesin that is required
for bipolar spindle assembly (Kapoor et al., 2000), we treated
p38-depleted and control cells with 100 M monastrol (an Eg5
inhibitor) for 4 hours and then removed the drug by extensive
washing. The proteasome inhibitor MG132 was added to the culture
medium after monastrol washout to maintain mitotic arrest. In the
p38-depleted population 37±8% of the cells (n100) had arrested
in mitosis with monopolar spindles during the 4 hour drug
incubation, whereas 95±1% of the control cells (n100) were
monopolar (Fig. 4D). This difference was due to the large
population of p38-negative cells that were already arrested at M
phase at the time monastrol was added, and these cells remained
bi- or multipolar during the 4 hour incubation (Fig. 4D). The
majority of control cells (83±3%, n100) subjected to monastrol
washout into MG132-containing culture medium formed bipolar
spindles with full chromosome alignment within an hour. In contrast
to this, only 24±4% of the p38-depleted cells formed bipolar
spindles upon release from monastrol, whereas the majority of
cells formed multipolar spindles (76±3%, n100. This indicated
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Journal of Cell Science 124 (2)
Fig. 3. Impaired chromosome alignment and anaphase defects in cells depleted of p38. (A)Still images from time-lapse films of p38-depleted HeLa H2B–
GFP cells. Typically, p38-depleted cells exhibited either multiple misaligned chromosomes (upper cell, arrowheads) that never achieved metaphase alignment or
first showed normal congression of chromosomes to the spindle equator but later failure to maintain the metaphase chromosome alignment (lower cell, arrows).
Numbers indicate hours and minutes. (B)Quantification of the duration and phases of mitosis in control, p38- and p38-depleted HeLa H2B–GFP populations.
(C)A few p38-depleted cells enter anaphase in the presence of unaligned chromosomes (arrows) or exhibit the cut phenotype (arrowhead) more frequently
compared with the control and p38-depleted cells. *P<0.05, **P<0.01, ***P<0.001. Data are means ± s.e.m. Scale bars: 10m.
that p38-negative monopolar cells became multipolar upon Eg5
reactivation (Fig. 4D). We also observed that most of the p38depleted cells with bipolar spindles exhibited several unaligned
chromosomes after monastrol washout (Fig. 4E,F, yellow arrows
in bottom panel).
Active p38 MAPK localizes to kinetochores and spindle
poles and is created by bound mitotic kinases
To investigate localization of active MAPKs in mitotic cells, we
stained cells with an antibody that recognizes the dually
phosphorylated p38 MAPKs (Thr180,Tyr182; phosphorylated p38)
(Fig. 5A-E). Upon staining cells in different mitotic phases, we
detected activated p38 MAPK at the kinetochores from prophase
to late telophase, at the spindle poles from prometaphase to late
telophase and at the mid-body in telophase. Also, a notable
cytoplasmic fraction of phosphorylated p38 was present throughout
mitosis. To confirm that the antibody against phosphorylated p38
recognized the p38 isoform, we immunoprecipitated p38 from
taxol-arrested HeLa cells and performed western blotting. A strong
protein band corresponding to the expected size of the
phosphorylated p38 MAPK was present below the heavy chain in
the p38 pull-down fraction, whereas no detectable amount of
phosphorylated p38 remained in the supernatant (Fig. 5F). The
immunoprecipitation assay was also performed reciprocally with
similar results (data not shown). To exclude the possibility that the
anti-p38 antibody either recognizes or pulls down the p38
isoform we blotted the p38 immunoprecipitate with an antibody
specific to p38 (ms p38). There were no detectable levels of
p38 present in the p38 immunoprecipitate, but instead all of the
p38 isoform remained in the supernatant fraction. We conclude
that the antibody against phosphorylated p38 also recognizes the
p38 isoform in mitotic cells and that the p38 does not interact
with the p38 isoform.
To investigate which kinases are involved in p38 MAPK
activation in early mitosis, we used the lysed cell assay (Ahonen
et al., 2005; Daum and Gorbsky, 2006). In this assay, cells
growing on coverglasses are extracted with a detergent and
incubated in a buffer without the phosphatase inhibitor
microcystin-LR (MC) or PhosSTOP phosphatase inhibitor cocktail
allowing removal of the soluble cytoplasm and dephosphorylation
of phosphoepitopes at kinetochores and other subcellular
structures by the active phosphatases. Supplementation of the
lysed cells with a buffer containing MC or PhosSTOP and
ATP allows the re-creation of those phosphoepitopes that are
targeted by adjacent stably bound kinase(s) (endogenous
rephosphorylation). As expected, phosphorylated p38 signal was
abolished to background levels after dephosphorylation (Fig. 6A).
However, bound endogenous kinase(s) were able to re-create
the epitope on kinetochores and spindle poles during
rephosphorylation without any external source of kinase activity
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p38 MAPK in mitosis
Fig. 4. Spindle abnormalities in cells depleted of p38. (A)Loss of p38 results in mitosis with multipolar spindles. Immunolocalization of pericentrin foci
(arrows) and -tubulin in HeLa cells 36 hours after transfection with non-targeting control siRNA or siRNA to knockdown p38. The numbers indicate the average
pole-to-pole distance (n25 spindles per group). In the merge image, DAPI-stained chromosomes (blue), microtubules (red) and spindle poles (green/yellow) are
shown. (B)Loss of p38 reduces interkinetochore distance. In the graph, the black, white and grey lines indicate the mean, s.e.m. and range of the interkinetochore
distance, respectively. The micrographs show cells transfected with control (±3 hours nocodazole treatment) or p38 siRNA after staining of the kinetochores
(Crest) and DNA. The insets show pairs of sister kinetochores near the spindle equator. (C)Representative images of cells from control and p38 siRNA transfected
populations fixed either directly (control) or subjected to cold calcium buffer lysis before immunostaining for -tubulin. The non-stable astral microtubules (insets)
are preserved in the control conditions (arrows) and lost after cold lysis (arrowheads). In the merge, DAPI-stained chromosomes (red) and microtubules (green) are
shown. (D)In p38-depleted cells, Eg5 is required for multipolar spindle formation. The flowchart of the monastrol assay and percentages of mono-, bi- or
multipolar spindles in control and p38 siRNA transfected cell populations before and during monastrol treatment, and after drug washout is shown. (E)Control
and p38 siRNA transfected cells fixed and labelled with anti-tubulin (red) and anti-pericentrin (green) antibodies 4 hours after the addition of monastrol or 1 hour
after release into MG132-containing medium. The yellow and white arrows indicate pericentrin foci in bipolar and multipolar cells, respectively. The arrowheads
indicate unaligned chromosomes in DAPI (blue)-stained cells. (F)The lower graph shows percentages of cells with bipolar spindles with either normal or perturbed
chromosome alignment. ***P<0.001; means ± s.e.m. Scale bars: 10m.
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Fig. 5. Localization of active p38 MAPK in mitosis. (A)Antibody recognizing the dually phosphorylated p38 MAPK (Thr180/Tyr182) stains kinetochores from
prophase to late telophase, the spindle poles from prometaphase to late anaphase and the mid-body at telophase (red in the merge) in HeLa cells. Removal of
microtubules with short (1 hour) nocodazole treatment does not markedly change the phosphorylated p38 (p-p38) signal intensities whereas treatment with the
blocking peptide totally abolishes the signals. The arrows indicate the spindle poles, the arrowhead the mid-body and the insets show higher magnification views of
the kinetochores at prophase and telophase. DNA is stained with DAPI (green in the merge). (B)Line graphs showing the colocalization of active p38 MAPK (red)
with Hec1 (green, left-hand panel) at outer kinetochores but not with Aurora B (green, right-hand panel) at inner centromeres. (C)Western blotting with the antip38-P antibody detects the active p38 MAPK (arrowhead) in mitotic HeLa cell extracts. (D)The anti-p38-P antibody also recognizes the active p38 MAPK at
kinetochore and spindle poles of U2OS cells and Ptk1 cells. The micrographs show representative prometaphase cells. The merge shows overlay of DNA (grey),
Crest (green) and phosphorylated p38 (red). (E)The phosphorylated p38 epitope is conserved between the four p38 MAPK isoforms. (F)Western blots from
immunoprecipitation assay using a rabbit anti-p38 antibody. In the upper panel, the immunoprecipitated sample is blotted with anti-p38-P antibody, which detects
a strong protein band corresponding to the anticipated size of the phosphorylated p38 MAPK. The heavy chain (Hc) is detected because of use of rabbit antibody
both in the immunoprecipitation and western blotting. In the lower panel, the same sample is blotted with an antibody detecting the p38 isoform. No apparent
band is present in the immunoprecipitate whereas a strong band is detected in the supernatant (Sup) indicating that p38 does not coprecipitate with p38.
than the ATP in rephosphorylation buffer (Fig. 6B). Therefore,
kinases that are responsible for creating phosphorylated p38 were
stably bound to these subcellular structures. To investigate which
mitotic kinases control phosphorylation of p38, we tested a
number of mitotic kinase inhibitors in the rephosphorylation
assay. Interestingly, the inhibition of Polo-like kinase 1 (Plk1) by
ZK-thiazolidinone (TAL) (Santamaria et al., 2007) during the
rephosphorylation step significantly reduced phosphorylated p38
signal intensity at kinetochores and spindle poles (Fig. 6B)
whereas inhibition of Aurora B by ZM447439 (Ditchfield et al.,
2003) did not affect the phosphorylated p38 epitope. For
quantification of phosphorylated p38 at the kinetochores, we
measured the integrated fluorescence intensities minus the
background per 10 kinetochores in each of seven cells analyzed
per group as described elsewhere (Beardmore et al., 2004). When
the activities of the bound enzymes were abolished with Nethylmaleimide (NEM) and the lyzed cells were supplemented
with exogenous kinases present in M phase extract prepared from
nocodazole-blocked cells, the phosphorylated p38 epitope
reappeared at kinetochores and spindle poles (Fig. 7A,C). By
contrast, the addition of S phase extract to the lyzed cells after
NEM treatment did not lead to the re-phosphorylation of
phosphorylated p38. This indicates that kinase activity required
for creating phosphorylated p38 was present in mitotic but not in
interphase cells. When M phase extract depleted of Plk1 was
added together with ATP or when Plk1-depleted M phase extract
was washed out before addition of ATP on NEM pretreated cells,
the phosphorylated p38 signal was not regenerated (Fig. 7B,C).
This strengthens the notion that Plk1 is required for creation of
the Thr180, Tyr182 epitope of p38 MAPK.
p38 is needed to target Plk1 to the kinetochore
A recently reported mitotic substrate of the p38 MAPK pathway
is Plk1, which is phosphorylated at Ser326 by MAPK-activated
protein kinase 2 (MK2) in vitro and in vivo (Tang et al., 2008).
To analyze the effect of p38 depletion on Plk1 levels at poles and
Journal of Cell Science
p38 MAPK in mitosis
223
Fig. 6. The phosphorylated p38 epitope is created by bound mitotic kinases. (A)The phosphorylated p38 epitope is preserved at kinetochores in lyzed HeLa
cells that were fixed in the presence of the phosphatase inhibitor microcystin-LR (MC) but is lost from lyzed cells that were subjected to dephosphorylation by
endogenous phosphatases in the absence of MC. The same applies for phosphorylation of CenpA, a centromeric histone H3 homologue and target of Aurora B
kinase, detected with an antibody recognizing the phosphorylation at Ser7 (CenpA-P). (B)In the cells that were first allowed to undergo dephosphorylation by
extraction in the absence of MC, the endogenous kinetochore and spindle-pole-bound kinases can regenerate the p38-P and phosphorylated CenpA epitopes upon
incubation in buffer containing ATP and MC (upper row). When Plk1 inhibitor ZK-thiazolidinone (TAL) is added to the rephosphorylation buffer, the
phosphorylation of phosphorylated p38 epitope is prevented. Inhibition of Aurora B kinase activity by ZM477439 has no effect on p38 phosphorylation. Addition
of ZM477439 to the rephosphorylation buffer abolishes signals from the Aurora B target residue of CenpA (Ser7). Lambda phosphatase treatment abolishes p38-P
signals to background levels. The graphs show the average p38-P and CenpA-P signal intensities at the kinetochores (n10–15 in each of the 5–10 analyzed cells).
**P<0.01, ***P<0.001. Data are means ± s.e.m. Scale bars: 10m.
kinetochores, we stained p38-depleted and control cells with
antibodies against Plk1. A dramatic reduction was observed in the
kinetochore level of Plk1 (down by 93±9%, P<0.001) after p38
depletion in comparison with control cells (Fig. 8A). In these
cells, the loss of kinetochore-bound Plk1 was accompanied with
an increase in the cytosolic fraction of Plk1 (Fig. 8A). Plk1 levels
at the main spindle poles were also decreased by 41±7% in the
p38-depleted cells compared with levels in the control (P<0.01,
Fig. 8A), but unexpectedly the signal intensity of phosphorylated
Plk1 (Ser326) was not affected (Fig. 8B). This suggests that p38
MAPK isoforms other than p38 are responsible for
phosphorylation of Plk1 at Ser326 and that only a proportion of
pole-located Plk1 is phosphorylated at this residue. Further support
for this notion was provided by the observation that the
phosphorylation of MK2 at Thr334 was not reduced in the p38depleted cells (data not shown).
Discussion
The p38 MAPK pathway consists of four kinase isoforms (p38,
p38, p38, and p38) that have been implicated in inflammation,
cell differentiation, cell growth, cell death, senescence and
tumorigenesis (Aouadi et al., 2006; Thornton and Rincon, 2009).
The pathway is also an interesting therapeutic opportunity for the
treatment of various diseases and over 20 different p38 MAPK
inhibitors have entered clinical trials (Goldstein and Gabriel, 2005).
p38 was the first p38 MAPK isolated (Han et al., 1994) and is
now the most thoroughly studied isoform, whereas much less is
known about the other three family members. In addition to the
functions mentioned above, the p38 MAPK pathway also controls
the cell cycle by regulating both G1–S and G2–M transitions
(MacCorkle and Tan, 2005). A role for the pathway in mitotic entry
has been well established (Cha et al., 2007; Bulavin et al., 2002;
Mikhailov et al., 2005), but data regarding the control of mitotic
224
Journal of Cell Science 124 (2)
Journal of Cell Science
Fig. 7. Plk1 is required for the
regeneration of the
phosphorylated p38 epitope.
(A)HeLa cell M phase (panel 5)
but not S phase extract (panel 6)
contains kinase activity that can
regenerate the p38-P
phosphoepitope at kinetochores
and spindle poles of
permeabilized, dephosphorylated,
and N-ethylmaleimide (NEM)treated cells (panel 4). (B)HeLa
cell M phase extract depleted of
Plk1 by RNAi cannot regenerate
the p38-P phosphoepitope (panels
5 and 7). (C)The graph on the
right corresponds to A and shows
the average p38-P signal
intensities at kinetochores (n10
in each of the seven analyzed cells
per sample). The graph on the left
corresponds to B and shows the
average p38-P intensities at the
kinetochore in the analyzed cells
(n5). **P<0.01, ***P<0.001.
Data are means ± s.e.m. Scale
bars: 10m.
events by p38 MAPKs is very limited and controversial. Here, we
present evidence that p38 acts as important modulator of mitotic
spindle architecture and that its loss of function has an impact on
mitotic cell survival.
Role of p38 in mitotic progression and mitotic cell
survival
Our studies revealed that p38 and p38 were the most abundant
p38 MAPK isoforms expressed in HeLa cells. The presence of
p38 in cycling HeLa cells was an unexpected finding because
earlier work demonstrated that this isoform is predominantly
expressed in skeletal muscle (Lechner et al., 1996; Li et al., 1996),
whereas p38 is more ubiquitously expressed (Jiang et al., 1996).
Analyses after silencing individual p38 MAPKs revealed isoformspecific mitotic effects. In sharp contrast to p38 siRNA treatment,
which did not yield any notable cell cycle defects or induction of
apoptosis, the specific depletion of p38 led to a transient
accumulation of mitotic cells starting 24 hours after transfection.
The p38-depleted mitotic cells exhibited spindle anomalies that
prevented normal chromosome alignment and exit from mitosis.
The spindle checkpoint remained activated because these cells
spend several hours in mitosis and no notable escape from taxolinduced block was observed. The viability of the mitotic cells was
dramatically reduced in the p38-depleted populations and the
cells were eliminated via caspase-dependent apoptosis.
Our live cell assays showed that the p38-depleted cells had
several unaligned chromosomes. This could be explained by the
spindle anomalies observed in the p38-depleted cells. The cells
exhibited several microtubule-nucleating foci that perturbed the
normal spindle architecture. The multipolar phenotype was
dependent on the normal function of Eg5. Furthermore, the Eg5
reactivation assays demonstrated that loss of p38 impaired normal
chromosome congression, even in cells with bipolar spindles,
suggesting a role for p38 in the correction of erroneous
microtubule–kinetochore attachments or motor functions that move
the chromosomes. However, as the k-fiber attachments in p38depleted cells were stable according to their resistance to the cold
Ca2+ buffer treatment, chromosome alignment defects were probably
not due to mistakes in microtubule–kinetochore attachments.
Interestingly, when p38 was depleted, the interkinetochore tension
was reduced to the same level as that seen in the absence of
microtubules. Together, these phenotypes resemble the situation in
cells after treatment with low doses of microtubule drugs that
diminish interkinetochore tension but not the k-fiber occupancy
(Waters et al., 1998; Skoufias et al., 2001) and therefore suggest
that loss of p38 interferes with normal microtubule dynamics.
Mitotic substrates of the p38 MAPK pathway
In the literature, only a few mitotic targets of the p38 MAPK
cascade, such as Plk1 (Tang et al., 2008) and cytoplasmic dynein
p38 MAPK in mitosis
225
compromised the spindle assembly checkpoint. Nevertheless, the
observed requirement of p38 for the normal kinetochore
localization of Plk1 suggests a potential connection between the
p38 MAPK cascade and Plk1 as determinants of cell fate at M
phase.
Journal of Cell Science
Control of p38 MAPK activity during mitosis
Fig. 8. Loss of p38 modulates subcellular localization of Plk1. (A)Plk1 is
abolished from kinetochores and reduced at spindle poles in p38-depleted
cells. (B)Phosphorylation of Plk1 at S326 by MK2 is not affected by p38
depletion. The graphs show kinetochore (n10 kinetochores per five cells per
assay) and/or spindle pole (two main poles from five cells per assay) signal
intensities of Plk1 and phosphorylated Plk1 (p-Plk1) (Ser326). **P<0.01,
***P<0.001. Data are mean ± s.e.m. from three separate assays. Scale bars:
10m.
(Whyte et al., 2008), have been described. Both Plk1 and dynein
are implicated in several mitotic processes including spindle
organisation, chromosome motion and spindle checkpoint signaling
(Karki and Holzbaur, 1999; Petronczki et al., 2008). MK2
colocalizes with activated p38 MAPK and Plk1 at spindle poles,
where it phosphorylates Plk1 at Ser326 to promote normal mitotic
progression (Tang et al., 2008). The expression of a Plk1 phosphomutant (S326A) causes spindle pole defects and the accumulation
of cells at M phase (Tang et al., 2008).
Another phenotypic link between Plk1 and p38 is the
elimination of the M-phase-arrested cells after RNAi. Depletion of
Plk1 causes mitotic arrests, albeit with monopolar spindle, and a
dramatic decrease in cell viability (Spänkuch-Schmitt et al., 2002;
Liu and Eriksson, 2003; Reagan-Shaw and Ahmad, 2005; Guan et
al., 2005; Bu et al., 2008). Moreover, induction of apoptosis by
Plk1 RNAi is associated with p53 malfunction (Guan et al., 2005),
but whether this is linked to p38 function remains to be determined.
Interestingly, mislocalization of kinetochore-associated Plk1 by
RNAi of Polo-box domain-binding protein PBIP1 has been reported
to impair kinetochore–microtubule attachment and the generation
of tension, thus causing severe chromosome misalignment (Kang
et al., 2006). This is a reminiscent of the p38-depleted phenotype,
although PBIP1 RNAi also caused other mitotic defects and
In earlier studies, the active form of p38 MAPK was found in the
cytosol of mitotic cells (Fan et al., 2005) or localized to the mitotic
spindle poles (Tang et al., 2008). Here, we extend these findings
by reporting kinetochore signals after immunofluorescent detection
with an antibody recognizing the conserved Thr180,Tyr182
phosphoepitope present in all p38 MAPK isoforms. Activated p38
MAPK localized on kinetochores and spindle poles during mitosis.
Silencing of p38 alone was not sufficient to abolish the phoshoepitope signals, suggesting that p38 is not the only active p38
isoform present at these subcellular foci (unpublished data). Using
the lyzed cell model (Ahonen et al., 2005; Daum and Gorbsky,
2006), we tested the conditions that are required for creating
phosphorylated p38 in mitotic cells. First, kinetochore and spindlepole-bound kinases were able to re-phosphorylate the epitope
pointing to the presence of immediate upstream kinases of p38
MAPK at these subcellular locations, or TAB1-dependent
autophosphorylation mechanism of p38 MAPKs (Ge et al., 2002;
Salvador et al., 2005). Second, the capability of the M phase but
not the S phase extract to re-create phosphorylated p38 epitope
demonstrated the mitotic origin of the elements required for p38
MAPK activation. Plk1 was implicated in the process by the
observation that chemical perturbation of Plk1 but not Aurora B
kinase or loss of Plk1 from the mitotic cell extracts during the
rephosphorylation reaction was sufficient to suppress
phosphorylated p38 epitope phosphorylation by the bound kinases.
Together, the data strengthen the notion that Plk1 communicates
with p38 MAPK to control its mitotic activity. Alternatively, Plk1
might stimulate the autophosphorylation of the Thr180,Tyr182
phosphoepitope of p38 MAPKs in the TAB1-dependent reaction.
In summary, the pleiotropic mitotic errors and massive cell
death caused by loss of p38 suggest that the kinase has several
mitotic substrates whose perturbed function contributes to the
observed phenotypes. Moreover, our results provide the first
evidence for functional links between the p38 MAPK cascade and
Plk1 in the control of mitotic progression and in promotion of cell
viability at M phase. Details of such mitotic signaling reactions are
subject to further experiments in various model systems.
Materials and Methods
Cell culture
HeLa and U2OS (human osteosarcoma) cells obtained from the American Type
Culture Collection (Manassas, VA) were maintained in DMEM (Invitrogen, Carlsbad,
CA) and Ptk1 (rat kangaroo kidney epithelial) cells (a gift from Gary Gorbsky,
OMRF, Oklahoma City, OK) in MEM (Invitrogen) supplemented with 10% fetal
bovine serum, 2 mM L-glutamine, 0.1 mg/ml penicillin-streptomycin, 1 mM sodium
pyruvate, 0.1 mM non-essential amino acids and 20 mM HEPES. The medium of
stable H2B–GFP HeLa cell line (a gift from Geoffrey Wahl, Scripps Research
Institute, La Jolla, CA) included 2 ng/ml blasticidin. Cells were grown in +37°C with
5% CO2.
Antibodies and reagents
In the study we used antibody against phosphorylated p38 (Cell Signaling Technology,
Danvers, MA, 1:500), CREST autoimmune serum (Antibodies Incorporated, Davis,
CA, 1:200), anti-Plk1 (Abcam, Cambridge, UK, 1:100), antibody against
phosphorylated CenpA (Upstate Biotechnology Incorporated, Lake Placid, NY,
1:200), antibody against phosphorylated Plk1 (S326, 1:200), anti--tubulin (DM1A,
Abcam, 1:200), anti-pericentrin (Abcam, 1:2000), anti-p38 (Cell Signaling
Technology, Danvers, MA, 1:100 in IF or 1:1000 in WB) and anti-p38 (1:1000 in
WB) antibodies. The secondary antibodies anti-Cy3 (1:1000), anti-FITC (1:600) and
226
Journal of Cell Science 124 (2)
anti-Cy5 (1:400) were from Jackson ImmunoResearch (Newmarket, UK) and Alexa
Fluor antibodies from Invitrogen. Other reagents used in this study were purchased
from Sigma if not stated otherwise.
siRNA transfections
Oligofectamine (Invitrogen) was used according to the manufacturer’s instructions
using the following siRNA targeting sequences: AAC TGC GGT TAC TTA AAC
ATA (p38-oligo 1), CAG AGA ACT GCG GTT ACT TAA (p38-oligo 2), CAG
GAT GGA GCT GAT CCA GTA (p38), CTG GAC GTA TTC ACT CCT GAT
(p38-oligo 1), TGG AAG CGT GTT ACT TAC AAA (p38-oligo 2), CCG GAG
TGG CAT GAA GCT GTA (p38) and AAGATCACCCTCCTTAAATAT (Plk1). All
Death (AllStars Cell Death Control siRNA) was purchased from Qiagen (Stanford,
CA).
Immunofluorescence
HeLa, U2OS and Ptk1 cells growing on coverslips were fixed and permeabilized in
2% paraformaldehyde in 60 mM PIPES, 25 mM HEPES (pH 7.0), 10 mM EGTA,
4 mM MgSO4 (PHEM) containing 0.5% Triton X-100 and 400 nM microcystin-LR
for 15 minutes. For microtubule staining, 0.2% glutaraldehyde was added to the
fixative. Cells on coverslips were rinsed in 10 mM MOPS (pH 7.4), 150 mM NaCl
and 0.05% Tween 20 (MBST) and blocked with 20% boiled normal goat serum
(BNGS) for 1 hour at room temperature (RT). Cells were stained with primary
antibodies for 1 hour at RT. Cells on coverslips were washed with MBST and treated
with secondary antibodies for 1 hour at RT. Antibodies were diluted to MBST
containing 5% BNGS. Cells on coverslips were washed with MBST and DNA was
stained with DAPI (4⬘,6-diamidino-2-phenylindole, 10 ng/ml in water). Cells on
coverslips were washed with distilled water and mounted on microscope slides with
Vectashield mounting medium (Vector Laboratories).
Journal of Cell Science
Lyzed cell assay
HeLa cells on coverslips were rinsed briefly in rinse buffer (50 mM Tris-HCl, pH
7.5, 4 mM MgSO4, 5 g/ml protease inhibitor cocktail) and extracted in extraction
buffer (50 mM Tris-HCl, pH 7.5, 4 mM MgSO4, 0.5% Triton X-100, 1 mM DTT, 5
g/ml protease inhibitor cocktail) for 5 minutes. Samples were dephosphorylated in
dephosphorylation buffer (50 mM Tris-HCl, pH 7.5, 4 mM MgSO4, 1 mM DTT, 5
g/ml protease inhibitor cocktail) for 7 minutes (in some samples -phosphatase was
added to dephosphorylation buffer), rinsed twice in rinse buffer and treated with
either rephosphorylation buffer [50 mM Tris-HCl, pH 7.5, 4 mM MgSO4, 1 mM
DTT, 400 nM Microcystin-LR or PhosSTOP (Roche), 1 mM ATP, 5 g/ml protease
inhibitor cocktail] for 20 minutes or with buffer without ATP (50 mM Tris-HCl, pH
7.5, 4 mM MgSO4, 1 mM DTT, 400 nM Microcystin-LR or PhosSTOP, 5 g/ml
protease inhibitor cocktail). In some assays 10 M ZK-Thiazolidinone (TAL) or 20
M ZM447439 was added to the rephosphorylation buffer before incubation of the
samples for 20 minutes. In exogenous rephosphorylation assay cells were treated
with N-ethylmaleimide (NEM) buffer (5 nM NEM in 50 mM Tris-HCl, pH 7.5, 4
mM MgSO4, 5 g/ml protease inhibitor cocktail) for 10 minutes and washed in rinse
buffer. Rephosphorylation buffer ± ATP was added together with mitotic or S phase
HeLa cell extract or mitotic Plk1-depleted extract (5⫻106 cells/ml in extraction
buffer; 1:20 in rephosphorylation buffer) for 40 minutes at RT. In some samples
extracts were added in rephosphorylation buffer without ATP for 40 minutes, washed
out in presence of phosphatase inhibitor followed by ATP addition for 40 minutes.
Samples were fixed and subjected for immunofluorescence as described above.
Cold calcium lysis
Cells on coverslips were rinsed in 0.1 M PIPES (pH 6.95) and placed in ice cold (0–
2°C) 0.1 M PIPES, pH 6.95, 80 M CaCl2, 1% Triton X-100 for 5 minutes on ice
bath. After the lysis, cells were rinsed in PIPES before fixation and
immunofluorescence as described above.
Monastrol washout experiment
HeLa cells transfected with scrambled siRNA or siRNA to knockdown p38 or p38
were treated with 100 M monastrol for 4 hours. The cells were washed and released
into fresh growth medium containing 20 M MG132 for 1 hour before fixation and
staining for tubulin and pericentrin. Some samples were also fixed before monastrol
treatment or directly after the drug treatment before washout.
Microscopy and image analysis
Fixed samples were imaged with a Zeiss Axiovert microscope equipped with a 63⫻
objective, Hamamatsu Orca-ER camera (Hamamatsu Photonics Norden AB, Solna,
Sweden) and Metamorph imaging software (Molecular Devices, Downingtown,
PA). Quantification of signal intensities was done using Metamorph imaging software.
All values were corrected with background deduction.
Live cell imaging
Asynchronous or double thymidine synchronized HeLa and HeLa H2B–GFP cells
growing on a well plate or -Slide eight-well plates (Ibidi, Martinsried, Germany)
were imaged 48 hours after siRNA transfection for 16 hours. Images were captured
using a Zeiss Axiovert microscope equipped with a 40⫻ or 63⫻ objective,
Hamamatsu Orca-ER camera (Hamamatsu Photonics Norden) and Metamorph
imaging software (Molecular Devices) or with the 3I-spinning disc confocal
microscope (Intelligent Imaging Innovations, Goettingen, Germany) equipped with
63⫻ objective, Hamamatsu Orca-ER camera (Hamamatsu Photonics Norden) and
Slidebook imaging software (Intelligent Imaging Innovations). We also time-lapse
imaged asynchronous HeLa cell populations on 384-well plates immediately after
the siRNA transfection using incucyte imaging system (Essen Instruments, Welwyn
Garden City, UK) for a total of 66 hours. In this study, duration of mitosis was
defined as the time from nuclear envelope breakdown to end of telophase, or to the
time of mitotic cell death. To calculate the cell death indices we used the
morphological criteria of apoptosis described by Hacker (Hacker, 2000) and scored
1000 cells per each time point from three replicate assays.
FACS
HeLa cells transfected with scrambled, p38 or p38 siRNA were harvested after 48
or 72 hours after transfection. Cells were fixed with –20°C ethanol followed by brief
incubation in –20°C. Cells were centrifuged at 1300 rpm for 4 minutes. Cell pellets
were washed and resuspended in PBS in round-bottom 96-well plates. RNase (100
g/ml) and propidium iodide (20 g/ml) diluted in PBS were added to the resuspended
cells and the cells were incubated for 30 minutes with agitation at RT in a lightprotected box. The samples were measured with LSR II system (Becton Dickinson,
Franklin Lakes, NJ) and results were analyzed with FCS Express program.
RT-PCR
Nondepleted HeLa cells or cells depleted for p38 MAPK isoforms were harvested
48 hours after the siRNA transfection. RNA was isolated with the RNeasy Midi Kit
(Qiagen) and cDNA was prepared according to the manufacturer’s instructions using
iScriptTM cDNA Synthesis Kit (Bio-Rad, Hercules, CA). Different p38 MAPK
isoforms were amplified from cDNAs using specific primers for different isoforms
(see below). PCR products were separated on 1% agarose gel including 0.5 g/ml
ethidium bromide and imaged with UV (Gene Genius Bio Imaging System by
Syngene, Cambridge, UK). Band intensities were quantified using Metamorph
imaging software (Molecular Devices).
Q-PCR
RNA was isolated from depleted HeLa cells and treated with DNaseI (Invitrogen)
followed by cDNA isolation. p38 MAPK isoforms were amplified using isoform
specific primers with q-PCR (TaqMan®, Applied Biosystems, Foster City, CA).
Statistical analysis
Statistical analysis was performed with GraphPad Prism and Excel software using
Student’s t-test or two-factor ANOVA test. The significance threshold accepted was
P<0.05.
Caspase activity assay and inhibition
The Apo-ONE homogeneous caspase-3/7 assay (Promega, Madison, WI) was used
to evaluate the activities of caspase-3 and caspase-7. HeLa cells were depleted with
negative control, All Death control, p38 and p38 siRNAs in 384-well format using
oligofectamine reagent in a reverse transfection according to the manufacturer’s
instructions. After 24, 48 or 72 hours of incubation at 37°C, part of the medium was
aspirated and Apo-ONE substrate buffer mix was added to each well. The plate was
incubated for 30–60 minutes with agitation and was protected from light. EnVision
2100 multilabel reader (Perkin Elmer) was used to measure fluorescence (excitation
wavelength 485 nm, emission wavelength 535 nm). Caspase activity was inhibited
with 20 M pan-caspase inhibitor zVAD.fmk. Inhibitor was added 40 hours after
transfection to cells treated with siRNA against p38 in 96-well plates.
Immunoprecipitation and western blot
Dynabeads (protein G) were washed twice with citrate-phosphate buffer (pH 5)
containing 0.01% Tween-20 and left in citrate-phosphate buffer (pH 5). p38 antibody
was added on beads and incubated for 40 minutes at RT in rotation. Bead–antibody
complexes were washed three times with citrate-phosphate buffer (pH 5) containing
0.01% Tween-20. Mitotic cell extract (5⫻106 cells/ml in PBS-based IP buffer; 1⫻
PBS, 1 mM EDTA, 2 mM MgCl2, 0.2 mM CaCl2, 0.5% Triton X-100, 150 mM
NaCl) arrested for 16 hours was then incubated with 0.6 M Taxol at +4°C for 3
hours. IPs were washed three times with PBS-based IP buffer and bound proteins
were eluted by adding SDS buffer and boiling for 5 minutes. Proteins from IP
supernatant were precipitated with acetone and dried pellet was dissolved in SDS
buffer and boiled for 5 minutes. SDS-PAGE was carried out using 4–12% gradient
gels (Lonza) and proteins transferred to nitrocellulose membrane. Membrane was
blocked in 10 mM Tris-HCl (pH 8.0), 150 mM NaCl, and 0.05% Tween 20 (TBST)
containing 5% nonfat dry milk for 1 hour. Membranes were incubated with primary
antibodies overnight at 4°C, washed three times with TBST and incubated with
secondary antibodies for 1 hour at RT protected from light. After washing, the
Odyssey Infrared Imaging System (LI-COR Biotechnology) was used for detection
of proteins.
We thank Bayer Schering Pharma AG, Xiaoqi Liu and Geoff Wahl
for providing reagents for the study and Gary Gorbsky for discussions
p38 MAPK in mitosis
and critical reading of the manuscript. This study was supported by
grants to M.J.K. from the Academy of Finland (120804), EUFP6
(Marie Curie EXT grant 002697), the Centre of Excellence for
Translational Genome-Scale Biology, the Finnish Cancer Organisations
and the Foundation for the Finnish Cancer Institute.
Supplementary material available online at
http://jcs.biologists.org/cgi/content/full/124/2/216/DC1
Journal of Cell Science
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