1 Department of Cell Biology, Max-Planck-Institute for Biochemistry, Am
Klopferspitz 18a, D-82152 Martinsried, Germany
2 Department of Membrane Biology, Max-Planck-Institute for Biology,
Corrensstrasse 38, D-72076 Tübingen, Germany
* Present address: Division of Biology, California Institute of Technology, 1200
E. California Blvd, Pasadena, CA 91125, USA
Author for correspondence (e-mail:
nigg{at}biochem.mpg.de
)
Accepted 26 May 2002
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Summary |
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Key words: C-Nap1, Nek2, Centrosome, Mitotic spindle, Phosphorylation
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Introduction |
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C-Nap1 is known to interact with the mammalian serine/threonine kinase Nek2
(Fry et al., 1998a), a member
of the NIMA kinase family (Schultz et al.,
1994
). Both Nek2 abundance and activity are cell cycle regulated,
with peak levels in S/G2 and low activity in nocodazole-arrested and G1 phase
cells (Fry et al., 1995
). Both
C-Nap1 and Nek2 co-localize at the centrosome
(Fry et al., 1998a
), and
C-Nap1 most likely constitutes a physiological substrate of Nek2
(Fry et al., 1998a
).
Interestingly, protein phosphatase 1 (PP1) interacts with Nek2 and can
de-phosphorylate both C-Nap1 and Nek2
(Helps et al., 2000
). Thus, it
is attractive to speculate that phosphorylation may regulate C-Nap1 function,
and that the phosphorylation state of C-Nap1 may depend on the balance of Nek2
and PP1 activity. In support of such a model, centrosome splitting can be
induced by overexpression of active Nek2
(Fry et al., 1998b
) as well as
by drug-induced inhibition of PP1 (Meraldi
and Nigg, 2001
). From a cell cycle perspective, it is intriguing
that PP1 is inhibited at the onset of mitosis, possibly as a direct
consequence of phosphorylation by Cdk1
(Puntoni and Villa-Moruzzi,
1997
). As a result, Nek2 is expected to prevail and phosphorylate
C-Nap1 specifically at the onset of mitosis.
Although attractive, the above model remains speculative. In particular,
there is no previous evidence to indicate that endogenous C-Nap1 is
phosphorylated in vivo, let alone that it is phosphorylated in a
cell-cycle-specific manner. Furthermore, there is increasing evidence to
suggest that ubiquitin-dependent proteolysis of centrosomal proteins plays an
important role in the regulation of centrosome structure and function
(Freed et al., 1999;
Gstaiger et al., 1999
;
Fabunmi et al., 2000
). In the
present study, we thus asked whether proteolysis or phosphorylation is more
likely to explain the cell-cycle-regulated dissociation of C-Nap1. To this
end, we studied both the cell cycle regulation of endogenous C-Nap1, and the
consequences of overexpressing full-length exogenous C-Nap1. Our results
provide no evidence for proteolytic degradation of C-Nap1. Instead, they
strengthen the view that C-Nap1 localization and function is regulated
primarily by cell-cycle-regulated phosphorylation.
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Materials and Methods |
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Cell culture
HeLa and U2OS osteosarcoma cells were grown at 37°C in a 7% or 5%
CO2 atmosphere in Dulbecco's modified Eagle's medium (DMEM)
supplemented with 10% heat-inactivated fetal calf serum (FCS) and
penicillin-streptomycin (100 i.u./ml and 100 µg/ml, respectively). HeLa
cells were treated with 2 mM thymidine, 1.6 µg/ml aphidicolin, 0.5 µg/ml
nocodazole, 100 µM monastrol, 10 or 40 µg/ml MG132 (C-2211), 10 or 25
µg/ml MG115 (C-6706) or 5µg/ml ALLN (A-6185). (Monastrol was a kind gift
from Dr T. Mayer, all other drugs from Sigma.) For synchronization at G1/S,
dishes with 20% confluent cells were treated for 14 hours with thymidine,
washed three times with PBS, reincubated with normal medium for 10 hours and,
finally, treated for 14 hours with aphidicolin. For synchronization in
mitosis, cells were released for 10 hours from the G1/S block, treated with
nocodazole or monastrol for 4.5 hours and collected by mechanical shake off.
Alternatively, mitotic cells were collected by mechanical shake off after 16
hours of nocodazole treatment. For synchronization in G1, cells blocked with
nocodazole in mitosis were collected, washed and released for 6 hours into
normal medium. Mechanical shake off was then applied to discard non-adherent
cells.
For transient transfection studies, U2OS cells were seeded onto HCl-treated
glass coverslips and then transfected with 5 µg of plasmid DNA, using
calcium phosphate precipitates as previously described
(Seelos, 1997). Cells were
fixed 24 or 36 hours later in cold methanol (-20°C) for 6 minutes. To
arrest cells in mitosis, a mutated chicken cyclin B2 in pCMV-neo was used
(arginine in position 32 converted to serine,
Gallant and Nigg, 1992
).
Co-transfection was achieved by adding 2.5 µg of each of the two plasmid
DNAs. When active (wt) and inactive Nek2 kinases in pRc-CMV
(Fry et al., 1998b
) were
co-transfected with mycC-Nap1, 3.5 µg and 1.5 µg of plasmid DNA were
used for Nek2 and C-Nap1, respectively.
Enhanced GFP-C-Nap1 was stably integrated into T-REx modified U2OS cells according to manufacturer's instructions (Invitrogen), and cells were cloned by limiting dilution. Expression of the GFP-C-Nap1 fusion protein was induced for 24 hours with 1 µg/ml of tetracycline (Invitrogen, Q100-19).
Immunofluorescence microscopy
Cells were washed with PBS and blocked for 20 minutes with 1% BSA in PBS at
room temperature, before being incubated with primary antibodies for 1 hour at
room temperature. Primary antibodies were affinity-purified anti-C-Nap1 (C-Ab,
0.5 µg/ml), GTU-88 anti--tubulin mAb (1:1000; Sigma),
anti-
-tubulin mAb (1:2000; Sigma), 9E10 anti-myc mAb (undiluted tissue
culture supernatant) and GT335 anti-polyglutamylated tubulin mAb (1:2000; a
kind gift of B. Edde, CNRS, Montpellier, France). Secondary antibodies were
biotinylated donkey anti-rabbit or goat anti-mouse IgG (1:200; Amersham),
followed by Texas-Red-conjugated streptavidin (1:100; Amersham),
FITC-conjugated goat anti-mouse Fab fragment (1:100; Sigma), FITC-conjugated
goat anti-rabbit (1:100; Sigma) and Alexa-Fluor-488-conjugated goat anti-mouse
IgG (1:1000; Molecular Probes). All antibodies were diluted in 3% BSA in PBS.
Immunofluorescence microscopy was performed using a Zeiss Axioplan II
microscope and Neofluar 40x or Apochromat 63x oil immersion
objectives. Photographs were taken using Quantix 1400 (Photometrics) or
Micromax (Princeton Instruments) CCD cameras and IP-Lab or Metaview (Universal
Imaging) software.
Immunochemical techniques
To prepare extracts for immunoblotting, cells were lysed in RIPA buffer (50
mM Tris-HCl, pH 8, 1% Nonidet P40, 0.5% deoxycholic acid, 0.1% SDS, 150 mM
NaCl, 1 mM PMSF, 1 µg/ml aprotinin, 1 µg/ml leupeptin, 1 µg/ml
pepstatin A, 20 mM ß-glycerophosphate, 20 mM NaF and 0.3 mM sodium
vanadate). Samples were left for 30 minutes on ice, passed 10 times through a
27G needle and centrifuged at 16,000 g for 10 minutes at
4°C. Protein concentrations were determined using an improved Lowry assay
(Dc protein assay, Biorad). C-Nap1 antibodies were rabbit polyclonal
C-Ab (aa 1982-2442) and M-Ab (aa 1098-1248)
(Fry et al., 1998a;
Mayor et al., 2000
). All
antibody incubations were carried out in blocking buffer (5% nonfat dried milk
in PBS/0.1% Tween-20) at the following dilutions: C-Ab, 0.5 µg/ml; M-Ab, 1
µg/ml; anti-cdc27 serum, 1/1000; anti-cyclin A, 1/500; anti-
-tubulin
mAb, 1/1000 (Sigma); anti-GFP mAb, undiluted tissue culture supernatant. Bound
IgGs were visualized using alkaline phosphatase-conjugated anti-rabbit or
anti-mouse IgG secondary antibodies (1:7500; Promega).
Alkaline phosphatase treatment
HeLa cells were lysed in IP buffer (0.5% Igepal, 150 mM NaCl, 50 mM
Tris-HCl pH 8, 20 mM ß-glycerophosphate, 20 mM NaF and 0.3 mM
NaVO4). Samples containing equal amounts of proteins were mixed for
1 hour at 4°C with C-Ab (8 µg/ml) pre-bound to Affiprep protein A beads
(1 µl/µg of Ab; Biorad). Beads were washed three times with IP buffer
and resuspended in 2x200 µl of AP buffer (10 mM Tris-HCl, 10 mM
MgCl2, 50 mM NaCl, 1 mM DTT, pH 7.9, with 1 mM PMSF, 1 µg/ml
aprotinin, 1 µg/ml leupeptin and 1 µg/ml pepstatin). Reactions were then
carried out by incubating these 200 µl aliquots for 40 minutes at 37°C
with 20 units of either active or heat-inactivated (10 minutes at 95°C)
calf intestinal alkaline phosphatase (M0290; New England BioLabs). Finally,
beads were resuspended in sample buffer.
In vivo [32P] labeling
HeLa cells in G1 were collected 4 hours after release from a 16 hour
nocodazole block. Cells in S, G2 and M phase were obtained after releases for
2, 6 or 10 hours, respectively, from a double-block with
thymidine-aphidicolin. M-phase cells were further blocked with nocodazole for
an additional 5 hours and then collected by mechanical shake-off. All cells
were labeled by additional incubation for 4 hours in phosphate-free DMEM
medium that had been supplemented with 10% dialyzed FCS, 10% normal DMEM, 1%
normal FCS and 400 µCi/ml of phosphoric acid (1 mCi in total). Cells were
collected and lysed in 400 µl/2x106 cells of modified-IP
buffer (0.4% Igepal, 0.1% deoxycholic acid, 150 mM NaCl, 50 mM Tris-HCl pH 8,
20 mM ß-glycerophosphate, 20 mM NaF and 0.3 mM NaVO4).
Orthophosphate incorporation in the various cell extracts was measured by
scintillation counting, and equal protein levels were confirmed by Coomassie
blue staining. Samples were then pre-cleared using rabbit IgG bound to Protein
A-Dynabeads (Dynal, 10 µg IgG to 40 µl beads/400 µl extract), and
immunoprecipitations were performed with C-Ab (and rabbit IgG for control)
bound to Protein A-Dynabeads pre-blocked with HeLa cell extracts (5 µg IgG
to 20 µl beads/200µl extract). To ensure low background, samples were
resuspended in sample buffer (30 µl), heated for 5 minutes at 95°C,
diluted in 1 ml of modified-IP buffer and subjected to a second round of
immunoprecipitation.
Miscellaneous techniques
Immunoelectron microscopy was performed as described previously
(Fry et al., 1998a). For
assaying protein degradation, cyclin B2 and myc-C-Nap1 were in vitro
translated, in the presence of [35S]-Met (TNT coupled reticulocyte
system, Promega), and added to M-phase Xenopus egg extracts (12.5% of
final volume). These were prepared by complementing interphasic extracts with
recombinant
90-cyclin B and cycloheximide (0.25 µg/µl), followed
by a 90 minute incubation at 22°C
(Vorlaufer and Peters, 1998
).
Aliquots were removed every 15 minutes, mixed with sample buffer and heated
for 5 minutes at 95°C.
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Results |
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The above results suggested that C-Nap1 could be subject to
cell-cycle-regulated proteolytic degradation. To determine whether the
inhibition of the proteasome would result in a stabilization of C-Nap1,
M-phase arrest experiments were performed in the presence of increasing
amounts of the proteasome inhibitor MG132. As shown in
Fig. 2A, C-Nap1 levels were
reduced even in the presence of this inhibitor. In contrast, the
proteasome-target cyclin A was stabilized, as expected, demonstrating the
efficacy of proteasome inhibition (Fig.
2A). Virtually identical results were obtained using the
proteasome inhibitors MG115 and N-acetyl-leucyl-leucyl-norleucinal (ALLN)
(data not shown). In principle, C-Nap1 could be a substrate for other
proteases, but we emphasize that no distinct cleavage products could ever be
observed in any of the above experiments. To more directly assess C-Nap1
stability during M phase, we turned to Xenopus egg extracts, which
provide a convenient assay system for examining protein degradation
(Vorlaufer and Peters, 1998).
When [35S]-labeled, in vitro translated C-Nap1 and cyclin B were
added to such extracts, C-Nap1 was completely stable, whereas cyclin B was
almost totally degraded after 60 minutes of incubation
(Fig. 2B). Taken together,
these results indicate that C-Nap1 is not subject to degradation by the
proteasome, and they also argue against cleavage of C-Nap1 by other
M-phase-specific proteases.
Evidence for M-phase-specific phosphorylation of C-Nap1
As the fate of mitotic C-Nap1 did not seem to be governed by proteolytic
degradation, we next considered the possibility that the M-phase-specific
reduction in the C-Nap1 signal could reflect a post-translational
modification. In particular, we reasoned that M-phase-specific phosphorylation
might have caused an altered, more diffuse electrophoretic mobility of C-Nap1,
or a reduced transfer to nitrocellulose membranes. To explore this
possibility, C-Nap1 was immunoprecipitated from extracts at different cell
cycle stages, and then treated with calf intestinal phosphatase prior to
western blotting. As shown in Fig.
3A, the staining of mitotic C-Nap1 was increased in response to
active phosphatase but not the heat-inactivated enzyme. Similar results were
obtained when mitotic cell extracts were incubated at 37°C in the absence
of phosphatase inhibitors (data not shown). These data show that a
near-interphasic C-Nap1 signal could be recovered by treatment of mitotic
C-Nap1 with an active phosphatase, implying that C-Nap1 was phosphorylated
when isolated from mitotic cells.
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To directly demonstrate that C-Nap1 is phosphorylated during mitosis, in vivo labeling with [32P]phosphoric acid was performed in HeLa cells. Extracts were then prepared from different cell cycle stages, and C-Nap1 was immunoprecipitated and analyzed by SDS-PAGE. As shown by autoradiography and western blotting, immunoprecipitated C-Nap1 was labeled with [32P] specifically during M phase, but not during G1, S or G2 phases (Fig. 3B). No [32P]-labeling of C-Nap1 was observed in control immunoprecipitation experiments using nonspecific rabbit IgG (data not shown). Taken together, these data indicate that C-Nap1 is indeed phosphorylated specifically during mitosis.
Overexpression of C-Nap1 causes the formation of large
centrosome-associated structures
The above results strongly suggested that phosphorylation, rather than
proteolysis, regulates the displacement of C-Nap1 from the centrosome at the
onset of mitosis. However, because only a fraction of the total C-Nap1 protein
(estimated to about 30-50%) is associated with the centrosome, it is difficult
to obtain definitive information about local events occurring at the
centrosome. As a complementary approach to studying the fate and regulation of
C-Nap1 during the cell cycle, we thus decided to examine the phenotypic
consequences of overexpressing C-Nap1 in human cells. In particular, we asked
whether a high level of C-Nap1 could prevent the dissociation of C-Nap1 from
the centrosome, and thus possibly interfere with centrosome separation at the
onset of mitosis. Full-length C-Nap1, with or without an N-terminal myc-tag,
was therefore placed under the control of a CMV promoter and transiently
transfected into U2OS cells. Then, the localization of the ectopically
expressed protein was determined by immunofluorescence microscopy. In
parallel, the distribution and morphology of centrosomes was examined by
staining with antibodies against -tubulin. In cells expressing low
levels of exogenous C-Nap1, the full-length protein localized correctly and
almost exclusively to the centrosome (data not shown). With higher expression
levels, however, cytoplasmic staining became increasingly prominent, and, most
strikingly, one or several compact C-Nap1-positive aggregates (termed patches)
became visible (Fig. 4A). In
some cells, these patches grew into large structures occupying almost the
entire cytoplasm. Immunoelectron microscopy performed on cells overexpressing
C-Nap1 revealed that the centrioles were invariably embedded in large
structures showing extensive labeling with anti-C-Nap1 antibodies
(Fig. 4B). These structures
often emanated primarily from the proximal ends of centrioles
(Fig. 4Ba), but eventually
extended to engulf almost the entire centrioles
(Fig. 4Bb).
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To further characterize the relationship between the observed C-Nap1
patches and centrosomes, antibodies against different centrosomal proteins
were used for double immunofluorescence microscopy. Endogenous Nek2 protein
was found to accumulate rather uniformly in all C-Nap1 patches
(Fig. 5Ab), consistent with a
direct interaction between these two proteins
(Fry et al., 1998a). In
contrast, the distribution of
-tubulin appeared largely unaltered, in
that anti-
-tubulin antibodies stained the typical one or two dots per
cell (Fig. 5Ac). In cases where
only a single C-Nap1 patch was present within a cell, this patch was
invariably associated with the
-tubulin-positive centrosome. However,
if multiple patches were present, only one stained positive for
-tubulin (Fig. 5A).
Similar results were obtained for other centrosome-associated proteins,
notably centrin-2, p150Glued and Plk1, with none of them showing a
particular enrichment in the C-Nap1 patches (data not shown). These results
indicate that the observed patches are composed primarily of C-Nap1 itself,
and that they incorporate at most a limited subset of endogenous centrosomal
proteins. Consistent with this interpretation, none of the extra-centrosomal
C-Nap1 patches displayed any microtubule (MT) nucleation activity. On the
contrary, a MT re-growth assay showed that centrosomes embedded in C-Nap1
patches displayed severely reduced MT nucleation activity
(Fig. 5B). It seems plausible
that this reflects steric hindrance exerted by the C-Nap1 oligomer.
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In order to identify sub-domains within C-Nap1 responsible for patch formation, several truncation mutants of C-Nap1 were expressed in U2OS cells (Fig. 5C). We found that a mutant from which only the C-terminal domain had been deleted (T4) still formed extensive patches, but that further deletion of the C-terminal coiled-coil domain (T5) abolished patch formation. Furthermore, none of the mutants (T6, T7) lacking the N-terminal end domain formed patches. These data suggest that self-assembly of C-Nap1 into oligomeric structures requires the N-terminal end domain and both central coiled-coil domains.
Overexpression of C-Nap1 does not prevent centrosome separation
Next we sought to determine whether the overexpression of C-Nap1 would
deregulate centrosome separation and/or cell cycle progression. Initial
experiments based on flow cytometry failed to reveal any significant
differences between the cell cycle profiles of populations transfected with
either C-Nap1 or pBK-CMV empty vector (data not shown). To ask more directly
whether C-Nap1 overexpression would prevent entry of cells into mitosis,
C-Nap1 was co-expressed with a non-destructible mutant of cyclin B2, which is
known to cause the accumulation of mitotic cells
(Gallant and Nigg, 1992). If
the observed embedding of centrosomes into large C-Nap1 patches was to block
entry of cells into mitosis, co-expression of C-Nap1 should have suppressed
the accumulation of mitotic cells. However, this was not observed: upon
co-expression of C-Nap1 with non-destructible cyclin B2, about 50% of the
transfected cells arrested in mitosis (Fig.
6A). This extent of mitotic arrest was similar to that seen in
control experiments, in which non-destructible cyclin B2 was co-transfected
with the enhanced GFP marker alone (Fig.
6A). Conversely, the accumulation of mitotic cells was severely
suppressed upon co-expression of the non-destructible cyclin B2 with a
dominant-negative Cdk1 mutant. As this Cdk1 mutant is known to cause a G2
arrest (van den Heuvel and Harlow,
1993
), this result validates our experimental approach and
establishes that overexpression of C-Nap1 does not prevent cells from entering
mitosis.
|
From the above results it is difficult to escape the conclusion that cells
were able to undergo mitosis in spite of the fact that their interphase
centrosomes were embedded in large C-Nap1 structures. This begs the question
of how such embedded centrosomes could separate and contribute to the
formation of a functional mitotic spindle. To address this issue, we
established a U2OS cell line stably expressing a full-length GFP-C-Nap1 fusion
protein under tetracycline control. Within 24 hours of tetracycline addition,
exogenous GFP-C-Nap1 was readily expressed, as demonstrated by western
blotting (Fig. 6B). When
analyzed by fluorescence microscopy, GFP-C-Nap1 formed patches that were
indistinguishable from those described above (data not shown, but see below).
The stably transformed U2OS cells were then grown for 6 days in the presence
or absence of tetracycline, and their growth curves determined. In a third
U2OS cell population, GFP-C-Nap1 expression was pre-induced for 24 hours,
before the culture was continued for 6 days in the presence of tetracycline.
As shown in Fig. 6C, all three
cell lines showed indistinguishable proliferation rates, confirming that cells
proliferate normally in the presence of extensive C-Nap1 patches. To further
test this surprising conclusion, we directly examined spindle formation in
cells harboring C-Nap1 patches. Spindle MTs were stained with antibodies
against -tubulin (Fig.
7A), whereas centrioles were stained with antibodies against
poly-glutamylated tubulin [GT335 (Wolff et
al., 1992
; Bobinnec et al.,
1998
)] (Fig.
7Ba,b), and centrosomes with antibodies against
-tubulin
(Fig. 7Bc). At the level of
resolution provided by immunofluorescence microscopy, no obvious defects in
centrosome migration, spindle formation or chromosome segregation could be
detected (Fig. 7b,c). However,
a most remarkable change occurred in the relationship between centrosomes and
C-Nap1 patches. Whereas all interphase centrosomes were embedded in C-Nap1
patches (Fig. 7Aa, Ba), none of
the mitotic spindle poles seen in metaphase, anaphase or telophase cells ever
showed any association with such patches
(Fig. 7Ab,c, Bb,c, and data not
shown). In contrast, C-Nap1 patches that were not in direct contact with
centrosomes persisted in the cell body throughout mitosis
(Fig. 7). Most interestingly,
we also observed several examples of prophase cells in which one centrosome
was still embedded within a C-Nap1 patch, whereas the other was already free
and positioned at some distance (Fig.
7C). The one C-Nap1 patch that was still in contact with a
centrosome often appeared disturbed and partially dissolved
(Fig. 7C, see enlarged
insets).
|
Virtually identical results were obtained when examining cells after transient overexpression of a non-tagged version of C-Nap1 (data not shown), indicating that the GFP marker was not responsible for the loss of C-Nap1 interaction with mitotic spindle poles. From these results we conclude that the centrosome-associated C-Nap1 patches are subject to disassembly at the very onset of mitosis, when chromosome condensation begins and centrosomes separate. Furthermore, our observations strongly argue against a generalized mechanism for the disassembly of C-Nap1-containing structures. Instead, they point to a mechanism that is activated specifically at the centrosome. As a consequence, centrosome-associated C-Nap1 structures are locally dismantled, allowing centrosome separation and spindle formation to occur, although C-Nap1 patches elsewhere in the cell persist throughout mitosis.
Active Nek2 interferes with C-Nap1 patch formation
In previous studies, we have shown that the centrosome-associated protein
kinase Nek2 can bind and phosphorylate C-Nap1, both in vitro and in vivo
(Fry et al., 1998b). Here, we
have shown that C-Nap1 is almost certainly phosphorylated during M phase, and
that overexpressed C-Nap1 is specifically removed from centrosomes at the
onset of mitosis. Therefore, it was of obvious interest to ask whether
co-expression of Nek2 would influence the formation or stability of C-Nap1
patches. To this end, C-Nap1 was co-expressed with either wild-type Nek2 or a
catalytically inactive Nek2 mutant. Then, cells were scored for the frequency
and appearance of C-Nap1 patches (Fig.
8A). To facilitate data analysis, transfected cells were grouped
into three classes, depending on the observed C-Nap1 distribution
(Fig. 8B). The first class
comprised cells showing little, if any, unusual C-Nap1 structures
(Fig. 8Ba). In the second
class, we counted cells with one or few (<10) large patches
(Fig. 8Bb), and in the third
class cells harboring many, often small C-Nap1-positive patches (>10) or
speckles (Fig. 8Bc). Using this
classification, we observed a very striking effect of active, but not inactive
Nek2, on the formation of C-Nap1-positive structures. Cells expressing the
catalytically inactive Nek2 mutant in fact contained predominantly class II
cells with few but large C-Nap1 patches
(Fig. 8A), as observed for
cells transfected with C-Nap1 only (see above). In contrast, the presence of
active Nek2 strikingly diminished the frequency of cells with large C-Nap1
structures, and instead, most cells now fell into class III, showing a
multitude of small C-Nap1-positive speckles
(Fig. 8A). We interpret this
observation to indicate that Nek2 kinase activity interfered with the
formation of large C-Nap1 aggregates, thereby favoring the formation of a
multitude of small C-Nap1 structures. Although obtained in an artificial
experimental setting, these results support the view that C-Nap1 is an in vivo
substrate of Nek2, and that Nek2 activity may thus be responsible for the
dissociation of C-Nap1 from mitotic spindle poles.
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Discussion |
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C-Nap1 can self-assemble into large structures
In a second, complementary approach, we have examined the consequences of
overexpressing C-Nap1 in human cells. The main purpose of these experiments
was to ask whether deregulated expression of C-Nap1 would interfere with
centrosome separation at the onset of mitosis, and thus possibly block mitotic
progression. We found that exogenous C-Nap1 formed large structures (termed
patches) within the cell cytoplasm. The first of these patches invariably
assembled around the centrosome, but with increasing C-Nap1 levels, additional
patches formed throughout the cytoplasm. Of several centrosome-associated
proteins examined (Nek2, -tubulin, centrin-2, Plk1,
p150Glued), only Nek2 was recruited to a significant extent to the
C-Nap1 patches. This underscores the affinity of Nek2 for C-Nap1, but also
suggests that ectopically expressed C-Nap1 interacts largely with itself,
recruiting at most a limited subset of centrosomal proteins into the patches.
As shown by a MT re-growth assay, C-Nap1 patches interfered with MT-nucleation
activity. We believe that this reflects steric hindrance rather than a direct
role of C-Nap1 in MT-nucleation. Conversely, no increased MT-nucleation
activity could be observed, in line with the view that C-Nap1 plays a
structural role at the centrosome.
Although the C-Nap1 patches described here clearly represent
non-physiological structures, they are reminiscent of aberrant
centrosome-related structures that occur in human tumors. Centrosomal
aberrations have long been described in human cancers
(Boveri, 1914), and more
recently were shown to include supernumerary centrioles, excess pericentriolar
material and fragmented centrosomal structures (for reviews, see
Lingle and Salisbury, 2000
;
Brinkley, 2001
;
Duensing and Munger, 2001
;
Doxsey, 2001
). If C-Nap1
overexpression were to occur under pathological conditions, our present
results predict that this would probably not affect cell division. However,
considering that centrosomes play an important role in organizing the
microtubule network also during interphase of the cell cycle, it would be
premature to exclude that aberrant centrosomal structures formed by C-Nap1 and
other structural proteins could interfere with MT-dependent processes such as
cell shape, polarity or motility.
C-Nap1 patches are dismantled around centrosomes during prophase
Considering the extensive embedding of centrosomes into C-Nap1 patches, and
the concomitant suppression of MT nucleation, we were surprised to find that
C-Nap1 patches did not detectably interfere with cell proliferation. However,
detailed examination of the fate of C-Nap1 patches during the cell cycle
offered an explanation for this apparent paradox: staining of centrosomes with
appropriate antibodies in fact revealed that centrosomes are extracted from
the embrace of C-Nap1 patches during early prophase. As a result, bipolar
spindles readily formed, and spindle poles were never embedded within C-Nap1
patches. In striking contrast, C-Nap1 patches that were not in direct contact
with centrosomes persisted throughout cell division. These results strongly
suggest that centrosomes harbor an activity that, at the onset of mitosis, can
trigger the local disassembly of C-Nap1. As suggested previously
(Fry et al., 1998a), the
protein kinase Nek2 is a prime candidate for providing such an activity. This
notion was further corroborated here by our demonstration that co-expression
of Nek2 with C-Nap1 profoundly affected the formation of C-Nap1 patches,
strongly indicating that Nek2 is able to phosphorylate C-Nap1.
Although attractive, the idea that Nek2 triggers C-Nap1 dissociation from
centrosomes is confounded by two problems: first, as we have shown here, Nek2
is recruited to all C-Nap1 patches, and yet only the centrosome-associated
patch disassembles at the onset of mitosis. Hence, another
centrosome-associated activity may be limiting for C-Nap1 disassembly.
Alternatively, our observations could be explained if the Nek2 antagonist PP1
was also recruited to C-Nap1 patches, but only inactivated by Cdk1/cyclin B at
the centrosome. Although we presently have no appropriate antibodies to test
whether PP1 is recruited to C-Nap1 patches, the observed
centrosome-association of Cdk1/cyclin B
(Bailly et al., 1992) would be
consistent with such a scenario. A second problem stems from the previous
finding that Nek2 protein and activity levels are strongly reduced in
nocodazole arrested cells (Fry et al.,
1995
). At first glance, this would seem to argue against Nek2
being the kinase responsible for phosphorylating C-Nap1 in the experiments
described here. However, we emphasize that recent studies have revealed the
existence of two alternatively spliced forms of Nek2, Nek2A and Nek2B, not
only in Xenopus (Fry et al.,
2000
; Uto and Sagata,
2000
) but also in human cells
(Hames et al., 2001
). Although
Nek2A is degraded in nocodazole-arrested cells, due to ubiquitin-dependent
proteolysis, Nek2B is stable (Hames et
al., 2001
). Thus, this newly discovered, shorter form of the Nek2
kinase could readily continue to phosphorylate C-Nap1 even after the
destruction of Nek2A. To definitively prove that Nek2A and/or B is the kinase
responsible for C-Nap1 disassembly, it will be necessary to identify the in
vivo phosphorylation sites within C-Nap1 and compare them with the sites
phosphorylated by Nek2 in vitro. So far, all our attempts at identifying in
vivo phosphorylation sites within C-Nap1 have been unsuccessful, at least in
part due to low protein abundance. However, with the increasing sensitivity of
mass spectrometry, it seems legitimate to hope that this goal may come within
reach in the not too distant future.
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References |
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