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J. Biol. Chem., Vol. 278, Issue 10, 8407-8419, March 7, 2003
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,From the Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, C/Velázquez, 144. 28006 Madrid, Spain
Received for publication, October 31, 2002, and in revised form, December 20, 2002
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
APPENDIX
REFERENCES
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The kinetics of Taxol association to and
dissociation from stabilized microtubules has been measured by
competition with the reference fluorescent derivative Flutax-1
(Díaz, J. F., Strobe, R., Engelborghs, Y., Souto, A. A., and Andreu, J. M. (2000) J. Biol. Chem. 275, 26265-26276). The association rate constant at 37 °C is
k+ = (3.6 ± 0.1) × 106
M Taxol,1 a complex
diterpene found in the bark of the Pacific yew (1), is a recent
addendum to the pharmacopeia of cancer treatment. Taxol is extensively
used in the therapy of ovarian cancer, metastatic breast cancer, head
and neck cancer, and lung cancer (2). At the time of the discovery of
its action on microtubules (3), Taxol had a unique characteristic
against other microtubule-binding drugs (4). Classical antimitotics
(colchicine and vinblastine) bind to tubulin and prevent the formation
of microtubules (5-8). Taxol activates tubulin by binding to the
microtubules, stabilizing the assembled form, and blocking the
microtubule dynamics necessary for cell function (9-11). Taxol is able
to drive the assembly of the otherwise inactive GDP-bound tubulin into
microtubules (12). Recently, promising new microtubule-stabilizing
anti-cancer drugs have been discovered, including cytotoxic products
from myxobacterias (epothilones (13)), sea sponges (discodermolide (14)
and laulimalide (15)), and a soft marine coral (eleutherobin (16)).
Despite the different chemical structures of these compounds, they
share a common pharmacophore with Taxol (17-20) and bind to the
Taxol-binding site with different
affinities,2 except
laulimalide (15), which binds at a different site in microtubules (21).
The conformation of microtubule-bound Taxol has thoroughly been
investigated (20, 22, 23).
The binding site of Taxol has been mapped in the 
1 s1. The reaction profile is
similar to that of the first step of Flutax-1 binding, which probably
corresponds to the binding of the Taxol moiety. The rate constant of
the initial binding of Flutax-1 is inversely proportional to the
viscosity of the solution, which is compatible with a
diffusion-controlled reaction. Microtubule-associated proteins bound to
the microtubule outer surface slow down the binding of Flutax-1 and
Flutax-2 10-fold. The binding site is fully accessible to Flutax-2 in
native cytoskeletons of PtK2 cells; the observed kinetic
rates of Flutax-2 microtubule staining and de-staining are similar to
the reaction rates with microtubule associated proteins-containing
microtubules. The kinetic data prove that taxoids bind directly from
the bulk solution to an exposed microtubule site. Several hypotheses
have been analyzed to potentially reconcile these data with the
location of a Taxol-binding site at the model microtubule lumen,
including dynamic opening of the microtubule wall and transport from an
initial Taxol-binding site at the microtubule pores.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
APPENDIX
REFERENCES
-tubulin subunit
using photolabeling (24-26). The labeled amino acid residues are in
agreement with the 3.5-Å resolution electron crystallographic structure of tubulin in Taxol-stabilized zinc-induced two-dimensional crystals (23, 27). Tubulin zinc sheets, whose assembly is not
GTP-dependent (28), consist of protofilaments similar to those that form the microtubules although in an antiparallel array. The
docking of these protofilaments into electron microscopy density maps
of Taxol-containing 14 and 15 protofilament microtubules (29, 30)
results in an atomic model of microtubules in which the binding site of
Taxol is located on the microtubule inner surface (30, 31). Such
luminal location will in principle make the binding site difficult to
access for Taxol site ligands in assembled microtubules. However, it
had been shown previously that Taxol modifies the flexibility of
microtubules in a few seconds (32) and that the reversible binding of
Taxol and its side chain analog docetaxel to an accessible site of
microtubules changes the number of their protofilaments within a time
range of 1 min (33). The binding site of Taxol is easily accessible for
two fluorescent derivatives of Taxol, Flutax-1 and Flutax-2 (34, 35).
These probes easily bind to and dissociate from native cytoplasmic and
spindle microtubules and centrosomes and are able to induce cell death
(34, 36). The binding of these ligands takes place in a two-step
mechanism (35) as shown in Equation 1,
The first step comprises the fast binding of the ligand with
micromolar affinity. This reaction does not affect the mobility of the
fluorescent group (either fluorescein or difluorofluorescein), and
because it is blocked by docetaxel it seems to be contributed to by the
binding of the Taxol moiety itself. The kinetic binding constants
k+1 are of the order of 106
M
(Eq. 1)
1 s1 at 37 °C (35)
indicating binding to an exposed binding site, in contrast with the
current microtubule model. Subsequent to the bimolecular step, there is
a monomolecular reaction that involves a rearrangement in the system
resulting in the immobilization of the fluorescent group. This step
probably implies a weak binding of the fluorescein moiety to the
microtubules (35).
The purpose of this study was to definitely establish the accessibility
of the binding site of Taxol in microtubules. Therefore, the kinetics
of Taxol association and dissociation have been measured in order to
discard any possible enhancement of the association rates by the
fluorescent side chain of the analogs. Thus, these fluorescent
derivatives of Taxol were used to learn about the mechanisms that allow
such accessibility by investigating the effects of the solution
variables and possible perturbants on the binding kinetics, such as the
presence or absence of MAPS and the C-terminal acidic segment of
tubulin. The binding rates of fluorescent taxoids to the Taxol site in
native microtubule cytoskeletons have been measured as well. Finally,
several hypotheses have been analyzed to potentially reconcile the
location of the binding site of Taxol in the lumen of model
microtubules with fast binding kinetics.
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EXPERIMENTAL PROCEDURES |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
APPENDIX
REFERENCES
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Tubulin and Taxoids--
Purified calf brain tubulin and
chemicals were as described (12). For glycerol-induced assembly,
tubulin was directly equilibrated in buffer: 10 mM
phosphate, 1 mM EGTA, 0.1 mM GTP, 3.4 M glycerol, pH 6.8. All tubulin samples were clarified by
centrifugation at 50,000 rpm, 4 °C, for 10 min using TL100.2 or
TL100.4 rotors in Beckman Optima TLX centrifuges. After centrifugation,
6 mM MgCl2 and up to 1 mM GTP were
added to the solution, giving a final pH 6.5. Microtubular protein,
containing tubulin and MAPs, was prepared as described (37) in 100 mM Mes, 1 mM EGTA, 1 mM
MgSO4, 2 mM 2-mercaptoethanol, 1 mM
GTP, pH 6.5. MAPs were prepared from microtubular protein as described
(38). hTau40, the longest isoform of microtubule-associated protein Tau
(39), was a gift from Dr. Vincent Peyrot and François Devred
(University of Marseille, France). It was dissolved in 10 mM phosphate, pH 6.5, buffer, and its concentration was
determined spectrophotometrically in 6 M guanidinium
chloride (employing a practical extinction coefficient of 6500 M1 cm1 at 280 nm, computed
using the ProtParam tool at www.expasy.ch). The activity of both MAPS
and Tau was confirmed by means of a cosedimentation assay with
cross-linked microtubules. Docetaxel (Taxotere®) was kindly provided
by Rhône-Poulenc Rorer (92165 Antony, France). Flutax-1
(7-O-[N-(4'-fluoresceincarbonyl)-L-alanyl]-Taxol) and Flutax-2
(7-O-[N-(4'-2,7-difluoro-fluoresceincarbonyl)-L-alanyl]-Taxol) were synthesized as described (40).
The diffusion coefficients of the taxoids were measured at 20 °C in 10 mM phosphate, pH 6.5, buffer using a synthetic boundary cell in an Optima XL-A (Beckman Instruments) analytical ultracentrifuge as described (41) at a speed of 15,000 rpm and wavelength of 230 (Taxol and docetaxel) or 495 nm (Flutax-1 and Flutax-2). The data were analyzed using the program VELGAMMA (42). The diffusion coefficients in GAB (glycerol assembly buffer, 3.4 M glycerol, 10 mM sodium phosphate, 1 mM EGTA, 6 mM MgCl2, 0.1 mM GTP, pH 6.5) at 37 °C were calculated from the values in phosphate buffer using the viscosity of a 30% solution of glycerol in water at 37 °C (1.55 centipoise (43)).
Preparation of Stabilized Microtubules-- Solutions of 50 µM tubulin in GAB were assembled at 37 °C for 30 min, and 20 mM glutaraldehyde was added to the solution, which was kept at 37 °C for another 10 min. The remains of the cross-linking agent were quenched by adding 60 mM NaBH4 (Fluka), and the solution was dialyzed overnight using Slide-A-Lyzer 10K dialysis cassettes (Pierce) against the desired buffer and drop-frozen in liquid nitrogen (35, 44). After this treatment 90% of the tubulin was found to have incorporated into the microtubules, and 100% of the assembled tubulin dimers were found to bind taxoids immediately after dialysis (as measured by a sedimentation assay (35)). Each batch of cross-linked microtubules was found to be stable against dilution and low temperatures. The binding sites of Taxol in drop-frozen microtubules were found to be stable for at least several months, while they slowly decayed at 4 °C with a half-life of ~50 days (average of four batches).
The morphology of the cross-linked microtubules was checked with electron microscopy as described previously (45). Cross-linked microtubules before freezing were normal and indistinguishable from non-stabilized microtubules, whereas unfrozen microtubules showed many openings along their structures.
When indicated, cross-linked microtubules in GAB 0.1 mM GTP were digested with 0.7% w/w subtilisin Carlsberg (Sigma) for 30 min at 37 °C. The reaction was stopped with 2 mM phenylmethylsulfonyl fluoride (Calbiochem), and the cleavage of the C-terminal fragment of both tubulin subunits was checked by SDS-PAGE of peptides (46).
Kinetics of Taxoids Binding to and Dissociation from Microtubules-- The kinetics of binding and dissociation of Flutax-1 and Flutax-2 were measured by the change of fluorescence intensity using an SS-51 stopped flow device (High-Tech Scientific, UK) equipped with a fluorescence detection system, using an excitation wavelength of 492 and a 530-nm cut-off filter in the emission pathway. The fitting of the kinetic curves was done with a non-linear least squares fitting program based on the Marquardt algorithm (47) where pseudo-first order conditions were used; otherwise the FITSIM package (48) was employed.
Because the binding of Taxol to microtubules does not produce any optical signal, its binding kinetics was measured by its effect on the observable Flutax-1 binding to microtubules. The binding of Flutax-1 to its microtubule site in the presence of different concentrations of Taxol was measured as described above. The kinetic curves were fitted using the rate constants of binding and dissociation of Flutax-1 that had been determined independently (see Ref. 35 and this work).
The kinetics of dissociation of Taxol from its site in the microtubules was measured by adding 10 µM Flutax-1 to a solution containing 1 µM Taxol and 1 µM binding sites. The dissociation of Taxol can be assessed by the small (less than 5%) change in fluorescence intensity of the solution because of the binding of Flutax-1 to the sites left empty by Taxol. The low signal to noise ratio required averaging a minimum of 10 experimental curves per measurement. These experiments were measured with a photon counting instrument Fluorolog 3-221 (Jobin Yvon-Spex, Longiumeau, France) with excitation wavelength 495 nm (0.1-nm bandpass in order to prevent photolysis) and emission wavelength 525 nm (5-nm bandpass).
The equilibrium binding constants of Flutax-1, Flutax-2, and Taxol to microtubules were obtained from anisotropy titration measurements made with a PolarStar microplate reader (BMG Labtechnologies, Offenburg, Germany) at different temperatures, as described (44).
Cytoskeletons and Fluorescence Microscopy-- PtK2 potoroo epithelial-like kidney cells were cultured as described previously (49). Unfixed coverslip-attached PtK2 cytoskeletons were obtained by washing cells eight times with PEMP (PEM buffer containing 4% polyethylene glycol 8000, pH 6.8), and then the cells were permeabilized with 0.5% (v/v) Triton X-100 in PEM (100 mM Pipes, 1 mM EGTA, 1 mM MgCl2, pH 6.8) for 90 s at room temperature and finally washed eight times in PEMP microtubule stabilizing buffer (34) to avoid disassembly. The cytoskeletons were dipped into 0.2 or 1 µM Flutax-2 for different times. They were rapidly washed 12 times with 2 ml of PEMP changing the washing well after six washes, mounted with 20 µl of PEMP, and their images were recorded with a Zeiss Axioplan epifluorescence microscope using a 100× Plan-Apochromat objective and a Hamamatsu 9742-95 cooled CCD camera (36). Controls were performed, in which the incubation with Flutax-2 was made in the presence of 50 µM docetaxel (which was preferred to Taxol due to its higher solubility). The displacement of Flutax-2 was observed in cytoskeletons incubated for 2 min with 1 µM Flutax-2, washed eight times with PEMP in two different wells, incubated with 13.3 µM docetaxel for different times, and then washed again eight times with PEMP in two different wells to remove the displaced Flutax-2, and the images were recorded as above.
The fluorescence intensities of a minimum of 10 different fields per time point were integrated using Scion Image (Scion Corp.) and averaged. In order to measure the fluorescence intensity per unit of length of microtubules, 3 × 10-pixel (0.08 × 0.24 µM) rectangles where defined over single interphasic microtubules in the images. The intensity of the rectangles was found to be homogeneous within each microtubule and among microtubules at each reaction time (the S.D. was 8 ± 2% of the mean value). The values of the controls performed with 50 µM docetaxel were subtracted from the data.
The maximal amount of tubulin contained in the cytoskeletons can be
roughly estimated from the number of cells attached to the coverslip
(~200,000) and the volume of each cell (~1012
liters). Assuming that the concentration of cytoskeletal tubulin inside
the cells is in the order of 50 µM (5 mg/ml), the maximal amount of tubulin per coverslip is 1011 mol, which is 200 and 40 times lower than the amount of Flutax-2 (0.2 or 1 µM) in the 2-ml well. The fact that the concentration of
Flutax-2 remains constant during the experiment (pseudo-first order
conditions) was confirmed by spectrophotometric measurements of
Flutax-2 before and after staining of the cytoskeletons, with no
change detected.
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RESULTS |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
APPENDIX
REFERENCES
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Kinetics of Taxol Binding to and Dissociation from Stabilized
Microtubules--
In order to discard the possibility that the
fluorescein moiety of the fluorescent taxoids could contribute to the
fast initial binding steps of these ligands to their site in the
microtubules, the kinetic constants of Taxol association and
dissociation were measured using a competition method. This method
precludes analysis of the exact mechanism of association of Taxol,
which is assumed to occur in a single step, thought to be equivalent to
the first step of binding of Flutax-1 (Introduction). The microtubules
used for these experiments had been stabilized by gentle cross-linking (see "Experimental Procedures"), a procedure that has been shown not to modify the kinetics of binding of fluorescent taxoids (35). Because the microtubules were frozen in liquid nitrogen and unfrozen prior to use, adequate controls were performed in order to check that
the kinetics of binding was not modified by freezing, as was the case.
The binding time course of 500 nM Flutax-1 to 500 nM sites was monitored in the presence of increasing
concentrations of Taxol, using stopped flow techniques. If the
association of Taxol were much slower than that of Flutax-1, no effect
should be observed when increasing the concentration of Taxol. On the other hand, if the association of Taxol is faster than or comparable with that of Flutax-1, both Taxol and Flutax-1 will bind
simultaneously, and only part of the sites will fill with Flutax-1; in
this way an appreciable effect in the amplitude of the observed
kinetics should be noticed. This is the case, as can be seen in Fig.
1A. A complete set of curves
at each temperature were simultaneously fitted (using the rate
constants of association and dissociation of Flutax-1 (35)), rendering
the kinetic constants of Taxol binding to its site in the microtubules
(Table I). The binding rate constant
value determined at 37 °C is five times higher than that of the fast
step of Flutax-1 association (Taxol, 3.6 × 106
M1 s1; Flutax-1, 6.1 × 105 to 7.4 × 105
M1 s1 (35)) and less dependent
on temperature, which indicates lower activation energy (Fig.
2A).
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Additionally, in order to correlate the kinetic constants with the
diffusion of the ligands, the diffusion coefficients of Taxol,
Flutax-1, and Flutax-2 were measured in 10 mM phosphate buffer pH 7.0. The values determined in this aqueous buffer at 20 °C, D20H2O, are as follows:
3.1 ± 0.3 × 1010 m2 s1 for Taxol, 2.8 ± 0.2 × 1010 m2 s1 for Flutax-1, and 2.7 ± 0.1 × 1010 m2 s1 for Flutax-2.
After correction for the viscosity of GAB and temperature, the
diffusion constants under the experimental conditions,
D37,GAB, were 2.2 ± 0.2 × 1010 m2 s1, 1.9 ± 0.2 × 1010 m2 s1, and 1.8 ± 0.2 × 1010 m2 s1,
respectively. The value of Flutax-2 is very close to the rough spherical approximation used in our previous work, 1.6 × 1010 m2 s1 (35). The diffusion
coefficient values of the fluorescent ligands indicate, following the
Stokes-Einstein equation, an effective radius of 8 Å.
The kinetic constants of dissociation of Taxol were determined by
displacing the Taxol bound to its site in the microtubules with a
10-fold excess of Flutax-1 (Fig. 1B). The values obtained for k1 of Taxol (0.091 s1 at
37 °C) (Table I) and its activation energy (Fig. 2A) are in between those of k1 and
k2 of Flutax-1, so it is not possible to
assign directly the dissociation step to either of the two dissociation
steps of the fluorescent taxoid.
The ratio of k+ and k
renders equilibrium binding constant values of Taxol binding in the
order of 107 M1 s1
which is compatible with equilibrium measurements of competition with Flutax-2 (Table II). There is
a factor from 3 to 4 between both sets of values, coming from a similar
factor observed between the equilibrium constants of the fluorescent
taxoids used as reference values calculated from their kinetic
parameters and those measured directly (Tables II and III in Ref.
35).
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Effect of Solution Variables in the Kinetics of Binding of Flutax-1 to Stabilized Microtubules-- Once the fast binding of Taxol to stabilized microtubules had been characterized, and shown to be clearly related to the first step of the association of Flutax-1, and the accessibility of the taxoid-binding site in the microtubules proved to be independent from the existence of the fluorescent moiety of Flutax-1 and Flutax-2, these fluorescent taxoids could be used as bona fide probes of the kinetics of binding and therefore were used for the rest of the study. Because Flutax-1 has a larger fluorescence intensity change upon binding, it was preferred for the stopped flow studies.
The first step of Flutax-1 and Flutax-2 binding to microtubules has been proposed to be a diffusion-collision reaction (35). To confirm this, the dependence of the association rate constant on viscosity was studied using GAB buffers with different concentrations of glycerol (0-60% instead of the usual 30%). If one of the reactants is as large as a microtubule and diffusion controls the reaction, the association rate constant will depend on the diffusion coefficient of the ligand (see Equation 3 in Ref. 35), which is inversely proportional to the viscosity of the medium, as shown by the Stokes-Einstein equation. As can be seen in Fig. 2B, k+1 linearly depends on the reciprocal of the viscosity of the media as expected from a diffusion-collision reaction.
The association of Flutax-1 to cross-linked microtubules was also studied under different conditions e.g. buffer composition, pH, ionic strength, and Mg2+ concentration, in order to learn from the system by observing the changes induced by these solution variables. The results (Table III) can be summarized as follows: (a) lowering the pH accelerates the binding reaction; (b) a moderate ionic strength doubles the reaction rate; and (c) Mg2+ seems to be a requirement for Flutax-1 binding.
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Effect of Modifying the Outer Surface of Microtubules; Kinetics of
Binding of Flutax-1 and Flutax-2 to MAP-containing
Microtubules--
To know if the surface-bound MAPs affect the binding
of taxoids to the microtubules, kinetic studies were performed using microtubules stabilized by MAPs (instead of cross-linking). Fig. 3A shows the time course of
binding of Flutax-1 to microtubules assembled from microtubular protein
in AB buffer (microtubular protein assembly buffer, 100 mM
Mes, 1 mM EGTA, 1 mM MgSO4, 2 mM 2-mercaptoethanol, 0.1 mM GTP, pH 6.5) under
pseudo-first order conditions. Two main differences from microtubules
assembled from pure tubulin in GAB buffer can easily be observed (Fig.
3B, dotted line). First, the reaction is slower, with a
half-life of 250 ms compared with 75 ms in similar reactant
concentrations. Second, the curve cannot be described by single
exponential decay but by the sum of two exponentials, indicating the
presence of at least two different processes. Because, in order to
assemble and stabilize microtubules by MAPs, AB buffer had to be
employed, adequate control
measurements3 were performed
that discarded the possibility that the observed effects could be due
to the buffer change.
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In order to confirm that the kinetic changes are truly due to the MAPs,
and not to the differences in the tubulin purification methods,
isolated MAPs were added in AB buffer to cross-linked microtubules,
which remained stable in the presence of at least 15% w/w of MAPs. The
same proportion of MAPs to microtubules was needed in order to turn the
monophasic kinetics into biphasic kinetics, and 20% w/w was necessary
to reconstitute the parameters of binding of Flutax-1 to microtubular
protein in AB (Fig. 3B; 0% MAPS (dotted line),
monophasic kinetics, k+ 7.56 ± 0.49 × 105 M1 s1; 20%
MAPS (solid line), biphasic kinetics,
k+1 2.18 ± 0.35 × 105
M1 s1,
k+2 2.43 s1, 35 °C; the
amplitude of the slow phase was still smaller than in the case of
binding to microtubules assembled from microtubular protein in AB
buffer, even if 40% w/w MAPS were added). These results prove that the
slowing down of the kinetics is a genuine effect of MAPs, and moreover
support previous evidence that the cross-linking of purified tubulin
microtubules does not modify their fluorescent taxoid binding
properties, (35).
To know whether the kinetic changes might be due to shielding of the
acidic C-terminal domains of tubulin by MAPS, the effect of the
positively charged hTau40 on the kinetics were measured. The binding of
recombinant hTau40 to microtubules negligibly modified the rate of
binding of Flutax-1. Different proportions (from 0 to 20% mol/mol
tubulin) of hTau40 were added to cross-linked microtubules in GAB, and
the kinetics of association of Flutax-1 were measured at 37 °C (the
k+1 values are as follows: 0%, 6.32 ± 1.12 × 105 M1
s1; 5%, 8.26 ± 1.22 × 105
M1 s1; 10%, 7.42 ± 1.54 × 105 M1
s1; 15%, 7.08 ± 1.01 × 105
M1 s1; 20%, 6.40 ± 0.97 × 105 M1
s1; the association curves were monophasic in all cases).
The effect of 20% mol/mol hTau 40 in the rate constants of Flutax-1
was further studied at different temperatures, but no significant
differences with the constants in the absence of hTau40 were found. The
combination of changing the buffer to AB and the addition of 20%
mol/mol of hTau40 did not reproduce the effect of MAPS
(k+1 37 °C, 23.58 ± 0.67 × 105 M1 s1,
monophasic kinetics). These results suggested that shielding of the
acidic C-terminal domains of tubulin by the positively charged Tau is
not enough to slow down the binding of Flutax-1, but another MAP
component is required, for example, high molecular weight MAPs
providing steric hindrance to the approaching ligand. In fact, reducing
the negative surface charge of the microtubules by subtilisin
proteolysis of the C-terminal segments of assembled microtubules (where
the main differences between tubulin isotypes and most of the
post-translational modifications are located (50, 51)) had no
significant effect on the association rate (k+1 37 °C non-digested microtubules, 6.34 ± 1.00 × 105 M1 s1;
C-terminal cleaved microtubules 6.01 ± 0.40 × 105 M1 s1),
suggesting as well no influence of the tubulin isotypes or post-translational modifications in the kinetics.
The kinetics of binding of Flutax-1 and Flutax-2 to microtubules with
MAPS were quantitatively studied in AB buffer. Fig. 4 shows the dependence of the observed
rate constants of binding of Flutax-1 at 37 °C on the concentration
of sites. The fast constant depends linearly on the concentration of
sites, although the slow phase saturates with the concentration of
sites. This indicates a mechanism where binding is followed by a
monomolecular reaction, similar to the one observed in the binding to
microtubules assembled from pure tubulin in GAB (see Equation 1).
Nevertheless, here the rate constant of the second step, the monophasic
reaction, is still quite strongly dependent on the concentration of
sites, which indicates that the equilibrium constant of the first step is lower than in the absence of MAPs, and so the reaction is not completely displaced toward the end product. The second step of the
reaction is now observed, as it displaces the equilibrium toward the
final state. Within such a model, if the difference between both
constants is large enough (52), it is possible to determine the values
of the individual kinetic constants from the values of the observed
rate constants, as shown in Equations 2 and 3,
![kf<SUB><UP>obs</UP></SUB><UP> = </UP>k<SUB><UP>+1</UP></SUB><UP>·</UP>[<UP>sites</UP>]<UP> + </UP>k<SUB><UP>−1</UP></SUB>](/content/vol278/issue10/fulltext/8407/img001.gif)
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(Eq. 2) |
![ks<SUB><UP>obs</UP></SUB><UP> = </UP><FR><NU>K<SUB><UP>1</UP></SUB><UP>·</UP>k<SUB><UP>+2</UP></SUB><UP>·</UP>[<UP>sites</UP>]</NU><DE><UP>1 + K<SUB>1</SUB>*</UP>[<UP>sites</UP>]</DE></FR><UP> + </UP>k<SUB><UP>−2</UP></SUB>](/content/vol278/issue10/fulltext/8407/img002.gif)
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(Eq. 3) |
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From the data in Fig. 4 it is possible to determine the values of
k+1 and k+2. The value of
k1 can be determined (although with a large
error margin) because its value is significant enough with respect to
the product k+1·[sites]. Because
k2 is very small, the extrapolation to
[sites] equal to 0 is too large to determine its value properly,
which has to be determined by dissociation kinetics. The values of
k2 are around 2 orders of magnitude lower than
those of k1; k2 is the rate-limiting step of the dissociation reaction (as for the dissociation from microtubules assembled from pure tubulin in GAB). The
rate constants of dissociation of Flutax-1 and Flutax-2 from their site
in MAP-containing microtubules were measured in the same way as for the
complex with microtubules assembled from pure tubulin (Table III in
Ref. 35). The reaction kinetics was found to be monophasic (see Fig.
5), and the values obtained for the
koff rate were very similar to those obtained in
absence of MAPS in GAB. The rate constants determined for Flutax-1 and
Flutax-2 are summarized in Table IV. The
comparison of the kinetic constants to those obtained for the binding
of Flutax-1 and Flutax-2 to microtubules without MAPS (see Table II in
Ref. 35) shows that only the rate of the first step
(k+1) is significantly reduced by a factor of
3-4 in the presence of MAPS, whereas the values of the other kinetic
constants (k1, k+2, and
k2) are only marginally altered. The
thermodynamic parameters of the binding of Flutax-1 and Flutax-2 to
MAP-containing microtubules (Table V) do
not show great differences from those obtained with purified tubulin
microtubules, except for the reduction of the activation energy of the
first step of the binding, the one affected. Curiously, the presence of
MAPs reduces the activation energy of the binding of Flutax to values
similar to those of Taxol itself. The parameters of the second step of
the binding are very similar to those observed for the binding of
Flutax-2 to the microtubules in GAB, which suggests that the
immobilization of the fluorescein group is also very similar, and it is
not affected by the presence of MAPS.
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Kinetics of Binding of Fluorescent Taxoids to Microtubule
Cytoskeletons--
It might be argued that the kinetics of taxoid
binding to in vitro assembled microtubules is not
representative of cellular microtubules. In order to approach a more
physiological situation and to know whether the accessibility of the
binding site in native microtubules is similar to those assembled
in vitro, the kinetics of the binding of fluorescent taxoid
(Flutax-2 was chosen in this case due to its higher photostability) to
native cytoskeletons from PtK2 cells at 25 °C has been
semiquantitatively characterized by approaching pseudo-first order
conditions, employing CCD-detected epifluorescence microscopy images
(Fig. 6, A-D). The plot of
the averaged fluorescence intensity versus incubation time
(Fig. 6, E and F; 0.2 and 1 µM
Flutax-2, respectively) shows an exponential staining of the
cytoskeletons (the data do not permit us to distinguish between a
single or double-exponential best fit). At 25 °C the half-lives of
the binding process and 0.2 and 1 µM Flutax-2 are 80 and
12 s, respectively, which correspond to a value of
k+ of 6.1 × 104
M1 s1. This value is compatible
with the extrapolation to 25 °C of k+1 of
Flutax-2 binding to in vitro assembled microtubules with
MAPS which renders a rate constant of 1.3 × 105
M1 s1. Docetaxel displaces
Flutax-2 from its site in the microtubules with
kobs 0.75 × 102
s1 (half-life in the range of 90 s (Fig.
7)). The dissociation rate constant of
Flutax-2 from its site in microtubules assembled in vitro
(Table IV) is k2 1.04 × 10(2
s1 (half-life of 66 s) which is close to the rate of
the cytoskeleton destaining process.
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In order to check if the staining and destaining was spatially
homogeneous within a single microtubule (i.e. the complete microtubule stains and destains at the same time), the fluorescence intensity per unit of length of individual microtubules was measured. No differences were observed among different fragments of microtubules within the same image. It can be concluded from the data available that
the kinetics of Flutax-2 binding to native microtubules from PtK2
cultured cells was not significantly different from the kinetics of
binding to in vitro assembled MAP-containing microtubules.
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
APPENDIX
REFERENCES
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Kinetics and Thermodynamics of Binding of Taxol and Fluorescent
Taxoids to Microtubules--
Because the exact kinetic mechanism of
Taxol binding to microtubules is unknown, it had to be assumed that it
consists of a single-step reaction in order to calculate the
thermodynamic parameters (Table II), which have to be considered with
this caution. Nevertheless, the linearity of the Arrhenius plots (Fig.
2A) and the compatibility of the kinetic data with the
equilibrium measurements suggest that a single reaction (or a first
step of a two-step reaction) is in fact being observed. The apparent
reaction profile of Taxol binding is similar to the first step of
Flutax-1 and Flutax-2 binding (not shown; see Ref. 35). This strongly
supports the idea that this step mainly corresponds to the binding of
the Taxol moiety of the fluorescent derivatives. It has to be pointed out that the activation energy of the binding of Taxol is lower, indicating a easier entrance of the smaller ligand to its site. The
free energy of the binding of Taxol at 37 °C is around 
45 kJ
mol1, compatible with a predicted value
G 
48 kJ mol1 (53). The binding
reaction of Taxol is endothermic, which explains why Taxol induces
microtubule assembly at low temperatures, because the binding
compensates for the decrease in the free energy of assembly (the
extrapolated binding affinity of Taxol at 4 °C is 7.6 × 108 M1, so it may contribute to
the overall assembly-binding linked process with 47 kJ
mol1).
In a diffusion-controlled reaction, the binding rate of a small ligand to an accessible site in a relatively immobilized molecule depends on the diffusion coefficient of the ligand and also on the efficiency of the collisions. The diffusion constant of Taxol is 20% larger than those of Flutax-1 and Flutax-2. Because Flutax-1, Flutax-2, and Taxol compete for the same site and the surface areas of Flutax-1 and Flutax-2 are 40% larger than that of Taxol, assuming that the effective surface for binding is the same, the efficiency of collision is expected to be 1.4 times larger for Taxol. The combination of the larger diffusion constant with better collision efficiency renders an expected kinetic rate of association 1.7 times larger for Taxol than for Flutax-1 and Flutax-2 (if the reaction is diffusion-colliding controlled and the observed step is the binding of the Taxol moiety of the molecule). The observed ratio is larger, which implies lower efficiency than expected for Flutax. There are two possible reasons for this, the first one is that Taxol and Flutax might have to pass through a pore of a restrictive size. Because their sizes are different, the flux of Taxol through the pores would be larger. However, this does not explain the dependence on temperature observed (k+1 (Taxol)/k+1 (Flutax-2): 25 °C 11.5, 30 °C 7.3, 35 °C 5.2, 37 °C 2.6, 40 °C 2.7), unless the pore size is temperature-dependent. In addition, diffusional flux through a pore of a size similar to those of the ligands in the microtubule wall (31) will not allow the binding reaction at the observed kinetic rate (35). A second, more feasible explanation is that the fluorescein moiety is hindering the binding of the Taxol moiety, perhaps by collapsing over it, as suggested by the small difference in the Stokes radii of Flutax and Taxol (8 and 7 Å, respectively). The activation energy of the first step of the binding of Flutax is larger than that of Taxol. This increase may indicate the existence of active and inactive conformations of Flutax. Such conformational equilibrium, if fast enough, will not alter the observed kinetic mechanism but only reduce the observed rate constant. If this equilibrium is influenced by the temperature, this will also explain the differences in the measured activation energy of the observed reaction.
The Location and Access of the Taxol-binding Site of Microtubules;
Contradiction between Structural Models and Kinetic Results--
Early
low resolution microtubule models (45, 54) placed the Taxol-binding
site at the interprotofilament space of microtubules. A compatible
location was observed in a projection difference map of zinc-induced
tubulin sheets at 6.5 Å resolution (55). Such location of the binding
site would be compatible with the presently established fact that
taxoids are able to access their binding site very rapidly. But since
the three-dimensional high resolution model of microtubules appeared
(31), one apparent contradiction showed up. The binding site of Taxol,
one of the most effective drugs in favoring microtubule assembly, was
mapped into the lumen of the tube, hidden from the outer solvent. It was suggested that rapid luminal access could occur via fenestrations in the microtubule wall (31). The three-dimensional structure of
tubulin dimer, the location of the Taxol-binding site in the tertiary
structure of -tubulin, and the alignment of dimers to form
protofilaments (23, 25, 27, 56-58) are all together hardly
questionable. However, the association rate constants of the
fluorescent derivatives of Taxol, Flutax-1, and Flutax-2 to the
Taxol-binding site of microtubules, in the order of 106
M1 s1, have been shown to be
quantitatively incompatible with these ligands entering through the
open microtubule ends or through fenestrations in the microtubule wall
of 25 Å diameter (33-35). A simple way to place the Taxol-binding
site of -tubulin near the microtubule surface, i.e. a
large rotation of the protofilament that would be needed in order to
expose Taxol to the outer solvent (see Fig. 10 in Ref. 35), has been
shown to be incompatible with several fits of the same tubulin
structure into microtubules performed with different microtubule
density maps and fitting strategies (30, 31, 59). The current
microtubule models indirectly account for the well known fact that
Taxol has negligible affinity for isolated 
-tubulin dimers but
high affinity for microtubules or zinc-induced sheets (51, 60-62),
through the contacts made by the M-loop (31, 63-64).
The results of the present work prove, we believe beyond any reasonable doubt, that Taxol binds to microtubules at a site which is directly accessible from the bulk of the solution. (i) The association rate of Taxol to microtubules is slightly faster than those of the rapidly binding fluorescent derivatives. (ii) The association of the fluorescent taxoids has been shown to be diffusion-controlled and significantly slowed down by MAPs, known to bind at the microtubule surface and to reduce the microtubule dynamics. (iii) The study has been extended to native microtubules prepared from cultured cells, which bind fluorescent taxoid at a similarly high rate, indicating an equally accessible Taxol-binding site, although it should be noted that microtubule cytoskeletons from cultured cells and microtubular protein have typically different tubulin isotypes, post-translational modifications, and MAPS (51). In addition, although the electron micrographs of microtubules assembled in vitro from pure tubulin typically show many areas in which the tube is open, which in principle provides a better access of the ligand through these areas, the labeling of the Ptk2 cell microtubules by the fluorescent taxoid is spatially homogeneous, indicating that all parts of the microtubules are equally accessible.
Very recently it has been argued that the fenestrations in the
microtubule wall allow the passage of Taxol for binding at the
microtubule lumen (65). However, the relevant issue here is not whether
the fenestrations may eventually allow the ligand to squeeze in, but
whether the ligand diffusion through them can be rapid enough to
account for the fast binding observed. We previously discarded ligand
passage through the holes (35) because at that time the size of the
observed fenestrations in microtubule walls was roughly 10 Å (66),
smaller than the 15 Å shortest dimensions of Flutax and Taxol. Let us
now reexamine this possibility in view of the refined structure of
tubulin dimers (23), the 14-Å microtubule map (30), the 8-Å
microtubule map (65), and an independent model of microtubule structure
(59). Fig. 8 shows a comparison of the
dimensions of the taxoids used in this work with the fenestrations in a
model microtubule wall. The size of the pores is comparable with the
shortest dimension of the ligands; therefore, these pores should
strongly restrict ligand diffusion through them.
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To make an estimation of the order of magnitude of the maximum kinetic rate of binding of Taxol to a luminal microtubule site, it is convenient to divide the process into three steps with different factors whose product gives the rate constant: (a) the diffusion-limited ligand-microtubule encounter, (b) the efficiency of passage through the pores, and (c) the efficiency of productive collisions with the luminal binding site.
(a) For diffusion-limited ligand collision with
microtubules, a maximum estimation for the rate constant value for a
bimolecular reaction between Flutax and tubulin assembled in
microtubules is 6.4 × 107
M1 s1 under our experimental
conditions (see Equation 3 in Ref. 35). Note that due to the
association of the tubulin subunits into an aggregate, this is smaller
than the ~4 × 109 M1
s1 value that can be estimated as a hypothetical
diffusion limit for the binding of Taxol to unassembled tubulin (67).
The frequency of collision of a small molecule with a large aggregate
depends on the diffusion coefficient of the small molecule and the
radius of collision, which can be approximated by the effective radius of the aggregate. The decrease of the rate of collision with the number
of subunits in several polymers can be calculated by means of classical
approaches (67, 68). In order to obtain the collision rate of the
ligand with the subunit in the aggregate, the collision rate with the
aggregate has to be divided by the number of subunits. The effective
radius of the aggregate divided by the number of subunits is not
constant but markedly decreases with the size of the aggregate, at a
rate that depends on its geometry. Therefore, polymerization results in
a decrease of the diffusion limit of the bimolecular rate constant of
ligand binding per subunit (see Fig. 10 and "Appendix").
(b) Regarding the efficiency of ligand passing through
microtubule pores, in order to match the experimental association rate constant of 1.4 × 106 M1
s1 (35) with the 6.4 × 107
M1s1 diffusion limit, 2.2% of
effective collisions between Flutax and microtubules is required.
Similarly, 5.1% of effective collisions of Taxol with microtubules is
needed to match the measured rate constant of 3.6 × 106 M1 s1 (see
"Results"). A collision efficiency of 2-5% would be reasonable for taxoid binding at the microtubule surface, accounting for the
percentage of surface area of microtubules forming the binding interface (35). The minimum pore sizes that may provide efficiencies in
this range for ligand passing through them can be estimated from simple
geometrical calculations of effective surface and angle of passage
(35). For example, a 32 (diameter) × 30-Å (deep) cylindrical
pore (which is of a size nearly comparable with a tubulin monomer)
would allow 2.2% of successful collisions of a Flutax-equivalent 16-Å
diameter sphere, resulting in ligand diffusion into the microtubule
lumen. A 30-Å deep truncated and cone-shaped pore with 20-Å inner
diameter and 32-Å outer diameter would lead to 1.9% of successful
collisions. A more realistic (65) pore of truncated cone shape, say 30 Å (deep) × 17-Å inner diameter and 20-Å outer diameter, would
have an efficiency of only 0.08% successful collisions, yielding a
diffusion limit to the association rate constant of 5 × 104 M1 s1, which is
2 orders of magnitude smaller than the observed value.
(c) Due to the efficiency of collision with the binding site, accounting in each of the above cases for the orientation and kinetic energy factors of the collision of the internalized ligand with the luminal binding site will further reduce the effective rate constant by order(s) of magnitude (69, 70). Therefore, in order to allow the ligand flux required to account for the observed binding reaction, the openings in the microtubule wall would have to be totally non-restrictive, i.e. much larger than the ligands. Otherwise the low efficiency of ligand passing through the pores reduces the kinetic rate of binding by several orders of magnitude below the experimentally observed value. Such non-restrictive openings are certainly larger than the pores in the model of Chacón and Wriggers (17 Å (59); Fig. 8) or in the electron microscopy maps of Meurer-Grob et al. (30) (about 15 by 25 Å) and Li et al. (65) (~17 Å). This analysis shows that it is not possible for Taxol to passively diffuse through the presently known microtubule fenestrations rapidly enough to bind at a luminal site at the observed rate.
In summary, the puzzle remains of how taxoids bind so rapidly to an apparently inaccessible model binding site at the microtubule lumen. We offer and analyze three hypotheses to potentially reconcile the fast kinetics of Taxol binding with a Taxol-binding site at the microtubule lumen. (i) The Taxol-binding site is exposed in unliganded microtubules, but it is buried following Taxol binding. (ii) The microtubule inside dynamically opens to the solvent so that the lumen is not a separate solution compartment. (iii) There is an initial Taxol-binding site (or channeling) at the microtubule surface that facilitates its transport into the lumen.
Does the Taxol-binding Site Become Inaccessible After Binding?-- It might be conceivable that the Taxol-binding site is exposed in unliganded microtubules but buried in the Taxol bound ones. When Taxol binds rapidly to microtubules, they are known to undergo a slow conformational change entailing the loss of one protofilament on average (33), which may as well internalize the binding site. This type of binding mechanism is formally equivalent to Equation 1 (Introduction). Although this hypothesis may appear as an easy explanation, there are several lines of experimental evidence against it. First, the wall structure of Taxol-induced microtubules is similar to that of drug-free microtubules at 30-Å resolution (54), except for the change in the number of protofilaments. Second, the binding of docetaxel, a side chain analog of Taxol, does not induce this diameter change in microtubules, indicating that the Taxol-induced change is restricted to the rearrangement of the subunits to compensate for the loss of one protofilament (33, 45). Third, both 14-Å resolution electron cryomicroscopy maps of microtubules stabilized by GMPCPP and by docetaxel have similar fenestrations (30).
Does Taxol Bind to an Internal Site via Large Dynamic Openings in the Microtubule Wall?-- The hypothesis that Taxol accedes to its binding site via openings or defects in the microtubule wall was considered in our early publication (Fig. 7D in Ref. 33) and by Nogales et al. (31) to justify the fast binding. Two causes that might in principle contribute to the formation of such large pores, in the size range of tubulin monomers, are as follows.
(a) In the refined structure of tubulin at 3.5-Å resolution
(23), several features involved in protofilament interactions in the
microtubule (the M-loop and the loop between H1 and S2) have been
retraced, and a Zn2+ ion has been found interacting with
the M-loop of the -subunit of tubulin. This ion should distort the
natural lateral interaction between protofilaments, so that tubulin
assembles into zinc sheets instead of microtubules. Because the
microtubule models have been constructed using the tubulin dimer
structure obtained from these zinc-induced sheets, the real lateral
interactions between the tubulin dimers in microtubules, which have no
zinc bound, should be different, and fenestrations larger than in the
models might exist. The electron microscopy structure of
Taxol-stabilized microtubules at 8 Å resolution (65) indicates a large
reorganization of the Taxol-binding site with respect to the electron
crystallographic structure of tubulin zinc sheets, including helix H6
and the M loop. There is a density in the averaged - and -tubulin
monomer that corresponds to part of the regions occupied by Taxol in
the -tubulin structure and the B9-B10 peptide loop in -tubulin, which is apparently backed by holes of comparable size (see Figs. 6B and 9A in Ref. 65). However, the fenestration
size is still restrictive, not allowing ligand passage at the required
kinetic rate (see above).
(b) The interprotofilament contacts in the microtubules may be so weak that large openings might be continuously appearing and closing in the microtubule wall. Lattice defects such as longitudinal shifts between protofilaments and changes in the number of protofilaments within single microtubules have been observed by cryo-electron microscopy, suggesting the weakness of the interprotofilament contacts (71-73). If such openings were not periodical, they would not be easily observed in electron micrographs of microtubules without Taxol bound.
This explanation would be supported by the observation that microtubules are able to rapidly change the number of their protofilaments in response to taxoid binding (33). Even if cross-linking of microtubules would stabilize these interprotofilament contacts, microtubules would probably be frozen by fixation in a given situation, which would not affect the kinetics of the reaction, because the openings would be frozen. On the other hand, interprotofilament contacts would be expected to be stabilized by the presence of MAPs, which slow down the binding reaction and are known to block the ligand-induced changes in the number of protofilaments (54). Because these contacts would be stabilized as well by the taxoids, the taxoid off-rate could be similar in both types of microtubules, as observed.
In this way it might be possible that large holes open and close in the
microtubule wall, resulting in a non-restrictive pore, so that the
lumen ceases to be a separate chemical compartment. From a kinetic
point of view, this is equivalent to adding a monomolecular pre-equilibrium step before the bimolecular binding event in the scheme
of Equation 1 (Introduction), consisting of the conversion of inactive
(closed) into active (open) binding sites. For this equilibrium to have
passed undetected in the kinetic analysis of taxoid binding, it would
have to be either extremely slow or have a rate constant
k0 more than 10-fold larger than the next step,
that is k0 
10·k1·[ligand], which from typical values
(35) is in the order of k0 100 s1. The hypothesis is not supported by the fact that the
presence of a larger percentage of open areas in cross-linked frozen
and melted microtubules results in the same kinetic rate of binding as
in non-frozen ones. In addition, differences in the tubulin isoforms
and post-translational modifications, between tubulin from brain and
from cultured cells (51), which are known to affect the stability and
dynamics of the microtubules (74, 75), are not reflected in the
kinetics of binding of Flutax-2. This hypothesis of dynamic openings of
non-restrictive size for ligands may be tested measuring the
accessibility of the Taxol-binding site to macromolecular probes, whose
size would hinder entering the microtubule lumen even through such holes.
These openings would not correspond to the so-called "seam" of the microtubules, because microtubules with the 3-start helix, B-lattice, and 13 protofilaments, which are the majority in the conditions of the study, do not have a seam (52, 73) and would be expected to bind taxoids at a slower rate. Control experiments in which the cross-linked microtubules stained with Flutax-2 were observed both by fluorescence and phase contrast showed that all microtubules were immediately stained after Flutax-2 addition. Even more, because transitions in the number of protofilaments
