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J. Biol. Chem., Vol. 277, Issue 25, 22718-22724, June 21, 2002
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From the Dynamique du Cytosquelette, Laboratoire d'Enzymologie et
Biochimie Structurales, CNRS, 91198 Gif-sur-Yvette, France
Received for publication, December 5, 2001, and in revised form, April 11, 2002
Stathmin is a phosphorylation-regulated
tubulin-binding protein. In vitro and in vivo
studies using nonphosphorylatable and pseudophosphorylated mutants of
stathmin have questioned the view that stathmin might act only as a
tubulin-sequestering factor. Stathmin was proposed to effectively
regulate microtubule dynamic instability by increasing the frequency of
catastrophe (the transition from steady growth to rapid
depolymerization), without interacting with tubulin. We have used a
noninvasive method to measure the equilibrium dissociation constants of
the T2S complexes of tubulin with stathmin,
pseudophosphorylated (4E)-stathmin, and diphosphostathmin. At both pH
6.8 and pH 7.4, the relative sequestering efficiency of the different
stathmin variants depends on the concentration of free tubulin,
i.e. on the dynamic state of microtubules. This control is
exerted in a narrow range of tubulin concentration due to the highly
cooperative binding of tubulin to stathmin. Changes in pH affect the
stability of tubulin-stathmin complexes but do not change stathmin
function. The 4E-stathmin mutant mimics inactive phosphorylated
stathmin at low tubulin concentration and sequesters tubulin almost as
efficiently as stathmin at higher tubulin concentration. We
propose that stathmin acts solely by sequestering tubulin,
without affecting microtubule dynamics, and that the effect of stathmin
phosphorylation on microtubule assembly depends on tubulin critical concentration.
Microtubules are dynamic polymers that play a role in cell
morphology and cell division. The dynamics of microtubule assembly is
finely regulated during the cell cycle (1-3). In interphase, microtubules are mainly organized in a radial array, with minus ends
anchored at the centrosome; however, a fraction of the population has
two free ends (4-7). Upon entry into mitosis, microtubules disassemble
and then reassemble into a highly dynamic mitotic spindle, with minus
ends at the poles. Finally, at the end of mitosis, disassembly of the
mitotic microtubules is balanced by the formation of the interphase
array in the daughter cells (8).
In living cells, two main processes, treadmilling (6, 7, 9) and dynamic
instability (10), are responsible for monomer-polymer exchange
reactions leading to microtubule turnover. Both are driven by the
hydrolysis of GTP linked to tubulin assembly. Treadmilling derives from
the energetic imbalance between the plus and the minus ends of
microtubules and operates when the two ends are free. The steady-state
concentration of dimeric GTP-tubulin allows equal net rates of assembly
at the plus end and disassembly from the minus end. Dynamic instability
concerns microtubules that have either two or only one free end (the
plus end, generally), which switches infrequently between a rapidly
depolymerizing state and a growing state. The transitions between the
two states, called "catastrophe" and "rescue," describe the
stochastic loss and gain of a GTP cap at the end of the microtubule.
The steady-state concentration of dimeric GTP-tubulin in this case is
determined in part by the frequencies of catastrophe and rescue.
Many cellular factors affect the dynamics of monomer-polymer exchange
(see Ref. 11 for a review). Microtubule-associated proteins slow down
microtubule depolymerization in a phosphorylation-controlled fashion
(12). Kinesins of the KIF family bind to microtubule ends and catalyze
depolymerization (13), whereas end stabilizers like XMAP215 prevent
depolymerization (14). As a result of these activities, the
steady-state concentration of dimeric tubulin coexisting with
microtubules is regulated. The microtubule-severing factor katanin
modulates the fraction of microtubules with one or two free ends (15,
16). Microtubule dynamics in a living cell depends on the proportion of
microtubules with one or two free ends (17, 18), hence katanin also
affects the steady-state concentration of free GTP-tubulin by changing
the relative contributions of treadmilling and dynamic instability.
Thus far, it has not been technically possible to evaluate the
steady-state concentration of GTP-tubulin in cells with great accuracy.
Nonetheless, when the fraction of free minus ends increases,
e.g. by detachment of microtubules from centrosomes, the
concentration of free GTP-tubulin is expected to increase from a value
close to the critical concentration of the plus end to a value closer
to the critical concentration of the minus end. This shift has actually
been observed (18), supporting the view that the concentration of
GTP-tubulin may vary in vivo.
In contrast with regulatory factors that control microtubule assembly
dynamics, tubulin-sequestering factors bind tubulin in a
nonpolymerizable complex. These proteins establish a pool of
unassembled tubulin, built at the expense of the microtubule pool and
in equilibrium with free tubulin at its steady-state concentration. The
concentration of sequestered tubulin is therefore governed by the
dynamic state of microtubules.
Op18/stathmin is a 17-kDa protein that has been recently revealed to
bind tubulin and destabilize microtubules (19) and is negatively
regulated by phosphorylation by a variety of kinases (20-25). Stathmin
plays a crucial role in cell division (see Ref. 26 for review). It is
phosphorylated to a low basal level in interphase and becomes
hyperphosphorylated by cyclin-dependent kinases (27) and a
polo-like kinase (28) upon entry into mitosis. Progression through the
cell cycle requires phosphorylation of all four serines (Ser-16, -25, -38, and -63). The phosphorylation level is regulated by protein
phosphatase 2A phosphatase during mitosis (29), and
dephosphorylation occurs at the end of mitosis (30). Stathmin
sequesters tubulin in a T2S complex in which it interacts
with two In contrast with the above-mentioned studies, much larger differences
in affinity for tubulin were observed between wild-type, 4E-mutated,
and phosphorylated stathmins when their effects on nucleotide exchange
on tubulin were measured (44). We thought that the discrepancies
regarding stathmin function might originate from the lack of a
quantitative evaluation of the tubulin-sequestering activity of the
different stathmin derivatives. Here we set up a sequestration assay in
which the concentration of free GTP-tubulin coexisting with
microtubules at steady state is buffered to any desired value using
Taxotere. Using this assay, the affinity of tubulin for the different
stathmins differs to a greater extent than in previous measurements.
Phosphorylation exerts a regulatory effect on stathmin depending on the
concentration of free tubulin. We conclude that the simple sequestering
activity of stathmin can support its effects on microtubules in
vivo. We tentatively propose that the changes in microtubule
dynamics during the cell cycle are associated with variations in the
concentration of free tubulin that coexists with microtubules. Changes
in the concentration of free tubulin in turn modulate the effect of
phosphorylation of stathmin on its sequestering activity.
Proteins--
Phosphocellulose-purified bovine brain tubulin
(44) was used. The tubulin used in this work had been kept at
Recombinant wild-type stathmin and 4E-stathmin were expressed in
Escherichia coli and purified as described previously (43). Stathmin was phosphorylated on serines 16 and 63 by protein kinase A
(Sigma) as described previously (44).
Determination of the Equilibrium Dissociation Constant for the
T2S Complex--
Spontaneous polymerization of tubulin in
microtubules was monitored turbidimetrically at 350 nm in a Cary 1 spectrophotometer using a 1-cm path, 120-µl cuvette thermostated at
37 °C. Experiments were carried out in either glycerol-containing M
buffer or glycerol-free P buffer (0.1 M PIPES-KOH, pH 6.8, 0.5 mM EGTA, 0.5 mM GTP, and 6 mM
MgCl2) or glycerol-free H buffer (100 mM
HEPES-KOH, pH 7.4, 0.5 mM EGTA, 0.5 mM GTP, and
6 mM MgCl2) in the absence or presence of
Taxotere or stathmin as indicated. In preliminary assays, the range of
stathmin concentrations was selected to cause at most a 10% decrease
in the mass of microtubules (see "Results"). Polymerization was
started by the addition of MgCl2 and Taxotere to the
tubulin + stathmin solution that was immediately brought into the
prewarmed cuvette. The temperature reached 37 °C in less than
15 s. Critical concentration plots were derived from turbidity
measurements as described previously (31). Parallel samples were
polymerized identically in centrifuge tubes placed in a water bath at
37 °C and then centrifuged at 300,000 × g for 15 min at 37 °C in a TL 100 ultracentrifuge (Beckman). The supernatants
were denatured and subjected to SDS-PAGE electrophoresis for evaluation
of the amount of unassembled tubulin. In the absence of stathmin, the concentration of unassembled tubulin equaled the critical concentration [T]SS. In the presence of stathmin, the concentration of
unassembled tubulin was [T]U = [T]SS + 2 [T2S]. A series of tubulin standards in the appropriate
range were electrophoresed on the same gel. Gels were stained
with either Coomassie Blue or silver (45), depending on the amounts of
tubulin present in the supernatants. Gels were scanned and analyzed
using NIH Image software. The amounts of nonassembled tubulin in the
samples were determined by interpolation using the calibration curve
obtained with standards spanning the range of tubulin
concentrations found in the samples. The value of the equilibrium
dissociation constant for the T2S complex was determined as
follows at different total concentrations of stathmin [S]0.
Background: Cooperative Binding of Tubulin to
Stathmin--
Stathmin interacts with two
The general binding scheme of tubulin to stathmin can therefore be
described by an isoenergetic square model (Fig.
1) in which the intermediate 1:1
complexes TS and ST interact strongly with a second tubulin molecule,
leading to T2S. The TS and ST complexes account for the
binding of tubulin to the NH2-terminal or the COOH-terminal
Alternatively (Fig. 1b), stathmin can formerly bind strongly
to a poorly represented
The kinetic mechanism of binding of tubulin to stathmin is not known.
The TS, ST, and TT species are putative kinetic intermediates that play
an important role in the pathway leading to T2S but are not
detected at equilibrium because of their low stability (44).
Equilibrium binding of tubulin to stathmin is described by an extremely
cooperative scheme.
Sequestration of Tubulin by Stathmin--
In a solution of
microtubules at steady state, the concentration of free tubulin is
maintained at a steady-state value [T]SS (often called
critical concentration) that depends on the dynamics of microtubules
under these solution conditions. When stathmin is added to microtubules
at steady state, it binds to tubulin, and because the resulting
T2S complex is nonpolymerizable, microtubules depolymerize
to maintain the concentration of free tubulin equal to
[T]SS. The T2S complex is formed at the
expense of the microtubule pool, without affecting the value of
[T]SS. Depolymerization stops, and steady state is again
established when both the remaining microtubules and T2S
complex coexist with free tubulin at the unchanged concentration
[T]SS. The amount of T2S complex at steady state is determined by the values of KD and
[T]SS as follows.
Note that if stathmin is added at a concentration high enough to cause
complete depolymerization of microtubules, the concentration of free
tubulin is no longer buffered by microtubules, hence Eq. 5 is no longer
valid. The relevant description then is the binding and saturation of
tubulin by stathmin, and a cubic equation describes the dependence of
[T2S] on the concentrations of total tubulin and total stathmin.
According to Eq. 5, the value of KD can easily be
derived from measurements of the amount of sequestered tubulin upon
addition of stathmin to a solution of microtubules at steady state (see
"Materials and Methods"). In fact, in the conventional polymerization buffers used for microtrubule assembly in
vitro, all the added stathmin or 4E-stathmin or diphosphostathmin
was found in complex with tubulin (data not shown), in agreement with previous reports (31). This result indicates that the value of
[T] Taxotere Buffers Free Tubulin at a Low Concentration, Allowing
Measurement of KD--
Differences in the value of
KD for the different stathmin variants would be
revealed if the value of [T]SS could be lowered to a
value closer to KD1/2. Taxotere can
fulfill this function because this microtubule-stabilizing drug binds
specifically to microtubules with an affinity of 2 × 107 M
To obtain the value of KD for the different stathmin
variants, the dependence of [T]SS on Taxotere
concentration was established. Experiments were done in glycerol-free P
buffer, in which the critical concentration can be varied from 15 µM in the absence of Taxotere to <0.25 µM
at high Taxotere concentrations. Tubulin was polymerized at 40 and 15 µM in P buffer in the presence of different
concentrations of Taxotere, [X]0. The relationship between the concentration of tubulin in the supernatants of sedimented microtubules, [T]SS, and the concentration of free
Taxotere, [X], was established as follows. The concentration of
assembled tubulin ([MT]0) was derived from the difference
between the total ([T]0) and free ([T]SS)
tubulin concentrations. The concentrations of microtubule-bound
Taxotere [MTX] and free Taxotere [X] were calculated as
follows:
The value of [T]SS decreased with free Taxotere (Fig.
2). The curves representing
[T]SS versus [X] obtained at the two
concentrations of tubulin tested (
The fact that the curves obtained at 40 and 15 µM tubulin
are superimposable, within an experimental error of 0.2 µM tubulin at saturating Taxotere concentration, provides
an estimate of the maximum fraction of tubulin that was
unpolymerizable. Namely, if 10% tubulin was inactive and
unpolymerizable in the supernatants, then the amount of soluble tubulin
would be at least 4 µM at 40 µM tubulin and
at least 1.5 µM at 15 µM tubulin, whereas
much lower values were measured. Second, the difference in the
concentrations of tubulin in the supernatants at 40 and 15 µM tubulin would be 4 µM
Fig. 2 shows that in the presence of 3 µM stathmin, the
amount of sequestered tubulin decreased as the concentration of
Taxotere increased. In contrast, no appreciable sequestration of
tubulin by 3 µM diphosphostathmin was observed even in a
range of low Taxotere concentration. In conclusion, Taxotere reveals
the difference in KD between stathmin and diphosphostathmin.
The differences in the sequestering activities of wild-type stathmin,
4E-stathmin, and diphosphostathmin can be quantitated at different
Taxotere concentrations (Fig. 3). The
amounts of sequestering protein added to microtubules were such that
<10% of the microtubules disassembled. In doing so, the saturation level of microtubules by Taxotere was not greatly affected by the
partial depolymerization. Hence, the concentration of free tubulin
could be considered identical in the absence and presence of
sequestering agent.2 The
values of KD were derived from the measurement of unassembled tubulin in the absence and presence of stathmin at different concentrations, using Eq. 4 (see "Materials and
Methods"). Typical data are shown in Fig.
4. Values of KD are
summarized in Table I, together with the value of the concentration of
free tubulin at which 50% of the stathmin is in complex with
tubulin ([T]1/2 = KD1/2).
Comparison of the values of KD obtained for the
different stathmin derivatives at pH 6.8 or pH 7.4, in the presence or
absence of glycerol, leads to the following conclusions.
In glycerol-free P buffer (pH 6.8), the value of KD
is 750-fold higher for diphosphostathmin than for stathmin. This corresponds to a 30-fold difference (0.3 µM for stathmin
and 9 µM for diphosphostathmin) in the [T]1/2 values. The value of KD for 4E-stathmin is only
40-fold higher than that for stathmin, indicating that the serine to
glutamate mutation only partially mimics phosphorylation, in agreement
with previous works (31, 36, 44). In glycerol-containing buffer, the
ratios between the values of KD for the different stathmin variants are 1 order of magnitude lower than in the absence of
glycerol. The value of [T]1/2 for stathmin is about 5-fold
lower than that in the absence of glycerol, indicating that the
affinity of stathmin is enhanced by glycerol, consistent with the
existence of hydrophobic contacts between the two proteins in the
T2S complex (47).
In glycerol-free H buffer (pH 7.4), the values of [T]1/2 for
all stathmin variants are 3-fold higher than those at pH 6.8 (Table I).
Therefore, the ratios of [T]1/2 of 4E-stathmin and
diphosphostathmin to [T]1/2 of stathmin are not significantly different from those found at pH 6.8. In conclusion, the relative tubulin-sequestering efficiencies of 4E-stathmin and diphosphostathmin are not affected by pH.
The general conclusion of these experiments is that phosphorylation of
stathmin affects its tubulin-sequestering activity in a manner that
depends on the concentration of free tubulin maintained in the medium.
As an example, the values of KD found here for
stathmin, 4E-stathmin, and diphosphostathmin have been used to
calculate the amount of tubulin sequestered by stathmin or its variants
(set at 6 µM, a plausible cellular value) at different concentrations of free tubulin using Eq. 5, so as to mimic virtual cellular situations in which these proteins have been used (Fig. 5). At low concentration of free tubulin,
a large difference is observed in the amounts of tubulin
sequestered by stathmin, 4E-stathmin, and diphosphostathmin, whereas
the differences tend to vanish at high tubulin concentration. At low
concentration of free tubulin, 4E-stathmin has a low sequestering
activity, like diphosphostathmin. In contrast, at high concentration of
free tubulin, 4E-stathmin sequesters tubulin as efficiently as stathmin
and differs from diphosphostathmin.
We have developed a novel, non-equilibrium-perturbing assay to
measure the equilibrium dissociation constant of the tubulin stathmin
complex and to quantitate the differences in stability of the tubulin
complexes formed with stathmin, 4E-stathmin, and diphosphostathmin. The
same assay could be used for other stathmin variants and fragments. The
value of KD is increased by 3 orders of magnitude by
phosphorylation of stathmin and <100-fold by serine to glutamate
mutation at pH 6.8 and at pH 7.4. Nucleotide exchange measurements (44)
in glycerol-free buffer at 20 °C indicated that the stathmin-tubulin
complex dissociated very slowly, consistent with a value of
[T]1/2 around 0.02 µM, 1 order of magnitude
lower than the range of 0.1-0.3 µM measured here at
37 °C. The estimated value of [T]1/2 for diphosphostathmin
at 20 °C was, in contrast, only 2-fold lower than the one found here
at 37 °C. It is possible that the difference in stability of the
complexes of stathmin and diphosphostathmin with tubulin varies greatly
with temperature. The uncertainty in the measure of
KD can be evaluated as follows. As discussed earlier
in this paper, a maximum of 1% of tubulin may be inactive in the
supernatant of Taxotere-stabilized microtubules. Then, at the high
concentration of Taxotere used to measure KD for
stathmin, the actual concentration of free tubulin able to undergo
monomer-polymer exchange may be overestimated by 50%. In P buffer (not
in H buffer), this uncertainty limits our estimate of
KD to a range of 0.01-0.15
µM2, corresponding to a range of
[T]1/2 of 0.1-0.4 µM, which is reasonably
accurate for a thermodynamic parameter. The interference of 1%
inactive tubulin has less bearing on the values of
KD for 4E-stathmin and diphosphostathmin that are
measured at a higher concentration of free tubulin. At pH 7.4, the
value of [T]1/2 for stathmin is 3-fold higher than that at pH
6.8 but still lower than the one (2.5 µM) derived from
equilibrium-perturbing measurements (41) at the same pH. Comparison of
the values of [T]1/2 for stathmin and its derivatives shows
that stathmin is 6-7-fold more efficient than 4E-stathmin and
20-40-fold more efficient than diphosphostathmin, independently of pH.
These differences are larger than those in previous estimates (41).
Using plasmon resonance (47), tubulin-stathmin interaction was
expressed in terms of binding of The cooperative binding of tubulin to stathmin allows regulation by
phosphorylation in a narrow concentration range of free tubulin. This
range is narrower at pH 6.8 than at pH 7.4; however, the general
conclusion is valid in the whole range of physiological pH. The graphs
in Fig. 5 suggest that in vivo, under circumstances where
the steady-state concentration of free tubulin that coexists with
microtubules is low, a fine regulation of the sequestering function of
stathmin can be obtained. Desequestration of tubulin (i.e.
microtubule assembly) may be elicited in two ways, either by a decrease
in [T]SS or by phosphorylation of stathmin.
In view of these results, we propose that the variability in the
compared effects of 4E-stathmin and diphosphostathmin in living cells
can be accounted for in terms of dependence of the sequestration
activity on free GTP-tubulin concentration. Recently, 4E-stathmin has
been observed to depolymerize interphasic microtubules but not mitotic
microtubules. The catastrophe-promoting activity of stathmin has been
postulated to be required to disrupt the mitotic spindle but not
interphasic microtubules. An alternative interpretation of the
different effects of 4E-stathmin on interphasic and mitotic
microtubules, based on the present results (Fig. 5), is that the
concentration of free GTP-tubulin is lower in mitosis than in
interphase, hence in a range of low concentrations the sequestering
activity of 4E-stathmin is measurable only in interphase (we expect
that at higher concentrations, 4E-stathmin would be effective in
mitosis). Is this interpretation acceptable given the current knowledge
of microtubule dynamics in the cell cycle? Within experimental error,
the mass amount of microtubules in the mitotic spindle is practically
as large as that in interphase (8), but the amount of unassembled
tubulin is probably distributed in different pools in mitosis and in
interphase. For instance, due to dynamic instability of mitotic
microtubules, GDP-tubulin may represent a large fraction of the pool of
nonmicrotubular (perhaps oligomeric) tubulin in mitosis and a smaller
fraction in interphase. The minus end of mitotic microtubules is
anchored at the poles, hence the concentration of GTP-tubulin must
equal the low critical concentration at the plus end in mitosis.
Modeling of dynamic instability (48) expresses that the action of
cyclin A- and cyclin B-dependent kinases on the catastrophe
frequency (49) maintains the activity of free GTP-tubulin at a level
low enough for the average microtubule growth rate to be
negative. In contrast, interphasic microtubules treadmill;
therefore, the steady-state concentration of GTP-tubulin is maintained
at a value intermediate between the plus end and minus end critical
concentrations. The rate of growth at the plus end is as high as 12 µM/min (18), indicating that the concentration of
GTP-tubulin is high. In conclusion, although no direct proof exists in
support of our interpretation, at least it is not in disagreement with
available data.
The catastrophe-promoting activity of stathmin has been postulated to
solve apparently conflicting results on the function of stathmin
in vitro and in vivo. However, these results were perceived as conflicting because the effects that result from simple
sequestration have not been fully appreciated. Direct evidence supporting a catastrophe-promoting activity of stathmin is in fact
lacking. A catastrophe-promoting factor is expected to bind to
microtubule ends and enhance microtubule turnover and accompanying steady-state GTPase activity, which has not been observed (31). As
recently pointed out (28), no evidence has been provided for binding of
stathmin to microtubule ends in vivo or in vitro. In our hands, immunodetection failed to detect significant binding of
stathmin to sedimented Taxol-stabilized microtubules. Identical interstitial low amounts (about 5% of total stathmin or
diphosphostathmin) were measured in microtubule pellets. The same
amount was measured for long or fragmented microtubules, inconsistent
with specific binding of stathmin to microtubule ends (see Fig.
1S).
Stathmin was thought to promote catastrophe because when added to
interphasic Xenopus egg extracts supplemented with
centrosomes, it reduced the proportion of microtubules extending long
bidimensional sheets, typical of the growing state, and increased the
proportion of rapidly depolymerizing microtubules with frayed ends
(50). However, these assays are not steady-state measurements of
microtubule dynamics. Microtubules grow indefinitely from free tubulin
in interphasic extracts (49). Measurements of microtubule length and
end structure are performed at a given time after centrosome addition
(microtubules would be longer at a later time). Hence, length
measurements are equivalent to measurements of initial rate of growth
at a given concentration of free tubulin. Addition of stathmin to
interphasic extracts lowers the concentration of free tubulin,
therefore the rate of microtubule growth decreases and the frequency of
catastrophe consistently increases, as when measurements are done with
pure tubulin (51). In this case, as in in vitro assays (43),
stathmin indirectly promotes catastrophe by sequestering tubulin.
A catastrophe-promoting activity of stathmin has also been postulated
(40) to explain that overexpression of truncated stathmin derivatives,
which bind tubulin less tightly than wild-type stathmin, still results
in microtubule destabilization. The expectation that stathmin competes
with the truncated fragments results from the incorrect assumption that
the pool of free dimeric tubulin in living cells is not in a dynamic
monomer-polymer exchange state with microtubules. In fact, if two
stathmin variants, say S and S', which form T2S and
T2S' complexes with tubulin, with thermodynamic parameters
KD and K'D are added together to microtubules, they act in an additive fashion, not in competition. The
amount of sequestered tubulin is given by an extension of Eq. 5:
*
This work was supported in part by the Ligue Nationale
contre le Cancer.The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
Published, JBC Papers in Press, April 15, 2002, DOI 10.1074/jbc.M111605200
2
Under different conditions, for instance at
lower Taxotere concentration ([X]0) and high extent of
depolymerization of microtubules (high stathmin concentrations),
depolymerization of microtubules would be accompanied by increased
saturation of the remaining microtubules by Taxotere, causing an
increased polymer stability, i.e. a decrease in
[T]SS. Then, stathmin would have less and less sequestering efficiency at higher concentration. In vivo, a
similar effect could be mediated by microtubule-associated proteins
rebinding to remaining microtubules. In vitro, this
situation can be handled quantitatively, using the curve representing
the change in [T]SS versus free Taxotere (Fig.
2), but here we preferred to work under simpler conditions, where no
correction would have to be brought. This simplification is validated
by the fact that the same values of KD are found at
different concentrations of stathmin.
The abbreviations used are:
MES, 4-morpholineethanesulfonic acid;
PIPES, 1,4-piperazinediethanesulfonic
acid.
The Effect of Stathmin Phosphorylation on Microtubule Assembly
Depends on Tubulin Critical Concentration*,
ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
-tubulin heterodimers (31) in a polar -
arrangement (see Ref. 11 for review; Refs. 32 and 33). Recent chemical
cross-linking data indicate that the NH2-terminal region of
stathmin is at the end of the - dimer (34). Whether the
biological function of stathmin is supported by its tubulin-sequestering activity only or by some additional
catastrophe-promoting activity, independent from tubulin binding, is
not understood yet (see Ref. 35 for review). Conflicting results have
been obtained using pseudophosphorylated mutated stathmin (4E-stathmin) in which all four phosphorylatable serines were replaced by glutamate. The 4E-stathmin showed unaltered tubulin sequestering activity, yet it
failed to destabilize microtubules in vivo (36-38). Mutants affected in the coiled-coil domain of stathmin interacted with tubulin
like wild-type stathmin but failed to destabilize microtubules in
leukemia cells (36). When injected in living cells, NH2- and COOH-terminal-truncated fragments of stathmin also had different effects on the microtubule lattice, suggesting that different activities of stathmin were carried by different regions of the protein
(39, 40). Finally, 4E-stathmin, which seems to lack catastrophe-promoting activity and retain unaltered
tubulin-sequestering activity, is able to disrupt interphase
microtubules but not mitotic microtubules (41). Consistently,
4E-stathmin does not prevent normal development of Xenopus
embryo (42). Quantitative measurements of tubulin binding to wild-type
stathmin and mutated stathmin (4E-stathmin) or phosphorylated stathmin,
using plasmon resonance (43) or a pull-down assay (40, 41), showed that
the affinity of stathmin for tubulin was decreased only 3-4-fold by
the serine to glutamate mutation and that phosphorylated stathmin had
<2-fold lower affinity for tubulin than 4E-stathmin. It was thought
(40, 41) that these modest differences in affinity could not account for the large differences in the effects of the various stathmin variants on microtubules in cells.
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
80 °C for at most 3 weeks. Older preparations showed a measurable
amount of nonpolymerizable material, which increased with aging.
Before each experiment, tubulin was recycled by polymerization at
37 °C in M buffer (50 mM
MES1-KOH, pH 6.8, 4 M glycerol, 0.5 mM EGTA, 0.5 mM
GTP, and 6 mM MgCl2). Microtubule pellets were
resuspended in M buffer containing 0.5 mM MgCl2
on ice and centrifuged at 200,000 × g at 4 °C for 10 min to remove aggregates. Tubulin was equilibrated in the desired buffer by Sephadex G-25 gel filtration (PD-10; Amersham Biosciences). Experiments were performed immediately after the above-mentioned recycling procedure to avoid denaturation of tubulin. Before each experiment, it was verified that tubulin polymerized in microtubules that depolymerized at least 95% at 4 °C.
![K<SUB>D</SUB>=[<UP>T</UP>]<SUB><UP>SS</UP></SUB><SUP>2</SUP>×[<UP>S</UP>]/[<UP>T<SUB>2</SUB>S</UP>]](/content/vol277/issue25/fulltext/22718/img001.gif)
(Eq. 1)
![[<UP>T<SUB>2</SUB>S</UP>]=([<UP>T</UP>]<SUB><UP>U</UP></SUB>−[<UP>T</UP>]<SUB><UP>SS</UP></SUB>)/2](/content/vol277/issue25/fulltext/22718/img002.gif)
(Eq. 2)
![[<UP>S</UP>]=[<UP>S</UP>]<SUB>0</SUB>−([<UP>T</UP>]<SUB><UP>U</UP></SUB>−[<UP>T</UP>]<SUB><UP>SS</UP></SUB>)/2](/content/vol277/issue25/fulltext/22718/img003.gif)
(Eq. 3)
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
-tubulin heterodimers
in a T2S complex (31-33). At 5 µM tubulin
and at concentrations of stathmin as high as 50 µM, the
T2S complex was the only complex observed in the analytical
ultracentrifuge; no evidence was obtained for an intermediate 1:1 TS
complex (31, 44). This result, corroborated by structural (33) and
biochemical studies (40, 41), suggested that tubulin bound
cooperatively to stathmin, i.e. tubulin-tubulin interactions
as well as stathmin-tubulin interactions were responsible for the high
stability of the T2S complex. The lateral interaction of
the two -tubulin molecules with a tandem of two related
consecutive -helices in stathmin (33) stabilizes the longitudinal
interactions in the - tubulin dimer, under ionic conditions
where tubulin exists only in the -tubulin form.
-helix of stathmin, hence they have different structures (Fig.
1a). Whether only one of the two -helices is sufficient to induce dimerization of tubulin in a T2S complex is not
known.
View larger version (15K):
[in a new window]
Fig. 1.
Scheme for cooperative binding of tubulin to
stathmin. Two possible nonexclusive mechanisms for
formation of the T2S complex are displayed. a,
stathmin binds two molecules of -tubulin consecutively, leading
to T2S via TS or ST intermediates. b,
dimerization of tubulin precedes binding of stathmin. Detailed balance
implies that K1 × aK2 = K2 × aK1 = K × K' = KD = [T]2 × [S]/[T2S]. Parameter a
represents the change in stability due to the tubulin-tubulin
interaction involved in the formation of the T2S complex.
Positive cooperativity in tubulin binding implies that a < 1. Noncooperative binding (independent binding of tubulin to the two
sites on stathmin) would correspond to a = 1, and
negative cooperativity would correspond to a > 1. The
two -helices H1 and H2 form the interface of
stathmin with the two -tubulin heterodimers. The -subunit of
tubulin is identified by an indentation.
- tubulin dimer (TT), leading to T2S.
The overall equilibrium dissociation constant of the
T2S complex is as follows:
where [T] and [S] are the concentrations of free tubulin and
stathmin, and [T2S] is the concentration of the
tubulin-stathmin complex. Irrespective of the pathway leading to
T2S, KD has the dimension of a square
concentration, expressed in M2. A physically
significant parameter is the concentration of free tubulin at which
half of total stathmin is in complex with tubulin at equilibrium ([S] = [T2S]), called [T]1/2. Eq. 4 shows that
[T]1/2 = KD1/2. Because of the
strong cooperativity in stathmin-tubulin interactions, the ratio of the
free tubulin concentrations at which 90% and 10% of stathmin is in
complex with tubulin ([T]90%/[T]10%) equals 9, whereas it would have a value of 81 if binding was noncooperative.
![K<SUB>D</SUB>=[<UP>T</UP>]<SUP>2</SUP>×[<UP>S</UP>]/[<UP>T<SUB>2</SUB>S</UP>]](/content/vol277/issue25/fulltext/22718/img006.gif)
(Eq. 4)
The total concentration of stathmin is [S]0 = [S] + [T2S].
![[<UP>T<SUB>2</SUB>S</UP>]=[<UP>S</UP>]<SUB>0</SUB>×[<UP>T</UP>]<SUP>2</SUP><SUB><UP>SS</UP></SUB>/(K<SUB>D</SUB>+[<UP>T</UP>]<SUP>2</SUP><SUB><UP>SS</UP></SUB>)](/content/vol277/issue25/fulltext/22718/img007.gif)
(Eq. 5)
1 at 37 °C (46) and
lowers the critical concentration.
![[<UP>MT</UP>]<SUB>0</SUB>=[<UP>MT</UP>]+[<UP>MTX</UP>]](/content/vol277/issue25/fulltext/22718/img009.gif)
(Eq. 6)
![K<SUB><UP>X</UP></SUB>=[<UP>MT</UP>]×[<UP>X</UP>]/[<UP>MTX</UP>]](/content/vol277/issue25/fulltext/22718/img010.gif)
(Eq. 7)
![[<UP>MTX</UP>]=1/2{[<UP>X</UP>]<SUB>0</SUB>+[<UP>MT</UP>]<SUB>0</SUB>](/content/vol277/issue25/fulltext/22718/img011.gif)
(Eq. 8)
![+K<SUB><UP>X</UP></SUB>±(([<UP>X</UP>]<SUB>0</SUB>+[<UP>MT</UP>]<SUB>0</SUB>+K<SUB><UP>X</UP></SUB>)<SUP>2</SUP>−4[<UP>MT</UP>]<SUB>0</SUB>×[<UP>X</UP>]<SUB>0</SUB>)<SUP>½</SUP>}](/content/vol277/issue25/fulltext/22718/img012.gif)
where KX represents the equilibrium
dissociation constant for Taxotere binding to tubulin in microtubules.
![[<UP>X</UP>]=[<UP>X</UP>]<SUB>0</SUB>−[<UP>MTX</UP>]](/content/vol277/issue25/fulltext/22718/img013.gif)
(Eq. 9)
and 
) were superimposable, as
expected for the dependence of an equilibrium constant on free ligand
concentration, which should be independent of the total concentration
of tubulin.
View larger version (13K):
[in a new window]
Fig. 2.
Dependence of the critical concentration for
microtubule assembly on Taxotere concentration. Tubulin was
polymerized at 40 µM (closed symbols) or 15 µM (open symbols) in P buffer and
in the absence (circles) or presence of 3 µM
stathmin ( 
) or diphosphostathmin (
). Tubulin in the supernatant
was quantitated using the bicinchoninic acid assay. The concentration
of free Taxotere was calculated using Eq. 7.
1.5 µM = 2.5 µM. The maximal measured
difference at all Taxotere concentrations is 10-fold lower, hence we
can conclude that at most 1% tubulin is nonpolymerizable. For each
preparation of tubulin used in this work, it was routinely checked that
the critical concentration measured at saturating Taxotere was the same
within 0.1 µM. Accordingly, the values measured for
KD were reproducible in triplicate experiments, with
fairly good precision (see Table I). Note
that if a small proportion of tubulin was irreversibly assembled in
nonmicrotubular aggregates, then the concentration of free tubulin in
the supernatant would still represent the concentration of tubulin
undergoing monomer-polymer exchange with microtubules.
Equilibrium dissociation constant of the T2S complex formed
with stathmin, 4E-stathmin, and diphosphostathmin at 37 °C
View larger version (40K):
[in a new window]
Fig. 3.
Effect of Taxotere on the sequestering
activity of the different stathmin variants in P and M buffers.
Top four panels, tubulin was polymerized at 40 µM in P buffer in the presence of the indicated
concentrations of Taxotere and in the absence or presence of stathmin
variants at 3 µM. Bottom two panels, tubulin
was polymerized at 16 µM in M buffer in the absence or
presence of 16 µM Taxotere and in the absence or presence
of stathmin variants at 2 µM. The SDS-PAGE analysis of
the supernatants of microtubules is shown. The gels are stained with
Coomassie Blue.
View larger version (32K):
[in a new window]
Fig. 4.
Stability of T2S complex with
different stathmin variants. Tubulin (15 µM) was
polymerized in the presence of Taxotere at 20 µM
(top panel) or 6 µM (middle and
bottom panels) and with the indicated concentrations of
stathmin (top panel), 4 E-stathmin (middle
panel), or diphosphostathmin (bottom panel). The
concentration of unassembled tubulin in the supernatants is derived
from densitometric analysis of silver-stained gels and comparison with
standards of tubulin electrophoresed on the same gel as indicated. The
arrows indicate the data points on the calibration
curve.
View larger version (17K):
[in a new window]
Fig. 5.
The regulatory effect of phosphorylation on
stathmin function depends on free tubulin concentration. a,
amount of unassembled tubulin as a function of free tubulin
concentration. The concentration of total stathmin is assumed to be 6 µM, as indicated by cell biology data (11, 37). The
concentration of unassembled tubulin ([T] + 2 [T2S]) is
calculated using Eq. 5, and the values of KD were
determined in glycerol-free P buffer for stathmin (circles),
4E-stathmin (triangles), and diphosphostathmin
(squares). Blue lines and closed
symbols, pH 6.8; red lines and open symbols,
pH 7.4 (symbols identify the curves and do not represent
experimental data). b and c, different tubulin
sequestering activity of 4E-stathmin at 1 µM free tubulin
(b) and 10 µM free tubulin (c). The
amount of unassembled tubulin ([T] + [T2S]) is
calculated as a function of the total concentration of stathmin
(circles), 4E-stathmin (triangles), and
diphosphostathmin (squares), emphasizing that the behavior
of 4E-stathmin is similar to that of diphosphostathmin at low tubulin
concentration (b) or closer to that of stathmin at high
tubulin concentration (c). Colors and
symbols are as described in a.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
- dimers (T2)
to immobilized stathmin, i.e. Kd = [T2] × [S]/[T2S]. Comparison of this value of Kd with our estimates of
KD (Kd = KD × [T2]/[T]2) is not straightforward because
the equilibrium dimerization constant of tubulin,
[T2]/[T]2, is not known.
The results (40) are fully consistent with the additive tubulin
sequestration activities of stathmin and its truncated derivatives, and
a catastrophe-promoting activity is not required to account for those data.
![[<UP>T<SUB>2</SUB>S</UP>]+[<UP>T<SUB>2</SUB>S</UP>′]=[<UP>T</UP>]<SUP>2</SUP><SUB><UP>SS</UP></SUB>×{[<UP>S</UP>]<SUB>0</SUB>/(K<SUB>D</SUB>+[<UP>T</UP>]<SUP>2</SUP><SUB><UP>SS</UP></SUB>)+[<UP>S</UP>′]<SUB>0</SUB>/(K′<SUB>D</SUB>+[T]<SUP>2</SUP><SUB><UP>SS</UP></SUB>)}](/content/vol277/issue25/fulltext/22718/img014.gif)
(Eq. 10)
FOOTNOTES
The on-line version of this article (available at
http://www.jbc.org) contains Fig. 1S.
To whom correspondence should be addressed: Dynamique du
Cytosquelette, Laboratoire d'Enzymologie et Biochimie Structurales, 1 Avenue de la Terrasse, CNRS, 91198 Gif-sur-Yvette, France. Tel.: 33-1-69-82-34-65; Fax: 33-1-69-82-31-29; E-mail:
carlier@lebs.cnrs-gif.fr.
ABBREVIATIONS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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 K. Tamura, T. Hara, M. Yoshie, S. Irie, A. Sobel, and H. Kogo Enhanced Expression of Uterine Stathmin during the Process of Implantation and Decidualization in Rats Endocrinology, April 1, 2003; 144(4): 1464 - 1473. [Abstract] [Full Text] [PDF]
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