Wellcome Trust Centre for Cell Biology, Institute of Cell and Molecular Biology, University of Edinburgh, King's Buildings, Edinburgh, EH9 3JR, UK
* Author for correspondence (e-mail:a.merdes{at}ed.ac.uk )
Accepted 6 February 2002
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Summary |
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Key words: Spindle pole, Microtubule-associated protein, Mitosis
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Introduction |
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Other spindle pole proteins, NuMA and TPX2, have been identified only in
vertebrates (Compton et al.,
1992; Yang et al.,
1992
; Wittmann et al.,
2000
). Both NuMA and TPX2 show a cell-cycle-dependent
localization: they are nuclear during interphase, and re-localize to the
spindle poles in mitosis. Despite varying protein compositions, the mechanisms
for spindle pole organization in vertebrates and Drosophila seem to
share striking similarities: NuMA has been shown to associate with the
minus-end directed motor dynein and the activator dynactin and is transported
towards the poles at early stages of spindle formation
(Merdes et al., 1996
;
Merdes et al., 2000
). By
attaching to parallel microtubules in the spindle, the moving NuMA complex can
focus them in a zipper-like fashion. By analogy, it was proposed that the
Drosophila protein Msps is transported polewards by Ncd, where it has
a stabilizing effect on the microtubule ends
(Cullen and Ohkura, 2001
).
Whereas Msps is anchored to the spindle poles by D-TACC, the mechanisms that
retain NuMA at the poles are less clear: Although the association of NuMA with
dynein and dynactin in a multi-protein complex can explain the transport of
NuMA along spindle fibres and the process of microtubule focussing, the
question remains how NuMA is able to attach and accumulate at the poles, and
why the protein isn't falling off the microtubule minus-ends following
transport. Previously, we suggested that NuMA might possess a direct affinity
for microtubules (Merdes et al.,
1996
), which could provide stable crosslinking of spindle fibres
once NuMA is deposited at the poles. In this report, we provide direct
evidence for this model: we map and characterize a 100-residue region in the
tail domain of NuMA that binds directly to tubulin, and that induces formation
of microtubule bundles with an increased stability.
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Materials and Methods |
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Hexa-histidine tagged bacterial fusion proteins of Xenopus NuMA
tail and rod fragments were prepared as described
(Merdes et al., 1996). A
hexa-histidine tagged fusion protein of the Xenopus NuMA head domain
was expressed using pRSET-C (Invitrogen, San Diego, CA), in which
Xenopus NuMA nucleotides 196 to 885 were cloned at BamHI and
EcoRI sites, using PCR products of previously identified library
clones (Merdes et al., 1996
).
A vector encoding hexa-histidine tagged
SNAP
(Whiteheart et al., 1993
) was
obtained from C. Rabouille, University of Edinburgh. All fusion proteins were
solubilized from bacteria in 8 M urea, 50 mM sodium phosphate, pH 7.6,
purified over Ni-agarose (Qiagen, Hilden, Germany), and 50 µg of each were
precipitated with 1.7 volumes of saturated ammonium sulfate. The proteins were
solubilized in a total volume of 30 µl containing 23 µg of
phosphocellulose-purified tubulin in BRB80. Following 1 hour incubation at
room temperature, samples were diluted with 100 µl of cytoskeleton buffer
(0.1 M NaCl, 0.3 M sucrose, 10 mM PIPES, 3 mM MgCl2, pH 6.8),
incubated for a further 10 minutes, and mixed with Ni-NTA magnetic agarose
beads, pelleted from 50 µl of a 5% suspension (Qiagen). After 10 minutes of
incubation on a rotator, the beads were separated from the supernatant using a
magnet, and washed twice with 1 ml of cytoskeleton buffer containing 0.5 M
NaCl, and twice containing additional 0.2% Triton X-100. Proteins were eluted
in SDS sample buffer, separated by gel electrophoresis, and immunoblotted
using either monoclonal antibody DM1
against tubulin (Sigma, Dorset,
UK), or antibody against an epitope of 6xHis-Gly from Invitrogen (San
Diego, CA).
In a different set of experiments, NuMA fusion proteins were dialyzed
against PBS, and 1.5 µg were mixed with 15 µg of previously polymerized
and taxol-stabilized tubulin in a total volume of 20 µl BRB80. After 15
minutes of incubation at room temperature, the solution was carefully
underlayed by 20 µl of 30% glycerol in BRB80, and centrifuged for 10
minutes at 16,000 g. Supernatants and pellets were analyzed by gel
electrophoresis. For the estimation of the binding constant
(KA) of NuMA tail II to tubulin, increasing amounts of
NuMA tail II were used in this assay, and the material in supernatants and
pellets was quantified by scanning of Coomassiestained gels, and by
quantitative immunoblotting using a phosphoimager and antibodies against
tubulin or 6xHis-Gly, followed by 125I-labelled protein A.
Analogously, microtubule binding of 0.4 µg NuMA tail II that were
phosphorylated with recombinant cdc2/cyclinB (New England Biolabs,
Hertfordshire, UK) and [-32P]ATP was assayed after mixing
with increasing amounts of unlabelled NuMA tail II; bound material was
quantified directly from dried protein gels on a phosphoimager.
Morphological effects of NuMA tail fragments on microtubule formation with
pure tubulin were studied as described
(Merdes et al., 1996), with
the modification that rhodamine-labelled tubulin was added to the assay.
Microtubule aster formation in Xenopus egg extracts was studied by
mixing 2.5 µg NuMA tail II or tail IIA with 10 µl of metaphase extract
and incubating for 35 minutes (Merdes et
al., 1996
). Affinity adsorption experiments in Xenopus
egg extracts were carried out using bacterial 6xHis NuMA tail I and tail
II proteins dialyzed against PBS and diluted to 0.2 mg/ml in 50 µl samples
of extract. The NuMA tail proteins were recovered using 30 µl of Niagarose,
followed by two washes in PBS and one wash in 0.2% Triton X-100 in PBS, and
elution in 30 µl SDS gel sample buffer. Tubulin was identified by
immunoblotting.
Transfection experiments and microscopy
HeLa cells were grown in Dulbecco's modified Eagle's medium with 10% fetal
calf serum and transfected with calcium phosphate. To increase the mitotic
index, a single block of 15 hours with 2 mM thymidine was used. Cells were
fixed either in methanol for 20 minutes at -20°C, or with 3.7%
formaldehyde in 0.1 M NaCl, 0.3 M sucrose, 10 mM PIPES, 3 mM MgCl2,
pH 6.8 for 10 minutes at room temperature, followed by permeabilization in
0.2% Triton X-100. Cells were incubated with PBS, 0.1% Tween, 0.5% BSA for 5
minutes, then with primary antibody [i.e. mAb1F1 anti-NuMA (Compton, 1991);
anti tubulin DM1, or anti-acetylated tubulin (both from Sigma, Dorset,
UK) for 30 minutes], and with secondary antibody (Texas-Red-conjugated
anti-mouse, Sigma, Dorset, UK) for 30 minutes. For actin staining, cells were
incubated with TRITC-phalloidin (0.25 µM, Sigma, Dorset). After DNA
staining with DAPI (2.5 µg/ml), coverslips were mounted on microscope
slides with Vectashield (Vector Laboratories, Burlingame, CA) and sealed with
nail polish. Conventional fluorescence microscopy was performed as described
previously (Merdes et al.,
2000
). For electron microscopy, cells on coverslips were fixed
with 1% glutaraldehyde in PBS. Cells that expressed GFP-NuMA tail II were
identified by fluorescence microscopy and photographed, and coordinates of the
microscope stage were recorded for following relocation. Cells were
subsequently treated with 2% osmium tetroxide, dehydrated in a graded series
of ethanol, and flat-embedded in Araldite CY212 (Agar Scientifc, Essex, UK).
After relocation of the transfected cells, the glass coverslip surface was
removed with hydrofluoric acid, and the cells were mounted on blocks of
Araldite. Ultrathin sections were counterstained with lead citrate and viewed
on a Philips CM120 electron microscope.
Construction of GFP-NuMA derivatives
Constructs were derived from GFP-human NuMA in the eukaryotic expression
vector pCDNA3 (Merdes et al.,
2000). Fragments of NuMA tail were obtained from this template by
PCR, using the proofreading Pfu polymerase (Stratagene, Amsterdam,
Netherlands). All primers carried the restriction sites NotI
(5') and XbaI (3', following a STOP codon), allowing
substitution of the full-length NuMA in the parental plasmid by the tail
fragments. In our constructs, nucleotide positions of NuMA cDNA [GenBank
Z11584 (Compton et al., 1992
)]
were the following: 5101-5595 for tail I, 5602-6306 for tail II, 5602-5901 for
tail IIA, 5932-6306 for tail IIB. For the deletion of the nuclear localization
signal in GFP-NuMA, wild-type sequence between EcoRV and
XbaI in pCDNA3 GFP-NuMA was replaced by an
EcoRV/EcoRI fragment of pUC19 NuMA
NLS
(Saredi et al., 1996
). The
deletion constructs NuMA
tailII and
tailIIA+NLS were cloned by
substitution of the 3' end of NuMA in pCDNA3 GFP-NuMA with PCR products.
For NuMA
tailII, the PCR product extended from a single AatII
site to nucleotide 5604 (3' primer carrying an XbaI site
following a STOP codon). For NuMA
tailIIA+NLS, two PCR products were
used: the first extended from the AatII site to position 5598
(3' primer carrying a BglII site), the second extended from
nucleotide 5932 (5' primer carrying a BglII site) to 6306
(3' primer carrying an XbaI site following the endogenous STOP
codon). The introduction of BglII did not modify the amino acid
sequence at the deletion point. For both deletion constructs, the PCR products
were first cloned into pBluescriptKS NuMA using AatII/XbaI,
then transferred into pCDNA3 GFP-NuMA using EcoRV/XbaI. An
overview of the various constructs is given in
Fig. 3E.
|
Expression levels of GFP NuMA constructs were measured by trypsinising and counting transfected HeLa cells from a culture dish, and analysing the levels of GFP signal by quantitative immunoblotting of HeLa extract, using a GFP-specific antibody, a 125I-labelled secondary antibody, and a phosphoimager. Measured amounts of recombinant GFP were loaded on the same gel for calibration. The percentage of transfected cells was analysed by fluorescence microscopy of a glass coverslip with cells grown from the same culture dish, and the variation of NuMA levels was measured with a digital CCD camera (Zeiss Axiocam, Oberkochen, Germany) and Adobe PhotoShop software (Adobe, San Jose, CA).
Secondary structure prediction of NuMA tail IIA was conducted using
PredictProtein software (Rost et al.,
1994).
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Results |
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NuMA binds directly to tubulin
To test the possibility of a direct NuMA-tubulin interaction, we purified
bacterial fusion proteins covering the various domains of Xenopus
NuMA and assayed their binding to tubulin. NuMA has a tripartite structure,
comprising a central -helical rod domain flanked by globular head and
tail domains. Because previous reports pointed towards an interaction between
spindle microtubules and the tail domain of NuMA
(Compton and Cleveland, 1993
;
Maekawa and Kuriyama, 1993
;
Tang et al., 1994
;
Gueth-Hallonet et al., 1996
),
we studied whether tubulin could be isolated from Xenopus egg
extracts by affinity interaction with hexa-histidine tagged fusion proteins of
the first and second half of the NuMA tail domain (tail I and tail II,
respectively) (Merdes et al.,
1996
). As shown in Fig.
2A, only the distal half of the NuMA tail (tail II) was able to
bind to tubulin. To determine whether other regions of the NuMA molecule
possessed any affinity to tubulin, we also purified fusion proteins of the
head domain, as well as a 425 amino acid long region within the rod domain,
and
SNAP as a control protein, usually involved in membrane vesicle
fusion. All proteins were incubated with soluble pure tubulin, and isolated
using magnetic nickel agarose beads. Fig.
2B shows that the tail II region of NuMA has the highest affinity
to tubulin. A relatively high background binding of tubulin to beads alone
complicated this experiment. In a different assay, NuMA tail II fusion protein
bound quantitatively to taxol-stabilized microtubules
(Fig. 2C). This experiment also
demonstrated that the amount of soluble tubulin left in the supernatant was
significantly reduced after NuMA tail II incubation compared with controls
incubating with BSA or no additional protein. We used this assay to incubate
taxol-stabilized microtubules with increasing concentrations of NuMA tail II,
to estimate the binding constant under saturating conditions. Scatchard plot
analysis (Fig. 2E) revealed an
affinity constant KA of 4x106
M-1 (±2), which is in good agreement with values previously
published for microtubule associated proteins
(Andersen et al., 1994
;
Butner and Kirschner, 1991
).
Because putative cdc2 kinase phosphorylation sites had been reported in the
NuMA tail (Compton and Luo,
1995
), we tested the microtubule-binding properties of NuMA tail
II treated with recombinant cdc2/cyclinB protein. NuMA tail II was efficiently
phosphorylated using [
-32P]ATP (not shown), and was pelleted
with taxol-stabilized microtubules in a competition assay, at increasing
concentrations of unphosphorylated NuMA. Consistent with our data on equal
microtubule binding by interphase or metaphase NuMA (see above), this assay
revealed that phosphorylation by cdc2 kinase did not increase NuMA binding
(Fig. 2F), but led to a small
reduction of the microtubule affinity compared with unphosphorylated NuMA tail
II. This reduction was only about 1.6-fold and sometimes within the variance
of the experiment. The NuMA tail II fragment also had a striking morphological
effect on microtubule organization: when polymerized from phosphocellulose
purified tubulin, thick cables of microtubules formed in the presence of NuMA
tail II (Fig. 2D), each
containing parallel bundles of multiple microtubules, as shown previously by
electron microscopy (Merdes et al.,
1996
).
|
A 100-residue region in the NuMA tail induces stable microtubule
bundles in vivo
The observation of microtubule bundles in vitro led us to investigate
whether NuMA tail II had a similar effect on microtubule organization in the
living cell. A tagged form of human NuMA tail II, containing GFP at its
N-terminus, was overexpressed and followed by fluorescence microscopy in HeLa
cells. In interphase, the fusion protein segregated entirely into the nucleus
(Fig. 3A), due to its nuclear
localization signal between amino acids 1970 and 1991 [corresponding to
positions 1984 and 2005 in a longer NuMA isoform (see
Tang et al., 1994;
Gueth-Hallonet et al., 1996
)].
However, when NuMA tail II was further truncated to remove the nuclear
localization signal and all C-terminal amino acids
(Fig. 3E), the resulting
construct tail IIA decorated multiple thick fibres in the cytoplasm of
interphase cells (Fig. 3A). These fibres represented parallel bundles of microtubules, as shown both by
immunofluorescence microscopy and electron microscopy
(Fig. 3A,B). To test whether
tail II also aligned alongside actin-containing stress fibres, we performed
staining with phalloidin and demonstrated that NuMA tail IIA and actin bundles
did not co-localize (Fig. 3D).
We showed that the formation of microtubule bundles was a direct effect of the
NuMA tail IIA expression and not mediated by endogenous full-length NuMA,
which localized entirely to the nucleus and was not present in the cytoplasmic
fibres (Fig. 3C. In this
experiment, endogenous NuMA was detected with the rod-specific antibody 1F1
(Compton et al., 1991
), which
does not crossreact with the NuMA tail IIA construct. Other regions of NuMA
such as the distal half of the tail (tail I) had no effect on microtubule
organization (Fig. 3A). Also,
further truncation of tail IIA did not produce any fusion proteins capable of
microtubule binding (data not shown), suggesting that tail IIA defines the
minimal domain necessary for microtubule binding and bundling.
The microtubule bundles formed in the presence of NuMA tail IIA were
unusually stable and resisted prolonged cold treatment
(Fig. 4A). Consistent with
this, the bundles stained positively with an antibody against acetylated
tubulin (Fig. 4B), a previously
characterized marker for stable microtubule arrays
(Webster and Borisy,
1989).
|
We measured the expression levels of GFP NuMA tail IIA causing this
phenotype by quantitative immunoblotting and by fluorescence microscopy, and
determined a range between 0.1 and 1 pg protein per cell, equivalent to 2-20
million copies per cell. This range is between 10 and 100 times more than the
number of copies of endogenous NuMA per cell
(Compton et al., 1992). We
found that the strongest effects on microtubule bundling were obtained above
0.3 pg GFP NuMA tail IIA per cell.
The NuMA tail IIA fragment induces abnormal spindle poles
In mitosis, the 100-residue region of NuMA tail IIA caused the formation of
abnormal spindle poles when overexpressed. A variety of phenotypes was
observed: in 47% of the cells (n=150), additional spindle poles were
observed (Fig. 5A) that also
contained endogenous, full length NuMA
(Fig. 5B); in 27% of the cells,
bipolar spindles formed with one of two poles being unusually large, and with
mono-oriented chromosome pairs grouped around the larger pole; in 14% of the
cells, virtually no pole separation could be seen, resulting in astral
microtubule arrays with rosettes of mono-oriented chromosomes. Only 12% of the
cells overexpressing tail IIA displayed apparently normal bipolar spindles.
Identical phenotypes were seen when the longer, 234-residue construct tail II
was overexpressed. The formation of asymmetric spindles with enlarged polar
microtubule asters or additional poles was reminiscent of mitotic cells
treated with taxol-related drugs (Paoletti
et al., 1997). As with taxol, this effect of tail IIA may be
explained by local microtubule stabilization. Similar to previous findings
with bacterially expressed NuMA tail II
(Merdes et al., 1996
), the
shorter fusion protein of NuMA tail IIA was able to induce large microtubule
asters when added to metaphase Xenopus egg extracts
(Fig. 5C). Any tail fragments
lacking the 100 amino acids of tail II, such as tail I or tail IIB, localized
diffusely in metaphase cells, without effects on spindle pole organization or
any other aspects of cell division (Fig.
5A).
|
Without tail IIA, NuMA can no longer bind to microtubules by
itself
Whereas full-length wild-type NuMA concentrates quantitatively in the
nucleus (Fig. 3C), mutant NuMA
in which the nuclear localization sequence has been rendered unfunctional,
accumulates in the cytoplasm and has a severe effect on microtubule
organization. In previous studies (Saredi
et al., 1996; Gueth-Hallonet
et al., 1996
; Gueth-Hallonet
et al., 1998
), such mutant protein was seen in cytoplasmic
aggregates with a fibrous substructure and colocalized with large amounts of
aggregated tubulin polymers. In a similar experiment in this report, tubulin
aggregates were induced by full-length NuMA in which the nuclear localization
signal was deleted (Fig. 6A,
top row), but did not form when the tubulin binding region was deleted. Both
deletion of the entire tail II region at the C-terminus (
tail II), as
well as specific removal of the nuclear localization sequence plus the
100-residue region of tail IIA (
tail IIA+NLS) produced NuMA molecules
that failed to concentrate tubulin during interphase
(Fig. 6A, middle and bottom
rows). However, in mitosis, both mutant forms of NuMA were still able to bind
to the spindle and to concentrate at the poles
(Fig. 6B), presumably by
interaction with endogenous full-length NuMA.
|
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Discussion |
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A surprising finding was that NuMA tail IIA not only decorated microtubules
along their length, but also massively induced the formation of strong
microtubule bundles, both in cells and in microtubule polymerization assays in
vitro (see also Merdes et al.,
1996). The induction of bundles could be explained either by the
existence of multiple tubulin binding sites within the 100 amino acid region
of NuMA tail IIA, or by the ability of multiple NuMA tail IIA polypeptides to
bind each other. According to the first model, this small region would have to
contain two tubulin binding sites, separated by sufficient linker sequence to
bridge two adjacent microtubules. The second scenario seems more likely: NuMA
has been shown to form large fibrous networks
(Saredi et al., 1996
;
Gueth-Hallonet et al., 1998
;
Harborth et al., 1999
) that
are based on dimerization of the NuMA rod domains and on the subsequent
association of multiple NuMA dimers via their tail domains. Harborth et al.
reported that the binding of multiple NuMA tail domains to each other does not
depend on the last 112 amino acids of the tail, indicating that
oligomerization is largely mediated by tail I or tail IIA
(Harborth et al., 1999
). Thus,
the binding of multiple tail IIA polypeptides to each other could well mediate
the association of parallel microtubules. Moreover, the formation of spindle
poles during cell division could be explained in two steps: first, the polar
accumulation of NuMA driven by dynein/dynactin
(Merdes et al., 2000
) and,
second, the direct binding of the NuMA tail domain to the microtubule surface,
whereby networks of multiple NuMA dimers linked at their tail domains would
maintain a focussed array of spindle fibres. The binding of NuMA to the
microtubule surface could provide an additional step of regulating spindle
dynamics by preventing uncontrolled disassembly of microtubule minus ends and
by increasing the stability of the mitotic apparatus, as shown by our finding
that NuMA tail IIA increases microtubule stability.
It is still unclear which mechanisms regulate NuMA binding to the spindle
and what causes the release of NuMA from the microtubule ends and its
re-import into nuclei during telophase. Phosphorylation of NuMA by cdc2/cyclin
B has been suggested to affect binding to spindle microtubules
(Compton and Luo, 1995;
Gaglio et al., 1995
). We have
tested this possibility directly by phosphorylating NuMA tail II in vitro with
cdc2 kinase, but could not detect major differences in microtubule binding
affinity. Although there is no doubt that full length NuMA is phosphorylated
during mitosis (Sparks et al.,
1995
; Gaglio et al.,
1995
; Compton and Luo,
1995
), details on the regulation through specific kinases still
remain to be investigated. It is possible that a large protein such as NuMA,
with multiple potential phosphorylation sites in various domains, is regulated
by more than one kinase. Furthermore, the regulation of NuMA binding to
microtubules might involve additional factors and might not be directly
affected by phosphorylation. Given that overexpressed NuMA tail IIA as well as
full-length NuMA in frog egg extracts can bind avidly to microtubules both in
interphase and mitosis, participation of other factors seems to be the most
likely explanation. One recently suggested mechanism involves the regulation
by importin ß and Ran GTP (Wiese et
al., 2001
; Nachury et al.,
2001
). Recent reports showed that the tail II region of NuMA can
bind and focus microtubules into mitotic asters after an inhibitor, importin
ß, is released from NuMA by Ran GTP. Consequently, at the exit of
mitosis, NuMA could re-associate with importin ß, detach from the spindle
and subsequently get transported into the nucleus. Using our microtubule
pelleting assay, we tested whether importin
or ß prevented NuMA
tail II binding to microtubules, but were unable to detect any effect (A.M.,
unpublished). During the course of this study, another binding partner of
NuMA, the protein LGN, has been identified and its interaction domain
characterized (Du et al.,
2001
): LGN binds to NuMA amino acids 1818-1930, which largely
overlap with the microtubule binding site. Most interestingly, LGN seems to
negatively regulate the interaction between NuMA and microtubule asters and
might therefore be an important factor during mitotic spindle organization. Of
course, the mechanisms that regulate microtubule aster formation in vivo might
be far more complex, and other components in addition to importin
,
ß, LGN and NuMA are currently being identified
(Gruss et al., 2001
). Based on
the present work, we propose that NuMA can organize microtubules by a direct
interaction, and that the segregation of NuMA into the nucleus after mitosis
is necessary to prevent interference with the microtubule network.
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Acknowledgments |
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