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J. Biol. Chem., Vol. 280, Issue 44, 37048-37060, November 4, 2005

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Alternating Site ATPase Pathway of Rat Conventional Kinesin^*

From the Institute for Cellular and Molecular Biology, University of Texas, Austin, Texas 78712

Received for publication, March 17, 2005 , and in revised form, August 12, 2005.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The pathway of ATP hydrolysis by rat kinesin was established by pre-steady-state kinetic methods. A 406-residue long N-terminal fragment was shown by sedimentation equilibrium analysis to form a dimer with a K_d of 46 nM. The pathway of ATP hydrolysis follows the Gilbert-Johnson pathway determined previously for a similarsized N-terminal fragment of Drosophila conventional kinesin. However, the rates of ADP release were at least 3-fold faster, and ATP hydrolysis was 5-fold faster. Paralleling our previous mechanistic data, these results support an alternating site ATPase pathway, including a captive head state as an intermediate in the kinesin ATPase cycle. The kinetic data presented in this report once again point to the importance of the captive head state and argue against a pathway that short-circuits this key intermediate. In addition, several unique aspects of the rat kinesin kinetics reveal new aspects of the ATPase-coupling mechanism. These studies provide a baseline set of kinetic parameters against which future studies of rat kinesin mutants may be evaluated and directly correlated with the structure of the dimeric kinesin.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Conventional kinesin is a plus-end-directed microtubule motor driving fast axonal transport of vesicles. Kinesin superfamily genes are represented in every eukaryotic genome examined, ranging in number from 6 genes in Saccharomyces cerevisiae to 45 in the human genome (1, 2). Members of this family are implicated in the transport of vesicles and organelles (3), as well as a variety of other microtubule-dependent processes, such as meiotic and mitotic chromosome movement (4), the maintenance and function of cilia and flagella (5), and the regulation of microtubule dynamics, as in the case of mitotic centromere-associated kinesins (6).

Kinesin is a dimer of mutually entwined -helices forming a coiled-coil structure, flanked on both ends with N- and C-terminal globular domains (7). The N terminus forms a globular motor domain of conserved sequence and structure, responsible for microtubule binding and ATP hydrolysis. Structures of the motor domain of rat conventional kinesin in both dimeric and monomeric forms containing ADP have been solved (8, 9), but subtle structural differences between the two heads suggest different conformational states that may be related to motility.

Conventional kinesin is a processive motor; for example, Drosophila kinesin dimer hydrolyzes 100 molecules of ATP before dissociating from the microtubule (10-12). Processivity is maintained by an alternating site ATPase pathway, in which one head of kinesin remains tightly associated with a microtubule while the other rapidly diffuses to the next binding site (13-15). Kinesins appear to achieve motility via a hand over hand mechanism, in which the two microtubule-binding heads of the protein alternate in relative position, advancing the molecule by 8 nm, the distance spanned by one -tubulin dimer, with each step (16-19). This model is supported by structural studies of fixed kinesin·microtubule complexes, x-ray crystallography of monomeric and dimeric kinesin, spectroscopic analysis of kinesin dynamics using fluorescent and spin-label probes, physical measurements of kinesin force-generation using laser-trapping, and kinetic analysis of kinesin ATPase activity (14, 15, 20-23).

Despite extensive analysis by a variety of techniques, there remains some uncertainty concerning the enzymatic pathway by which processive motility is achieved, and the nature and order of the conformational changes. Taking advantage of rapid, transient-state kinetic analysis of Drosophila conventional kinesin a detailed model of kinesin ATPase emerged, based upon measurements of the rate and order of each step in the pathway (10, 11, 14, 15, 24-26). Similar studies using human kinesin (27-30) have produced a similar ATPase cycle with kinetic parameters that differ somewhat, especially with the extent to which ADP release may be partially rate-limiting. Data on each of the kinesin·microtubule ATPases provide evidence for the central role of a "captive head" state in the ATPase cycle where a nucleotide-free head is attached strongly to the microtubule while the other head retains ADP and interacts only weakly with the microtubule. Evidence for the alternating site ATPase pathway relies upon the observed effect of ATP binding to the open site stimulating the release of ADP from the second site. It can be argued that all of the data in support of an alternating site ATPase stem from experiments that are dependent upon the captive head state. In contrast, based upon purely structural and equilibrium data, Rice et al. (31) proposed a truncated model in which the captive head state is bypassed during processive movement.

One drawback of our kinetic studies using Drosophila kinesin is the absence of an x-ray crystal structure for this molecule. Given the potential utility of a crystal structure for structure-function relationship analysis, in which interactions between residues appearing in the crystal structure might be targeted for mutagenesis, it is unfortunate that one of the best-understood and most thoroughly characterized conventional kinesins has proved refractory to crystallization. Because Rattus norvegicus kinesin is the only source that has yielded the structure of the dimeric form of kinesin, we have turned our attention to rat kinesin for more detailed structure/function studies. In the present study, we examined the transient state kinetics of the ATPase hydrolysis cycle of a 406-residue N-terminal fragment of rat conventional kinesin. The results of this study establish the ATPase pathway for another kinesin whose structure is known and provide a standard against which the kinetics of rat kinesin mutants can be compared in the accompanying paper (32).

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Materials—Escherichia coli BLR(DE3) was obtained from Novagen Inc. (Madison, WI). E. coli XL1-Blue was from Stratagene (La Jolla, CA). QIAprep Plasmid DNA extraction kits were purchased from Qiagen Inc. (Valencia, CA). QuikChange site-directed mutagenesis kits were purchased from Stratagene. Bio-Rex 70 resin (75- to 150-µm wet bead size) and Bio-Gel P-4 Gel (45- to 90-µm particle size) were from Bio-Rad Laboratories (Hercules, CA). DEAE-, Q-, and S-Sepharose Fast Flow chromatography resins were from Amersham Biosciences. Taxol (paclitaxel) was purchased from Sigma Co. Radiolabeled ATP ([-³²P]ATP, >3000 Ci/mmol) was from PerkinElmer Life Sciences, and N-methylisatoic anhydride and N-[2-(1-maleimidyl)ethyl]-7-diethylaminocoumarin-3-carboxamide (MDCC)² were from Molecular Probes (Eugene, OR). Polyethyleneimine-cellulose F TLC and silica gel 60 F₂₅₄ plates (EM Science) were purchased from VWR Scientific (West Chester, PA). Other chemicals were from Sigma or Fisher Scientific.

Media and Buffers—The following media and buffers were used for the experiments described: Buffer A (30 mM HEPES, pH 7.2, 4 mM MgCl₂, 0.1 mM EDTA, 20 µM ATP); Buffer B (50 mM Tris-HCl, pH 8.2, 4 mM MgCl₂, 0.1 mM EDTA, 20 µM ATP); ATPase Buffer (40 mM HEPES, pH 7.2 with KOH, 5 mM magnesium acetate, 50 mM potassium acetate, 0.1 mM EDTA, 0.1 mM EGTA, 0.5 mM dithiothreitol); and PM buffer (100 mM Na-PIPES, pH 6.6, 4 mM magnesium acetate, 1 mM EGTA). The pH of each buffer was adjusted at the temperature at which it was to be used.

Data Analysis—Thin-layer chromatography of radiolabeled nucleotide measured formation of ATPase hydrolysis product, which was quantified using a Storm 860 PhosphorImager and ImageQuaNT software (Amersham Biosciences). Linear and nonlinear regressions to kinetic data were performed using Excel (Microsoft Corp., Redmond, WA) and GraFit (Erithacus Software, Horley, Surrey, UK). Graphs were prepared using GraFit. Simulations of kinetic data were performed using the kinetic simulation software KinTekSim (33, 34) from KinTek Corp. (Austin, TX, www.kintek-corp.com).

Construction of pKHC407A—The plasmid used in the expression and purification of an N-terminal fragment of rat conventional kinesin heavy chain was derived from pKHC406 (35), a gift from Scott Brady (University of Texas Southwestern Medical Center, Dallas TX). The C-terminal polyhistidine tag encoded in the plasmid was removed by substitution of the first non-kinesin codon in the open reading frame of the plasmid with a nonsense codon, using site-directed mutagenesis (36). Kinesin mutants were likewise generated using PCR-mediated codon substitution. Oligonucleotides for site-directed mutagenesis were purchased from Integrated DNA Technologies (Coralville, IA). For removal of the polyhistidine tag, the complementary primers 5'-CACCTGTGGTTGACTAGCTTGCGGCCGCAC-3' and 5'-GTGCGGCCGCAAGCTAGTCAACCACAGGTG-3 were used. The resulting plasmid, pKHC407A, expresses an N-terminal fragment of rat conventional kinesin terminating at asp407. A QuikChange site-directed mutagenesis kit (Stratagene), was used with a GeneAmp PCR System 2400 (PerkinElmer Life Sciences) for polymerase chain reaction (37). Temperature cycles were as recommended by the kit manufacturer: 95 °C (30 s), 55 °C (1 min), and 68 °C (6 min) for 16 cycles.

Expression of Kinesin—The pKHC406 plasmid and its derivatives are based on the isopropyl 1-thio--D-galactopyranoside-inducible pET expression vector (38-40). Colonies of transformed E. coli BLR(DE3) were grown overnight with shaking at 37 °C in 2 liters of LB broth plus 1% dextrose, 50 µg/ml kanamycin, and 10 µg/ml tetracycline. Induced cells were centrifuged and pellets were frozen at -80 °C.

Purification of Kinesin—Kinesin preparations were kept on ice or at 4 °C. Cells from 4-liter-induced culture were resuspended in 50 ml of Buffer A plus 95 mM NaCl, 1 mM EDTA, 1 mM EGTA, 0.5 mM phenylmethylsulfonyl fluoride, 0.5 µg/ml leupeptin, 0.25 µg/ml lysozyme. The suspension was disrupted using a Branson Sonifier Model 450 for 4 bursts on ice, power setting 4, 20 s/burst. The lysate was then cleared by centrifuging (Beckman JA-25.50 rotor, 23,000 x g, 30 min), and loaded onto a 34-ml BioRex70 column (2 cm² x 17 cm) pre-equilibrated with Buffer A plus 95 mM NaCl. The column was washed with 3 column volumes of the equilibrium buffer. Elution of protein was performed using a 4-column volume gradient spanning 95-485 mM NaCl, and 4-ml fractions were collected. The fractions were evaluated by SDS-PAGE, and kinesin appeared as an abundant 45.5-kDa protein. Fractions were pooled and dialyzed for 2 h against 1 liter of Buffer B plus 95 mM NaCl. The dialyzed kinesin was loaded onto a 10-ml Q-Sepharose column (2 cm² x 5 cm) pre-equilibrated with the same buffer. After a 3-column volume wash, kinesin was eluted with a 6-column volume gradient from 95 to 475 mM NaCl, and fractions were collected. Those fractions in which a 45.5-kDa protein was predominant were selected and pooled. Pooled fractions were dialyzed twice for 2 h apiece against 1 liter of Buffer A plus 40 mM NaCl. After dialysis, the preparation was loaded onto a 3-ml Q-Sepharose column (0.8 cm²x 3.75 cm), pre-equilibrated with the same buffer. Elution was done with a 3-column volume gradient from 40 to 240 mM NaCl. Those fractions containing the 45.5-kDa protein were dialyzed twice for 2 h apiece against ATPase buffer plus 0.1 µM ATP. Aliquots of purified protein were snap-frozen in liquid N₂ and stored at -80 °C. Before use, thawed kinesin aliquots were centrifuged (Heraeus Biofuge, 15,000 x g, 10 min) to remove debris. Active kinesin concentration was determined by measuring the time-dependent dissociation of kinesin-bound ADP as described (41).

Mammalian Brain Tubulin and Microtubule Preparation—Tubulin was extracted from bovine brain tissue by exploiting the temperature-dependent and reversible polymerization of the protein into microtubules (42, 43), and microtubule-associated proteins were then removed by DEAE-chromatography (44, 45). Microtubules were stored at -80 °C as centrifuged pellets. On the day of each experiment, pellets were thawed and resuspended in PEM buffer plus 1 mM GTP to a concentration of 10-15 mg/ml and depolymerized on ice for 20 min. Taxol was added stepwise to concentrations of 0.2, 2, and 20 µM, with 10-min incubations at 34 °C subsequent to each addition. The solution was then diluted 10-fold with PEM buffer plus 10 µM Taxol to stabilize the microtubules, and further incubated at 34 °C for 10 min. After centrifugation (Beckman JA-25.50 rotor, 39,000 x g, 30 min, 4 °C), the microtubule pellet was resuspended in ATPase buffer plus 20 µM Taxol. Microtubule concentration was determined using the method of Bradford (46), and reported molar concentrations of microtubules refer to the concentrations of -tubulin dimer (molecular mass of 110 kDa).

Nucleotide Analogs—N-Methylanthraniloyl (mant) derivatives of ATP and ADP were synthesized as described previously (47, 48). Spectrophotometric properties of the products were evaluated and conformed to published values. 2'(3')·mantADP and 2'(3')-mantATP are reported to have an A₂₅₅/A₃₅₆ ratio of 4.0, reflecting the optical densities of the N-methylanthraniloyl and adenine moieties (47). Previous work showed the 2'-mant-3'-dATP and 3'-mant-2'-dATP gave results similar to the mixture of isomers (2'-mant-ATP and 3'-mant-ATP) (14); therefore, we only used the mixture in these studies.

Phosphate Sensor—Phosphate release experiments measured the rate at which inorganic phosphate is released from kinesin, and relied on an engineered E. coli phosphate-binding protein (PBP-A179C) covalently coupled to a fluorescent dye MDCC (49, 50). The expression system for the protein component, consisting of the E. coli strain ANCC75 containing a pBR322-derived plasmid into which the modified phoS gene directing the production of PBP-A197C has been cloned, was obtained from M. Webb (National Institute for Medical Research, London, UK). PBP-A179C was expressed and purified as described (49). Conjugation of the protein with MDCC, and subsequent purification of the phosphate sensor, was performed according to the method described by Brune et al. (49), with subsequent modifications (50). The concentration of MDCC-PBP-A197C was determined spectrophotometrically, assuming an extinction coefficient at 280 nm of 68,575 M^-1 cm^-1. MDCC-PBP-A197C was divided into small aliquots, snap-frozen in liquid N₂, and stored at -80 °C.

Steady-state ATPase Assays—The hydrolysis of [-³²P]ATP by kinesin·microtubule complex was monitored at 35 °C by mixing labeled nucleotide with the enzyme, quenching the reaction after a predetermined time, separating the products on a TLC plate, and measuring the hydrolysis of the labeled nucleotide by standard radiation monitoring techniques as previously described (41). Velocity data for a range of substrate concentrations were then plotted and fit by nonlinear regression to a hyperbolic model, k_obs = k_cat[ATP]/(K_m-ATP + [ATP]) + C, to determine values of k_cat and K_m-ATP.

Sedimentation Equilibrium Study—A Beckman-Coulter Optima XL-1 analytical ultracentrifuge was used; it was fitted with an AnTi60 rotor and absorbance optics. Using three 6-channel charcoal-filled Epon centerpieces, nine kinesin concentrations could be evaluated simultaneously. Rotor speed was 13,000 rpm and run temperature was 24 °C. Equilibrium data were collected at 230 nm at a spacing of 0.003 cm with five averages in a step scan mode. Data sets were collected at 2-h intervals between 16 and 22 h after run initiation, and equilibrium was verified by comparing successive scans. Optical data were edited in Excel to extract data from individual channels and analyzed by nonlinear least-squares fitting to a self-association scheme using NONLIN (51), obtained from the Center for Analytical Ultracentrifugation of Macromolecular Assemblies at University of Texas Health Science Center at San Antonio. An estimate of 0.7350 cm·g^-1 for the partial specific volume of the kinesin monomer was made based on amino acid sequence contribution, taking into account a contribution of -0.0030 cm·g^-1 made by the bound ADP. Solvent density was determined volumetrically to be 1.0056 g·cm^-1. The extinction coefficient at 230 nm, the wavelength monitored during the analytical ultracentrifugation experiment, of KHC407A, was determined at 31,260 M^-1 cm^-1, and this value was used to convert the apparent association constants determined by NONLIN from optical density units to those of molarity according to the relationship K₂(M^-1) = K₂(abs^-1) x (1.2/2), where 1.2 is the optical path length in centimeters of the rotor centerpiece. NONLIN was used to fit equilibrium optical profiles of the sample channels to a simple monomer-dimer association reaction, with no assumed nonideality.

Rapid Quench Experiments—Transient-state kinetic analysis of kinesin ATPase in ATPase buffer was performed at 35 °C using a KinTek RQF-3 chemical quench flow instrument (KinTek Corp.). Reactions were quenched with 2 N HCl, and neutralized with 2 M Tris-3 M NaOH as described (24). 1.5 µl of each quenched sample was examined by polyethyleneimine-cellulose thin layer chromatography, developed with 0.6 M KH₂PO₄, pH 3.4.

Stopped-flow Experiments—A KinTek Stopped-Flow apparatus (Model SF-2001, KinTek Corp., Austin, TX) was used for stopped-flow experiments. Experiments were performed as previously described (14, 52) at 35 °C in ATPase buffer. Indicated reagent concentrations represent concentrations achieved after mixing.

Phosphate release kinetics were measured in the stopped-flow instrument, using the fluorescent phosphate reporter MDCC-PBP-A197C (11, 49) at a concentration of 4 µM and including the "phosphate mop," consisting of 0.1 mM 7-methylguanosine and 0.01 units/ml purine nucleoside phosphorylase. Excitation was at 425 nm, and fluorescence was measured using a 450 nm cutoff long-wave filter. Post mixing concentrations were 50 nM kinesin, 75 nM microtubules, and 500 µM ATP. Phosphate concentration was computed from the fluorescence based upon a standard curve. The time dependence of phosphate production was fitted to the burst equation [P_i] = A·exp(-k_obst) + k₂t + C, with k_obs representing the rate of phosphate release under the conditions tested.

The rate constants governing the binding of nucleotide to kinesin were estimated using mantADP and mantATP (15). Fluorescence data were fit to a single exponential function, F = A·exp(-k_obst) + C, where k_obs is the rate constant governing nucleotide binding and k₂ accounts for a slow, linear phase. Values of k_obs were plotted against nucleotide concentration and fit to a hyperbolic model: k_obs = k_max[mAD(T)P]/(K_d + [mAD(T)P]) + k_off. In experiments measuring rates of dissociation of mantADP, excitation of the fluorophore was direct, at 360 nm, and a decrease in fluorescence accompanied dissociation of mantADP from the active-site as described previously (14). Fluorescence traces were either fit to a double-exponential model of the form F = A₁·exp(-k₁t) + A₂·exp(-k₂t) + C, or else fit by simulated kinetic data.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
ATPase Pathway—The results presented in this report define the alternating site ATPase pathway shown in Fig. 1, which will be referenced to clarify the relationship between the data and this minimal model.

Active-site Titration—Isolated kinesin contains Mg-ADP tightly bound to the enzyme active-site (41, 53). To determine the concentration of kinesin active-sites in each preparation, the protein was incubated with [-³²P]ADP, and then the concentration of the bound radiolabeled nucleotide was measured by quantifying the amount that was inaccessible to a regenerating system that converted all free ADP to ATP (41). When phosphocreatine kinase, phosphocreatine, and cold ATP were added to a kinesin·[-³²P]ADP complex, all free [-³²P]ADP was rapidly converted to [-³²P]ATP while the remaining [-³²P]ADP was slowly lost from the kinesin active-site, observable by its slow conversion to [-³²P]ATP. The rate of disappearance of [-³²P]ADP equals the rate of dissociation of ADP from kinesin, and the amplitude extrapolated to t = 0 provides an estimate of the concentration of bound [-³²P]ADP at the start of the reaction. In Fig. 2, the results of an active-site determination experiment are shown. The concentration of the kinesin preparation was estimated by the absorbance at 280 nm as 56.5 µM. A 1:1 mixture of kinesin and [-³²P]ATP at 68.2 µM was prepared and allowed to come to equilibrium. The fraction of radiolabeled nucleotide, [ADP]/([ADP] + [ATP]), was plotted and fitted to a single exponential curve, F = A·exp(-k·t) + C, to yield rate of 0.0062 ± 0.0003 s^-1 and an amplitude of A = 0.286 ± 0.005. To correct for the dilution of radiolabeled ATP by the ADP already bound to the enzyme, we used the relationship [kinesin] = [ATP added]·A/(1 - A) to calculate the active-site concentration of 27.3 ± 0.5 µM. This value was then used as the active-site concentration of the preparation in all subsequent experiments. The active-site concentration decreased by <10% after 6 months at -80 °C and was equally stable after 5 days at 4 °C.

The rate of ADP release from the kinesin dimer determined in this assay was 0.0062 ± 0.0003 s^-1, a value comparable to that obtained by other methods. This measurement defines the rate-limiting step during the steady-state ATPase reaction when microtubules are absent.

Sedimentation Equilibrium Analysis of KHC407A—To determine whether KHC407A formed a dimer in solution, analytical ultracentrifugation using the sedimentation equilibrium protocol was used. Analysis of truncated Drosophila conventional kinesin constructs showed that a 366-residue N-terminal fragment of the protein does not self-associate in solution, whereas a 401-residue fragment forms a dimer with a dissociation constant of 36 nM. Thus, it has been shown that the domain necessary for self-association is between residues 367 and 401 in this protein, corresponding to the coil-coil domain seen in the crystal structure of rat kinesin (9). Based upon the structural and sequence similarities between Drosophila and rat conventional kinesins, we constructed a 407-residue N-terminal fragment of rat conventional kinesin expecting it to form a dimer. Nevertheless, it was necessary to examine the dimerization state of the purified protein.

View larger version (40K): [in this window] [in a new window]   FIGURE 1. Gilbert-Johnson Alternating site ATP hydrolysis pathway. This schematic was drawn using PDB file 3kin [PDB] (9). The red segment designates the position of the neck linker, which alternates between docked and undock positions, corresponding to tight and weak nucleotide binding states, respectively. The neck linker is undocked on the leading head (toward the right) leading to fast release of ADP. ATP does not bind to the leading head until the strain between the two heads is relaxed by the release of the trailing head from the microtubule, allowing the neck linker to dock. Steps a and b may be considered as priming steps that bring kinesin onto the microtubule to begin the alternating site cycle (steps 1-4). Release of ADP from the captive head state in the absence of ATP (step c) may become important at low ATP concentrations.

View larger version (8K): [in this window] [in a new window]   FIGURE 2. Active site titration of KHC407A. KHC407A-[-32P]ADP complex (28.3 µM kinesin and 34.1 µM [-32P]ADP) was rapidly mixed with phosphocreatine kinase (0.3 mg/ml), phosphocreatine (4 mM), and Mg-ATP (5 mM) in ATPase buffer. The ratio of [-32P]ADP to total labeled nucleotide is plotted against reaction time, as well as a best-fit single exponential curve F = A·exp(-k·t) + C. Parameter values obtained from the fitting were A = 0.286 ± 0.005, k = 0.0062 ± 0.0003 s-1, and C = 0.023 ± 0.004. An active-site concentration of 27.3 ± 0.5 µM was estimated, based on values for A and the concentration [-32P]ATP used in the initial labeling, according to the relationship [kinesin] = ([ATP]·A)/(1 - A).

View larger version (26K): [in this window] [in a new window]   FIGURE 3. Sedimentation equilibrium analysis of KHC407A. KHC407A was analyzed by analytical ultracentrifugation using the sedimentation equilibrium protocol. Rotor speed (AnTi60) was 13,000 rpm, run time 20 h, and run temperature 24 °C. Enzyme concentrations, according to active-site titration, were 44, 154, 264, 374, 484, 594, 704, 814, and 924 nM. A, optical profiles at 230 nm of protein samples at equilibrium, taken at 0.003 cm resolution (x) and curves representing a global fitting of data using NONLIN to a monomer-dimer model (solid curves). Global fitting indicated a value for K2, the dimer association equilibrium constant, of 2.15 x 107 M-1 (9.25 x 106 and 5.70 x 107). Values in parentheses are the 95% confidence interval calculated by NONLIN. B-D, residuals for each of the nine samples.

View larger version (8K): [in this window] [in a new window]   FIGURE 4. Steady-state ATP hydrolysis by KHC407A in the presence of microtubules. Hydrolysis rates of [-32P]ATP by KHC407A were determined at substrate concentrations of 1 to 200 µM. 5 nM KHC407 and 60 µM microtubules were incubated with labeled substrate, and a time course created. The observed relationship between hydrolysis rate and substrate concentration was used to generate a best-fit curve kobs = kcat[ATP]/(Km-ATP + [ATP]) + C, whose parameters yield estimates of kcat and Km-ATP. From the data and subsequent curve fitting, kcat = 40.3 ± 1.1 s-1 and Km-ATP = 53.6 ± 4.8 µM.

View larger version (16K): [in this window] [in a new window]   FIGURE 5. Pre-steady-state binding of mantATP to KHC407A·microtubule complex. A, a representative trace of fluorescence change incurred by KHC407A·microtubule complex upon rapid mixing with mantATP. Concentrations after mixing were 2 µM kinesin, 10 µM microtubules, and 50 µM mantATP. The dashed line indicates a best-fit curve F = A·exp(-kobst) + k2t + C. Similar curves were generated for mantATP concentrations of 5, 10, 20, 30, 50, 70, and 100 µM mantATP. B, values for kobs obtained from fluorescence traces at each mantATP concentration are plotted against the independent variable. The solid curve represents the best-fit hyperbola kobs = kmax[mATP]/(Kd + [mATP]) determined by nonlinear regression. Convergence of parameters provide estimates of kmax at 210 ± 25 s-1 and Kd at 38 ± 10 µM. The ratio kmax/Kd provides a lower limit to the apparent second-order rate constant for mantATP binding to the complex, and is 5.6 ± 1.7 µM-1 s-1.

Binding of mantATP to KHC407A—To obtain an estimate of the rate constant governing the binding of ATP to KHC407A, a fluorescent ATP analog was used in a stopped-flow experiment, in which the kinesin·microtubule complex was mixed with mantATP. Fluorescence resonance energy transfer between optically excited tryptophan residues in the protein and the N-methylanthraniloyl moiety of mantATP provided a means by which the binding rate can be measured. Excitation was at 280 nm, and fluorescence was detected by a photomultiplier tube fitted with a 400 nm cutoff long-wave pass filter. Although the experiment measures the binding kinetics of a substrate analog rather than those of the substrate itself, the results are considered a close approximation of the behavior of the enzyme toward its natural substrate, because k_cat and K_m for the fluorescent analog are within a factor of two of the corresponding values for ATP (15). Concentrations after mixing were 2 µM KHC407A, 10 µM microtubules (in a preformed kinesin·microtubule complex), and mantATP at concentrations ranging from 5 to 100 µM. Fig. 5A shows a representative trace at 50 µM mantATP. A binding rate (k_obs) was extracted from each trace by fitting the data by nonlinear regression to a single exponential function, F = A·exp(-k_obst) + k₂t + C. The linear constant k₂ accounted for a slow, linear increase in signal after the transient, which is not deemed to be mechanistically significant, but improves the accuracy of the fit to the transient.

In Fig. 5B values of k_obs determined for different concentrations of mantATP are plotted. A best-fit hyperbola of the form, k_obs = k_max-[mATP]/(K_d + [mATP]) + k_off, was obtained by nonlinear regression. The rate constant for ATP dissociation (k_off) could not be accurately determined by this experiment and was set to zero. The maximum achievable binding rate, to which k_obs converges as the concentration of mantATP increases (k_max), was evaluated at 210 ± 25 s^-1 and, the apparent K_d was 38 ± 10 µM. We also considered an alternative interpretation of the data based upon a two step binding sequence where the first step was not a rapid equilibrium with k₁ = 5.6 µM^-1 s^-1 and k₂ = 210 s^-1. However, this model would predict that a lag should be seen in the kinetics at moderate concentrations (10-20 µM), and the failure to see a lag argues in favor of a rapid equilibrium binding for the collision complex.

View larger version (9K): [in this window] [in a new window]   FIGURE 6. Pre-steady state hydrolysis of ATP by KHC407A. Microtubule-dependent ADP formation by KHC407A was monitored in the quench-flow instrument. KHC407A·microtubule complex was rapidly mixed with [-32P]ATP (post-mixing concentrations 2.18 µM KHC407A, 10 µM microtubules, and 100 µM [-32P]ATP) and [-32P]ADP formation examined by TLC. A linear fit (dashed line) was generated from the data (circles), estimating the hydrolysis rate at 34.5 ± 1.7 s-1 and the burst amplitude at 0.85 ± 0.07. A KinTekSim simulation using the reaction depicted in Fig. 1 (solid line) suggests values for kinetic parameters k2 (ATP hydrolysis) at 523 ± 58 s-1, and k3 (ADP release) at 43.6 ± 2.9 s-1. The rate of ATP binding was set at 5.6 µM-1 s-1 based on results from mantATP binding experiments.

Reaction 1

The existence of such a complex, although not certain, is supported by evidence from kinetic and structural studies, which suggested the possibility of two distinguishable sequential kinesin·ATP complexes (11, 28, 55). The observed single exponential in the fluorescence traces at all mantATP concentrations supports the conclusion that the initial binding step is a rapid equilibrium, and so, the fit to a hyperbola defines a ground state dissociation constant of 1/K₁ = 38 µM, which is followed by a rate-limiting isomerization step (k₂ = 210 s^-1), which could be coincident with mantATP hydrolysis (see below, 520 s^-1 rate for ATP). The initial slope of the concentration dependence of the rate defines the apparent second order rate constant for ATP binding and is equal to K₁k₂ = 5.6 ± 1.7 µM^-1 s^-1, which puts a lower limit on the magnitude of the rate of step 1 (Fig. 1).

Pre-steady-state Kinetics of ATP Hydrolysis—To examine the kinetics of ATP hydrolysis at the active-site, a rapid quench experiment was performed, whereby the KHC407A·microtubule complex was rapidly mixed with [-³²P]ATP, allowed to react for a predetermined period, and then quenched with acid.

A kinesin·microtubule complex was mixed with [-³²P]ATP in the quench-flow instrument such that post mixing concentrations were 2.18 µM KHC407A, 10 µM microtubules, and 100 µM [-³²P]ATP. Fig. 6 shows the concentration of [-³²P]ADP versus time. The data are fit best to a linear model (dashed line) with slope and y-intercept determined at 75 ± 2 µM^-1 s^-1 and 1.8 ± 0.2 µM, respectively. This reaction appears to have a steady-state rate of 34 ± 2 s^-1, calculated from the hydrolysis rate divided by the enzyme concentration. This value, short of k_cat (40 ± 1 s^-1) for the reaction by 14%, reflects the sub-saturating concentration of labeled ATP used in this experiment (100 µM) and is consistent with the results of steady-state determination. The rapid evolution of product, preceding steady-state turnover, is expected to conform to a burst equation [ADP] = A·exp(-k_obst) + k_sst + C. The burst phase occurred too rapidly to be resolved by the procedure used and was essentially completed before the collection of earliest data at 5 ms. However, we could estimate a rate of the burst of 520 s^-1 by comparison of the amplitude of the burst and the value of k_cat according to the following mechanism (Reaction 2).,

REACTION 2

The rate constants k₂ and k₃, representing the rate constants governing ATP hydrolysis and ADP release, respectively, were estimated indirectly from the burst amplitude of the quench flow data and the value of k_cat, determined directly by steady-state methods. The burst amplitude (0.85 ± 0.07), a dimensionless number, is defined by the y-intercept of the linear extrapolation of the data in Fig. 6 divided by the active-site enzyme concentration. The values of k₂ and k₃ can be calculated from the relationships, burst amplitude A = [k₂/(k₂ + k₃)]² and k_cat = k₂k₃/(k₂ + k₃), with known amplitude A and steady-state rate constant k_cat (56), assuming that k_-2 is negligible. Simultaneous solution of these two equations yields values of k₂ = 520 ± 60 s^-1 and k₃ = 44 ± 3 s^-1. This analysis depends upon the assumption that hydrolysis is not readily reversible; however, the consequences of this assumption are minor, and in either case the apparent rate of the burst = k₂ + k_-2 = 520 s^-1. For example, if K₂ = 4 as in the case of skeletal muscle myosin, then the data would be fit by the parameters k₂ = 416 s^-1, k_-2 = 104 s^-1, and k₃ = 56 s^-1. If the rate of the burst had been measurable, then the assumption that k_-2 was negligible would not have been necessary and all rate constants could have been solved by simultaneous solution of three equations defining the burst rate, burst amplitude, and k_cat.

To illustrate the expected burst kinetics of KHC407A, the reaction was simulated using KinTekSim software, programmed with a simple three-step reaction mechanism described above and using estimates of each rate constant in the pathway. The apparent second-order rate constant for ATP binding to kinesin·microtubule complex (k₁) was previously estimated to be 5.6 µM^-1 s^-1, so this value was used in the simulation. Fig. 6 shows the data, the linear fit to the data (dashed line), and the simulated curve (solid line). It is important to note that values obtained for k₂ and k₃ describe the hydrolysis of ATP and release of ADP, respectively, according upon the three-step model described above. If kinesin hydrolyzes ATP using an alternating site mechanism as has been proposed (13-15), then the hydrolysis of [-³²P]ATP by the active-site to which it has bound will not immediately follow the binding step but will be delayed until the prescribed conformational changes occur within the other subunit of the kinesin dimer. Similarly, the alternating site model suggests that the release of product from one site may occur only after prerequisite rearrangements occur at the other. For this reason, values of k₂ and k₃ describe what are likely to be composite reactions that incorporate more than one distinguishable step. Nonetheless, the data demonstrate two important points. ATP hydrolysis is faster than steady-state turnover, and only one ATP is hydrolyzed prior to the step limiting the steady-state rate. We next examine the kinetics of phosphate release.

View larger version (14K): [in this window] [in a new window]   FIGURE 7. Pre-steady-state phosphate release kinetics of KHC407A. Kinesin·microtubule complex (0.05 µM kinesin and 0.075 µM microtubules) was mixed with ATP (500 µM)in the stopped-flow instrument in the presence of MDCC-PBP (4 µM) and a "phosphate mop" (1 unit/ml purine nucleoside phosphorylase and 0.1 mM 7-methylguanosine). For the upper trace, 100 mM KCl was included. All concentrations are post-mixing. Fluorescence change was converted to phosphate concentration based on a standard curve and normalized by dividing by the kinesin concentration. Fitting to a double- or single-exponential equation, including a linear phase generated the solid curves representing experiments performed with and without KCl, respectively. Kinetic parameters are summarized in TABLE ONE.

Fig. 7 shows the release of phosphate from KHC407A·microtubule complex after rapid mixing with ATP, normalized by dividing by the kinesin concentration. Final concentrations after mixing were 0.05 µM kinesin, 0.075 µM microtubules, and 500 µM ATP. The experiment was performed in ATPase buffer, with and without 100 mM KCl (upper and lower traces, respectively). The purpose of the added salt was to destabilize the kinesin·microtubule interaction, so as to promote dissociation after a single turnover as described previously (54) in an attempt to measure the rate of phosphate release after the hydrolysis of ATP. Kinetic parameters are summarized in TABLE ONE.

In the presence of KCl, the time dependence of phosphate release fits a double exponential. The fast phase has a rate of 510 s^-1 and amplitude of 0.8 per kinesin, whereas the slow phase occurs at a rate of 62 s^-1 and amplitude of 1.2 per kinesin. Interestingly, the elevated salt concentration appears to accelerate the rate of release of the first phosphate after ATP hydrolysis. Because the rate of phosphate release is linked to dissociation of the trailing head from the microtubule, it appears as though salt weakens the interaction of the head with the microtubule to accelerate its release.

Binding of KHC407A to Microtubules—The binding of kinesin to the microtubule is the first step in microtubule-dependent kinesin motility. Two methods were employed to examine the kinetics of kinesin binding to microtubules. Using the first method, binding was measured directly by exploiting the change in turbidity that accompanies kinesin·microtubule association. In the second method, which will be described in the next section, the release of mantADP from kinesin upon binding to microtubules served as a reporter for the initial association reaction.

KHC407A (2 µM after mixing) was rapidly mixed with microtubules at concentrations between 5 and 15 µM (after mixing) in the stopped-flow apparatus in the absence of nucleotide, and the intensity of 340 nm light transmitted through the mixture was monitored, which was used to compute the turbidity (defined by the natural logarithm of the intensity change). Fig. 8A shows a trace for 2 µM KHC407A rapidly mixed with 5 µM microtubules. The dashed line represents a fitted double exponential curve of the form T = A₁·exp(-k₁t) + A₂·exp(-k₂t) + C. A similar analysis was performed on each turbidity trace in the microtubule concentration dependence series. For each microtubule concentration, a fast and a slow phase were identified based on the relative magnitudes of k₁ and k₂, and these rates were plotted against microtubule concentration, as depicted in Fig. 8B.

The rate of the fast phase (circles) shows a linear increase with microtubule concentration, whereas the slow phase (squares) is nearly constant. The slope of a linear fit to the fast phase data (2.9 ± 0.2 µM^-1 s^-1) provides an estimate of the apparent second-order binding rate constant for KHC407A·microtubule association, whereas the y-intercept (19 ± 2.2 s^-1) estimates the rate constant governing dissociation. A dissociation equilibrium constant K_d = 6.7 ± 0.9 µM is indicated from these two measurements. The significance of the slow phase data is not immediately clear. If the measured rates of the slow phase are relevant to the kinetics of kinesin·microtubule interaction, their independence from microtubule concentration suggests that they describe a process that is most likely first-order with respect to kinesin, the reaction component that is held constant at 2 µM throughout the series of experiments. For this reason, the fast phase rates are considered as indicating a second-order binding process, whereas the slow phase rates indicate an unknown first-order process, perhaps reflecting an isomerization or aggregation of the microtubule·kinesin complex. The slow phase could represent a change in structure leading to tighter binding of the kinesin to the microtubule complex; however, the measured rates are too slow to be part of the ATPase cycle.

View larger version (16K): [in this window] [in a new window]   FIGURE 8. Pre-steady-state binding of KHC407A to microtubules. A, representative turbidity trace at 340 nm illumination of kinesin and microtubules after mixing (2 µM kinesin, 5 µM microtubules, post-mixing). The dashed line represents a best-fit double exponential curve T = A1·exp(-k1t) + A2·exp(-k2t) + C, whose parameters A1, A2, k1, k2, and C were determined by nonlinear regression. Similar traces were recorded for microtubule concentrations of 5, 7, 9, 11, 13, 15, and 17 µM microtubules, each of which were fit by double-exponential models. B, both fast (circles) and slow (squares) rates were plotted against microtubule concentrations. The ratios of the amplitudes (fast phase amplitude/slow phase amplitude) are also shown (triangles). Fast rates were fit by linear regression, whose gradient of 2.9 ± 0.2 µM-1 s-1 estimates the second-order rate constant for kinesin·microtubule association. A y-intercept at 19 ± 2.2 s-1 is the apparent rate constant for dissociation of the complex.

View larger version (15K): [in this window] [in a new window]   FIGURE 9. Microtubule dependence of mantADP release from KHC407A. A, representative trace of fluorescence decay resulting from rapid mixing of kinesin·mantADP complex with microtubules and ATP. Post-mixing concentrations were 2 µM kinesin, 4 µM mantADP, 18 µM microtubules, and 1 mM ATP. The data were normalized by dividing by the starting fluorescence. A best-fit double exponential curve F = A1·exp(-k1t) + A2·exp(-k2t) + C (dashed) was determined by nonlinear regression. Parameters A1, A2, k1, k2, and C were determined for microtubule concentrations at 8, 12, 13.5, 15, 16.5, 18, 19.5, 21, and 24 µM. At 30, 60, and 80 µM, model fitting was best achieved using a single exponential model. B, fast- (filled circles) and slow- (open circles) phase rates are plotted against microtubule concentration, and fast phase rates were fit by linear regression. The gradient of the linear fit, 4.6 ± 0.1 µM-1 s-1, predicts the second order rate constant for microtubule·kinesin association, whereas a y-intercept of 30 ± 3.3 s-1 is the apparent rate constant for dissociation.

KHC407A·mantADP complex was rapidly mixed with microtubules at concentrations from 8 to 80 µM (after mixing) in the stopped-flow instrument. Fig. 9A shows a representative trace of time-dependent fluorescent decay after mixing 2 µM KHC407A·mantADP complex with 18 µM microtubules and 1 mM ATP. Like the turbidity data described in the previous section, the fluorescence traces from this experiment were best fit to a double exponential rather than a single. Fluorescence data at the highest microtubule concentrations could only be fitted to single exponential models.

Fast and slow phase rates are plotted in Fig. 9B, as well as a linear fit to the fast phase rates. As described below, the signal in this case arises from the sequential release of two ADP molecules and the binding of one ATP molecule. Therefore, complete analysis of the time course requires fitting the data to four step model where represents kinesin with ADP bound to each site (Reaction 3).

REACTION 3

In this experiment, the high ATP concentration makes step 3 very much faster than step 2, and the microtubule concentration dependence of the rate of release of ADP therefore provides a measurement of the microtubule binding rate. The observed rate of a reversible binding reaction is the sum of the forward and reverse rates, k_obs = k₁[Mt] + k_-1. Curve fitting by linear regression sets the forward and reverse rates at 4.6 ± 0.1 µM^-1 s^-1 and 30 ± 3.3 s^-1, respectively. These values are in reasonable agreement with the corresponding rate constants obtained more directly by turbidity analysis (2.9 ± 0.2 µM^-1 s^-1 for binding, 19.5 ± 2.2 s^-1 for release), and cover a larger range of microtubule concentrations. Interestingly, despite the differences between the results obtained using turbidity and mantADP release methods, the apparent equilibrium dissociation constants for the initial microtubule·kinesin collision complex suggested by each experiment are nearly identical, 6.5 ± 0.7 µM indicated by the mantADP dissociation reaction, and 6.6 ± 0.9 µM by the turbidity assay. However, this may not represent a true dissociation constant, because the observed release of ADP is so fast. Therefore, ADP release (step 2) may be reversible in order for the net microtubule dissociation rate to contribute to the concentration dependence of the observed ADP release rate such that the observed rate of 20-30 s^-1 may represent a composite of k_-1 and k_-2.

View larger version (14K): [in this window] [in a new window]   FIGURE 10. Pre-steady-state ATP dependence of mantADP release from KHC407A. KHC407A·mantADP complex (1 µM enzyme active-site, 2 µM mantADP) was rapidly mixed with microtubules (20 µM) plus ADP (10 µM), or ATP (5, 10, 25, and 50 µM), and fluorescence decay monitored. Concentrations are post-mixing. Solid lines represent normalized fluorescence data. Dashed lines are kinetic simulations of the reaction, based on a reaction depicted in Fig. 1, rate constants in TABLE TWO, and initial reagent concentrations used in the experiments.

ATP Dependence of mantADP Release from KHC407A—In previous analysis of conventional kinesin motility, two ADP release events were observed after kinesin bound to microtubules (13-15), the second of which was dependent upon ATP concentration. To examine the ATP concentration dependence of mantADP release, a KHC407A·mantADP complex was rapidly mixed with microtubules (20 µM) plus nucleotide (ATP at 5, 10, 25, and 50 µM, or ADP at 10 µM) in the stopped-flow instrument, and the fluorescence was monitored by direct excitation of the fluorophore 360 nm.

The data from this experiment resulted in a family of curves displayed in Fig. 10. Each trace was normalized to its initial signal intensity and displaced along the x-axis to account for an instrument dead time of 1.5 ms so that the curves can be superimposed and compared with the results of computer simulation. In the absence of ATP, release of mantADP is clearly biphasic with rates of 140 ± 2s^-1 and 1.44 ± 0.02 s^-1 for the release of the first and second ADP, respectively. Observation of the second ADP release is dependent upon the addition of 10 µM ADP to prevent the rebinding of mantADP. Thus, the release of the second ADP reaches equilibrium at a point that favors rebinding of the second ADP, and interactions with the microtubule stimulate the ADP exchange.

Increasing the concentration of ATP increases the rate of release of the second mantADP, which results in an observed increase in the amplitude of the fast reaction phase. At intermediate ATP concentrations, the observed reaction is a function of at least three steps, and their rates are not sufficiently different to be resolved meaningfully by conventional data fitting to a sum of exponential functions. Rather, the reaction sequence summarized in Fig. 1 was used to globally fit the data by computer simulation. In Fig. 10, dashed lines represent simulated data based on rate constants summarized in TABLE TWO.

The most striking results of this experiment are the predicted rate constants for mantADP dissociation. A value of greater than 1000 s^-1 for both k₊₂ and k_+b were required for the fit seen in Fig. 10. Under conditions of the experiment, the release of the first ADP is limited by the rate of kinesin binding to the microtubule and therefore is not well defined. This was not the case for the Drosophila K401 N-terminal truncation of conventional kinesin, where the rate of release of mantADP was well defined at a rate constant of 300 s^-1 (14).

On the longer time scale, there is a regain in fluorescence due to the partial rebinding of mantADP after the completion of ATP hydrolysis. This behavior was not seen in similar studies performed using the Drosophila K401 dimeric N-terminal truncation, which generated a family of traces converging to a common minimum within 1-2 s after mixing (14). Thus, the observations made using KHC407A suggest a qualitative difference affinity of the two enzymes for mantADP. Rat kinesin in complex with microtubules binds one molecule of ADP more strongly than does Drosophila kinesin. The reaction pathway for KHC407A simulation required the inclusion of a mantADP release step in the absence of ATP (step 6), occurring with a rate constant k_+c of 1 s^-1, to account for the slow decay in the presence of 10 µM ADP (0 µM ATP).

ADP Dependence of mantADP Release from KHC407A—The ATP-independent release of mantADP observed in the experiment described in the previous section prompted an investigation of the ability of ADP to stimulate mantADP release. A KHC407A·mantADP complex was mixed in the stopped-flow instrument with microtubules plus ADP. Post mixing concentrations were 1 µM KHC407A (active-site), 2 µM mantADP, 20 µM microtubules, and ADP at concentrations at 5, 10, 20, 50, and 250 µM. Fig. 11A shows the family of fluorescence traces from this experiment, each normalized to its initial intensity. Note that the bottom curve, with the steepest rate of descent, represents the rate of ATP-stimulated mantADP release, having been generated by mixing KHC407A·mantADP with 20 µM microtubules plus 250 µM ATP. This curve does not contribute to the calculation of an ADP binding rate constant and is included here only to provide a reference amplitude.

View larger version (20K): [in this window] [in a new window]   FIGURE 11. ADP dependence of mantADP release from KHC407A. A, the set of fluorescence traces resulting from the rapid mixing of kinesin·mantADP complex with microtubules plus ADP. Post-mixing concentrations were 1 µM kinesin, 2 µM mantADP, 20 µM microtubules, and ADP at concentrations of 5, 10, 20, 50, and 250 µM, or ATP at 250 µM (top curve to bottom curve). Curves were fit to double-exponential models (F = A1·exp(-k1t) + A2·exp(-k2t) + C) by nonlinear regression (fit curves not shown), generating a set of parameters A1, A2, k1, k2, and C for each. B, fast-phase rates are plotted against ADP concentration, with a best-fit hyperbola of the form kobs = kmax[ADP]/(Kd-ADP + [ADP]) + koff. The parameters obtained by nonlinear regression estimate of kmax = 3.9 ± 0.3 s-1, Kd-ADP = 98 ± 31 µM, and koff = 0.8 ± 0.3 s-1. The ratio kmax/Kd-ADP (0.04 ± 0.01 µM-1 s-1) estimates the apparent second order rate constant for ADP binding to the stalled KHC407A·microtubule complex.

Binding of mantADP to KHC407A—We previously showed in Fig. 10 that after the release of mantADP from KHC407A upon binding to microtubules plus ATP there is an apparent reversal in the fluorescence decay that accompanies the conversion of mantATP to mantADP on the 0.1- to 3-s time scale. The most straightforward explanation for this behavior maintains that mantADP rebinds to the active-site of the enzyme. To further examine the interaction between mantADP and KHC407A, stopped-flow experiments were performed.

KHC407A·microtubule complex was formed and rapidly mixed with mantADP at various concentrations in the stopped-flow instrument. Based upon other measurements, we presume that at the start of this experiment kinesin will be bound to the microtubule in the captive head state, with one head tightly associated with the m