©1996 by The American Society for Biochemistry and Molecular Biology, Inc.
Equilibrium Studies of Kinesin-Nucleotide Intermediates (*)

(Received for publication, September 15, 1995; and in revised form, January 17, 1996)

Steven S. Rosenfeld (1) (2)(§) Brenda Rener (1) John J. Correia (5) Matthew S. Mayo (4) Herbert C. Cheung (3)

From the  (1)Departments of Neurology, (2)Cell Biology, and (3)Biochemistry and the (4)Division of Biostatistics, UAB Comprehensive Cancer Center, University of Alabama at Birmingham, Birmingham, Alabama 35294 and the (5)Department of Biochemistry, University of Mississippi Medical Center, Jackson, Mississippi 39216

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

We have examined the energetics of the interactions of two kinesin constructs with nucleotide and microtubules to develop a structural model of kinesin-dependent motility. Dimerization of the constructs was found to reduce the maximum rate of the microtubule-activated kinesin ATPase 5-fold. Beryllium fluoride and aluminum fluoride also reduce this rate, and they increase the affinity of kinesin for microtubules. By contrast, inorganic phosphate reduces the affinity of a dimeric kinesin construct for microtubules. These findings are consistent with a model in which the kinesin head can assume one of two conformations, ``strong'' or ``weak'' binding, determined by the nature of the nucleotide that occupies the active site. Data for dimeric kinesin are consistent with a model in which kinesinbulletATP binds to the microtubule in a strong state with positive cooperativity; hydrolysis of ATP to ADP+P(i) leads to dissociation of one of the attached heads and converts the second, attached head to a weak state; and dissociation of phosphate allows the second head to reattach. These results also argue that a large free energy change is associated with formation of kinesinbulletADPbulletP(i) and that this step is the major pathway for dissociation of kinesin from the microtubule.


INTRODUCTION

Molecular motors power a wide variety of physiologically important motile processes. These include movements of intracellular organelles, of chromosomes during mitosis and meiosis, and of cytoskeletal components during the process of ameboid motion (Vallee and Shpetner, 1990; Endow and Titus, 1992). These enzymes can be broadly classified into two categories: the myosins, which generate movement along actin-containing microfilaments; and a group of microtubule-based motors that include cytoplasmic dynein and the kinesin family of mechanoenzymes (Endow and Titus, 1992). The myosins remain the best studied group of molecular motors, and much effort has gone into identifying which of the steps in the actomyosin ATPase cycle are responsible for force generation.

Studying the nature of these myosin-nucleotide intermediate states has been facilitated by the use of transition metals, which bind to myosinbulletADP stoichiometrically and with high affinity (Goodno, 1979; Phan and Reisler, 1992; Maruta et al., 1993). Complexes of these metals with myosinbulletADP appear to mimic either the myosinbulletATP or myosinbulletADPbulletP(i) structures (Fisher et al., 1994). Thus, aluminum fluoride appears to induce a prehydrolytic myosin-nucleotide transition state, whereas beryllium induces a myosinbulletATP structure (Fisher et al., 1994). The stability of these complexes has allowed their study with spectroscopic, NMR, and crystallographic methods and has provided insight into the structure of the short lived myosinbulletATP and myosinbulletADPbulletP(i) intermediates. The validity of their use in studying myosin-nucleotide intermediates has also been supported by the effects of inorganic phosphate. Phosphate can bind to the active site of myosinbulletADP to generate a myosinbulletADPbulletP(i) state, and it has effects that are physiologically similar to those of vanadate and aluminum fluoride in reducing the affinity of myosinbulletADP for actin (Dantzig and Goldman, 1985).

Compared with the myosins, the kinesin family of microtubule motors appears to have to comply with a different set of physiologic constraints. Kinesin powers movement of organelles along microtubules and, unlike myosin, appears to operate in isolation (Walker and Sheetz, 1993). Motility studies in vitro are consistent with this assignment of function, as single kinesin molecules are able to translocate along microtubules for several micrometers at maximal velocity without detaching (Howard et al., 1989). These differences in physiology suggest that the nature of the force-generating transition(s) in the kinesin-microtubule ATPase cycle may likewise be different. Support for this comes from an in vitro motility study (Romberg and Vale, 1993) which demonstrated that ATPS, (^1)vanadate, and aluminum fluoride prolonged the lifetime of attachment of kinesin to the microtubule; and increasing concentrations of ADP shortened this lifetime. These findings were interpreted to mean that the kinesinbulletADPbulletP(i) state (presumably mimicked by aluminum fluoride, vanadate, and ATPS) was strongly bound, whereas the kinesinbulletADP state was weakly bound. However, more recent kinetic studies (Gilbert et al., 1995) of the kinesin ATPase cycle could be explained by either of two models. In one, dissociation of the kinesin-microtubule complex occurs in the kinesinbulletADPbulletP(i) state, suggesting that this state may be weakly bound, whereas in the other, dissociation occurs in a transition involving a kinesinbulletADP intermediate state:

where K is kinesin, M is microtubule, T is ATP, and D is ADP.

Determining which of the above models is the most accurate depiction of the kinesin ATPase cycle requires direct measurements of the binding affinities of the various kinesin-nucleotide intermediate states and the equilibrium constants for the various kinesin-nucleotide transitions. Efforts in this regard have been made by several laboratories, which have examined the steady and pre-steady-state kinetics of these transitions by utilizing bacterially expressed recombinant fragments of kinesin which contain the amino-terminal motor domain and variable amounts of the carboxyl-terminal tail (Huang and Hackney, 1994; Huang et al., 1994; Hackney, 1994a, 1994b; Gilbert and Johnson, 1993, 1994; Gilbert et al., 1995; Ma and Taylor, 1995a, 1995b). These studies have confirmed previous observations using intact kinesin and have extended them by demonstrating that: 1) ATP binding, hydrolysis, and phosphate release are rapid relative to subsequent steps in the hydrolysis cycle; 2) kinesinbulletADP is the predominant species in the system, and release of ADP is the rate-limiting step; 3) microtubules accelerate ADP release several thousandfold; and 4) for dimeric kinesin constructs that contain ADP in the active site, microtubules accelerate release of only one of the two bound ADP molecules. However, these studies have not measured binding affinities of stable kinesinbulletATP and kinesinbulletADPbulletP(i) intermediates and are thus not able to assign microtubule affinities reliably to several of the kinesin-nucleotide states depicted above.

In this study, we have generated two bacterially expressed constructs of human kinesin and examined the effects of ADP, aluminum fluoride, beryllium fluoride, and inorganic phosphate on their binding affinities and steady-state ATPase parameters. These studies indicate that ternary complexes of kinesinbulletADP with salts of aluminum and beryllium mimic the kinesinbulletATP state, which is strong binding; that inorganic phosphate reduces the affinity of dimeric kinesinbulletADP for microtubules; and that the dissociation of the kinesinbulletADPbulletP(i) complex from microtubules represents the major dissociation step in the kinesin-microtubule ATPase mechanism.


EXPERIMENTAL PROCEDURES

Materials

Oligonucleotides used in the polymerase chain reaction were synthesized by Oligos Etc. (Guilford, CT) and Cruachem (Sterling, VA). Restriction enzymes and modifying enzymes were obtained from Stratagene, Inc. (La Jolla, CA) and Life Technologies, Inc. Taq polymerase was also supplied by Life Technologies, Inc. Media components were obtained from Difco Laboratories. Protease inhibitors, antibiotics, GTP, malachite green oxalate, Triton X-100, and chemicals used for buffers and agarose gel electrophoresis were obtained from Sigma. Isopropyl-beta-D-thiogalactopyranoside was obtained from Fisher Scientific. Ribonuclease A and deoxyribonuclease I were purchased from U. S. Biochemical Corp. Prepacked Q-Sepharose columns, protein assay dye reagent, and chemicals used in sodium dodecyl sulfate-polyacrylamide gel electrophoresis were from Bio-Rad. Ni-NTA agarose was obtained from Qiagen (Chatsworth, CA). Centriprep-10 concentrators were obtained from Amicon, Inc. (Cherry Hill, NJ). N-Succinimidyl [2-3-^3H]propionate, was purchased from American Radiolabeled Chemicals, Inc. (St. Louis, MO). Taxol was generously provided by Nancita Lomax of the Drug Synthesis and Chemistry Branch, Division of Cancer Treatment, National Cancer Institute. mant and 2`-deoxy mant nucleotides were synthesized from the unlabeled nucleotides as described (Hiratsuka, 1983) and purified by chromatography in water on Sephadex LH-20.

Kinesin Expression in Escherichia coli and Purification of Recombinant Proteins

pXPE plasmid DNA encoding human kinesin was kindly provided by Dr. Ron Vale (described in Navone et al., 1992). DNA fragments encoding the first 413 amino acids (K413) and the first 332 amino acids (K332) of human kinesin were generated by polymerase chain reaction using the following oligonucleotide primers. The amino-terminal primer for both K413 and K332 was 5`-GCGGACCTGGCCGAGTGCAACATC-3`. The carboxyl-terminal primer for K413 was 5`-GGTACTCGAGAAAATTTCCTATAACTCCA-3`, containing an XhoI restriction site. The carboxyl-terminal primer for K332 was 5`-GGTACTCGAGATTGACACAAACTGTGTTC-3`, containing an XhoI restriction site. Each DNA fragment was ligated to the vector pET-21d (Novagen, Madison, WI), which had been digested with NcoI and repaired. This vector generates a fusion protein in which the kinesin insert is fused at the carboxyl terminus to a sequence of six histidine residues, which allows for affinity purification on Ni-NTA agarose (see below). The ligation mixtures were used to transform E. coli DH5F`alpha as described (Sambrook et al., 1989). DNA from these clones was prepared by a modification of the alkaline lysis minipreparation method, restriction digested, and analyzed by agarose gel electrophoresis. Positive DNA was used to transform E. coli BL21(DE3) as described (Sambrook et al., 1989) for expression of each recombinant protein. Transformants were selected on plates of LB with 100 µg/ml ampicillin. For purification of protein, 10 liters of culture were grown at 37 °C and 300 rpm in LB with 100 µg/ml ampicillin to an absorbance at 595 nm of 0.6. Cultures were induced for 4 h by the addition of isopropyl-beta-D-thiogalactopyranoside to 0.5 mM. Cells were harvested by centrifugation and stored at -70 °C. The frozen cells were thawed and resuspended in 3 ml of cold lysis buffer (50 mM Tris, pH 7.9, 10% sucrose, 0.3 M NaCl, 5 mM MgCl(2), 0.5 mM MgADP, 1 mM phenylmethylsulfonyl fluoride, 2 µg/ml aprotinin, 2 µg/ml leupeptin, 1 µg/ml pepstatin A)/g of cells. Lysozyme was added to 1 mg/ml, and the suspension was kept on ice for 30 min with occasional mixing. The sample was sonicated to shear the DNA. Ribonuclease A was added to 10 µg/ml, deoxyribonuclease I was added to 5 µg/ml, and the suspension was incubated at room temperature for 30 min. The sample was clarified by centrifugation at 27,000 times g for 20 min. The clarified sample was passed through a 0.45-µm filter. Twelve ml of a 1:1 suspension of Ni-NTA agarose preequilibrated in lysis buffer was added to the clarified lysate. The sample was mutated at 4 °C for 2 h. The resin was pelleted and loaded into a 0.5 times 30-cm column at 4 °C. The column was washed for 3 h at 0.5 ml/min with wash buffer (20 mM Tris, pH 7.9, 40 mM imidazole, 0.5 M NaCl, 5 mM MgCl(2), 0.5 mM MgADP, 1 mM phenylmethylsulfonyl fluoride). The desired protein was eluted with a 50-ml gradient of 40 mM-0.5 M imidazole in wash buffer. One-ml fractions were collected, and uv-absorbing fractions were pooled. Typical yields were 2-3 mg of protein/liter of cells. Purity was assessed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, which revealed a single band on Coomassie-stained gels. Samples were dialyzed against 20 mM HEPES, 50 mM potassium acetate, 5 mM MgCl(2), 1 mM DTT, 1 mM MgADP, 1 mM NaN(3), pH 7.20, and stored at -70 °C until use. Samples were used within 24 h after thawing.

Sedimentation Velocity Experiments

Sedimentation velocity experiments were conducted on a Beckman Optima XLA analytical ultracentrifuge equipped with absorbance optics and an An60Ti rotor as described (Correia et al., 1995). These experiments were done at 42,000 rpm, 24.6 °C in charcoal-filled Epon double-sector centerpieces. Velocity data were collected at an appropriate wavelength (232-237 or 280 nm, depending on the initial concentration) and at a spacing of 0.01 cm with four averages in continuous scan mode. The data were analyzed as described (Correia et al., 1995).

Preparation of Tubulin and Microtubules

Bovine brain microtubules were prepared by two cycles of temperature-dependent polymerization and depolymerization (Shelanski et al., 1973). Tubulin was purified further by phosphocellulose chromatography, as described (Weingarten et al., 1975; Sloboda et al., 1976). Aliquots of tubulin which were frozen at -80° C were thawed. PIPES was added to 100 mM, and MgSO(4) was added to 5 mM. GTP (from a frozen stock of 50 mM in water, pH 7.0) was added to 1 mM. The tubulin was allowed to polymerize at 35 °C for 20 min. Taxol was added to 100 µM, and polymerization was allowed to continue for 15 min. The sample was spun in a microcentrifuge for 30 min at 4 °C. The supernatant was discarded and the pellet resuspended in ATPase buffer (20 mM HEPES, 5 mM MgCl(2), 50 mM potassium acetate, 1 mM DTT, 1 mM NaN(3), pH 7.2) plus equimolar taxol. The suspension was left at 35 °C for 30 min and then centrifuged for 30 min at room temperature. The supernatant was discarded and the pellet resuspended as above. This procedure was repeated one more time to ensure that no free phosphate remained in the sample. The concentration was determined using Bio-Rad protein assay dye reagent.

ATPase Assays

A modification of the malachite green method (Kodama et al., 1986) was used for the detection of inorganic phosphate in ATPase assays. A solution containing K332 or K413 and varying concentrations of microtubules in ATPase buffer was combined for a total volume of 225 µl and allowed to sit in a 25 °C water bath for 1 h. Samples were assayed for ATPase activity. 2.25 µl of 50 mM MgATP in water was added, and the reaction was allowed to proceed for the desired length of time. The reaction was stopped by adding 225 µl of ice-cold 0.6 M perchloric acid. Samples were spun in a microcentrifuge for 4 min to pellet the precipitated protein. 400 µl of supernatant was transferred to a tube containing 400 µl of color development reagent (0.2% sodium molybdate, 0.03% malachite green oxalate, and 0.05% Triton X-100 in 0.7 M HCl) at 30-s intervals. Each sample's absorbance at 650 nm was read exactly 10 min after the addition of the color development reagent. The amount of inorganic phosphate was determined from a standard curve, and the ATPase rates were calculated.

Data for K332 ATPase rate versus microtubule concentration were fit to the Michaelis-Menten equation to determine k and K. For K413, rate versus microtubule concentration data had to be corrected for the percentage of K413 which was monomeric, by fitting the ATPase rate, , to

where is the fraction of K413 which is monomeric, 1 - is the fraction which is dimeric, and M is the concentration of tubulin dimer. The value of was determined by equilibrium sedimentation studies of K413 which will be reported elsewhere. (^2)The values of K and k for monomeric K413 were determined by measuring microtubule-activated ATPase activity at a K413 concentration of 50 nM, where the construct is >95% monomeric.^2

Mant ADP, Mant AMPPNP Binding Assays

Binding of mant ADP and mant AMPPNP to K413 was determined by a dialysis method. K413 in ATPase buffer was made nucleotide-free as described previously (Sadhu and Taylor, 1992) by the addition of EDTA to 10 mM, incubation at room temperature for 15 min, followed by gel filtration on Sephadex G-25 (PD-10, Pharmacia) which had been equilibrated in 20 mM HEPES, 50 mM potassium acetate, 0.1 mM EDTA, 1 mM DTT, 1 mM NaN(3), pH 7.20. Magnesium chloride was immediately added to 5 mM, and mant AMPPNP or mant ADP was added over a range of concentrations to samples of K413 at a concentration of 2 µM. Samples were briefly sedimented in Centriprep-10 concentrators, during which time approximately 10% of the volume had filtered through the semipermeable membrane. Equal volumes of filtrate (containing free nucleotide) and retentate (containing total nucleotide) were diluted into 2.0 ml of 6 M urea, 25 mM HEPES, 0.1 mM EGTA, pH 7.5, and the fluorescence of the samples was measured in an Aminco SLM 8000 fluorescence spectrophotometer. Fluorescence intensity was converted to concentration of nucleotide by comparing with the fluorescence of a series of samples of defined nucleotide concentration. The standard curve remained linear throughout the range of concentrations examined. Free and bound nucleotide concentrations were calculated, and equilibrium data were calculated by Scatchard analysis, using a plotting program (DeltaGraph 3.5).

The effect of EDTA treatment on nucleotide binding stoichiometry was measured independently for K413 by the following experiment. Twenty µM K413 in ATPase buffer was incubated with a 30-fold molar excess of mant ADP for 4 h at 4 °C. The sample was dialyzed against 1,000 volumes of ATPase buffer, and the remaining mant ADP was removed followed by gel filtration on Sephadex G-25 (PD-10), which had been equilibrated in ATPase buffer. The concentration of K413 was measured by its absorbance at 280 nm, calculated from its amino acid composition ( = 33,650 M cm; Gilbert and Johnson(1993)), and the concentration of mant ADP was measured by its absorbance at 356 nm ( = 5,800 M cm; Hiratsuka(1983)).

^3H Labeling of Kinesin

10 µCi of N-succinimidyl [2,3-^3H]propionate (80 Ci/mmol) was spotted on a wedge of filter paper and air dried in an Eppendorf tube. 500 µl of K413 or K332 in 20 mM HEPES, 5 mM MgCl(2), 50 mM potassium acetate, 0.1 mM EGTA, 1 mM NaN(3), pH 7.20, was added to the label and incubated at 4 °C for 4 h. Unreacted label was removed by gel filtration of the protein on Sephadex G-25 (PD-10). Labeling stoichiometry, determined from the specific activity and protein concentration, was less than 1 mol of label/1,000 mol of protein.

Microtubule Binding Assay

^3H-Labeled kinesin at a concentration of 50-250 nM was combined with varying concentrations of a >15-fold excess of microtubules in a final volume of 220 µl in 20 mM HEPES, 5 mM MgCl(2), 50 mM potassium acetate, 1 mM NaN(3), pH 7.20. For studies of binding in the presence of ADP, 1 mM MgADP was added to the sample mixture. Samples were equilibrated at 25 °C for 60 min in the presence of 0.5% bovine serum albumin, loaded into Beckman Airfuge centrifuge tubes, and sedimented at 125,000 times g for 20 min. 190 µl of supernatant was placed in 3.0 ml of scintillation fluid (Opti-fluor, Packard) and counted. Counts were normalized by comparing with kinesin samples sedimented in the absence of microtubules. Sedimentation of labeled K413 or K332 in the absence of microtubules reduced the concentration of kinesin in the supernatant by less than 5%.

Binding data for K332 could be fit to a binding isotherm of the form

where is the fractional binding, defined as the ratio of sedimented K332 to total K332, (max) is the maximum degree of binding, K(a) is the association constant, and [M] is microtubule concentration. For K413bulletADP ± beryllium fluoride, fitting required a correction for the fraction of K413 which is dimeric versus monomeric. This was accomplished by utilizing data from equilibrium sedimentation studies.^2 Data were fit to , which assumes: 1) that the fraction of K413 that is monomeric () binds to the microtubule with affinity K(a), determined by measuring microtubule binding affinity at low K413 concentration, where the construct is monomeric; 2) that the fraction of K413 which is dimeric is 1 - ; 3) that each head of a free dimeric K413 molecule can bind to the microtubule affinity constant K(1); and 4) that binding of the second head to the microtubule occurs with affinity constant K(2). , the fractional binding of K413, is defined as the ratio of unsedimented K413 to total K413. Under these conditions, the degree of binding, , is determined by the fractional binding of the monomeric and dimeric species.

Expressing these quantities in terms of the various affinity constants reveals the following (Tanford, 1961).

The validity of using values of in which were derived from equilibrium sedimentation studies was tested by setting as an independent variable in fitting to for the data in Fig. 3A. The value of derived by this method for K413bulletADP at a concentration of 250 nM ( = 0.62, r^2 = 0.96) was very close to that derived from equilibrium studies ( = 0.60; Footnote 2). Free energies of binding for dimeric K413 were calculated using the microscopic binding constant, K, whose relationship to the individual macroscopic binding constants, K(i),in is as follows (Tanford, 1961)


Figure 3: Fractional binding versus tubulin dimer concentration for kinesin constructs. Panel A, K413 at 250 nM (closed squares), 50 nM (open triangles), and 250 nM K413 + 1 mM BeSO(4) + 5 mM NaF (closed circles). Data for K413 at 50 nM fit a rectangular hyperbola, which defines a value of K of 7.8 µM and stoichiometry of 0.9. Conditions: 20 mM HEPES, 50 mM potassium acetate, 5 mM MgCl(2), 1 mM DTT, 1 mM NaN(3), 1 mM MgADP, pH 7.20, 25 °C. Data for K413bulletADP ± beryllium fluoride at 250 nM were fit to (see ``Experimental Procedures''), which corrects for the reversible dimerization of this construct. Values of were determined from sedimentation equilibrium studies (Footnote 2). This reveals 1/K(1) = 0.5 µM and 1/K(2) = 7.1 µM for dimeric K413bulletADP (stoichiometry 0.8) and 1/K(1) = 0.6 µM and 1/K(2) = 0.4 µM for dimeric K413bulletADP + beryllium fluoride (stoichiometry 0.8). Panel B, 100-200 nM K332 (closed squares), K332 + 1 mM BeSO(4) + 5 mM NaF (closed circles), K332 + 1 mM AlNO(3) + 5 mM NaF (open triangles), K332 + 1 mM AMPPNP (closed diamonds), and K332 + 10 mM sodium phosphate (open squares). Conditions as in panel A. Data for each sample could be adequately fit to a rectangular hyperbola, defining values of dissociation constants as follows: K332bulletADP, 20.8 µM (stoichiometry 0.9); K332bulletADPbulletBeF, 5.6 µM (stoichiometry 0.8); K332bulletADPbulletAlF, 1.4 µM (stoichiometry 0.8); K332bulletAMPPNP, 1.1 µM (stoichiometry 0.8); K332bulletADPbulletP(i), 18.2 µM (stoichiometry 0.8).



where n, the number of microtubule binding sites on K413, is equal to 2. Thus, a prediction of the model described by is that the individual values of K(i) are highly correlated with each other. In the presence of cooperativity, K(2) may be larger (positive cooperativity) or smaller (negative cooperativity) than expected from that predicted by the value of K(1). In the presence of cooperativity, the free energy of binding of the second site to a microtubule would be (Tanford, 1961)

where DeltaG is the intrinsic free energy of binding in the absence of cooperativity, and DeltaG^0() is the interaction free energy. The free energy of binding to the second site on K413 was derived by using the value of the microscopic constant, K (derived from K(1)) to determine DeltaG^0 and by using the difference between the predicted value of K(2) (determined from K(1) by assuming no interactions) and that determined from .

Effect of Inorganic Phosphate on KinesinbulletADPbulletMicrotubule Affinity

The effect of inorganic phosphate on the affinity of K413bulletADP for microtubules was measured using the microtubule binding assay as described above. ^3H-Labeled K413 in the presence of 1 mM ADP was mixed with a range of microtubule concentrations, and the binding affinity was measured as a function of increasing phosphate concentration. The concentration of potassium acetate was varied to keep the ionic strength constant. Data were fit to both and , and best fitting was found with , which assumes that only one of the two heads can attach to the microtubule. Apparent dissociation constants (K(d)) were calculated from these fits. The relationship between K(d) and phosphate concentration was determined from a four-state equilibrium model as follows.

which assumes that only one head of the K413 dimer can attach to the microtubule when phosphate occupies the active site. In this scheme, K is kinesinbulletADP, MK is kinesinbulletADP bound to microtubules, P(i) is inorganic phosphate, and K are association constants for each step. The apparent dissociation constant, K(d), at a given phosphate concentration, [P(i)], is defined by

which yields the following.

Statistical Methods

Initial values for the desired parameters from and 8 were derived from a least squares curve fitting program (DeltaGraph Pro3), yielding fits with r^2 values >0.95. Fitting was then refined by using a multivariate algorithm from a commercially available statistics package (PROC NLIN; SAS 6.10) to generate parameters ± 1 S.D.


RESULTS

Characterization of Human Kinesin Constructs

Previous studies of recombinant kinesin from a Drosophila clone had demonstrated that constructs containing the amino-terminal motor domain could be dimeric or monomeric (Huang and Hackney, 1994; Huang et al., 1994; Correia et al., 1995; Lockhart et al., 1995; Young et al., 1995), depending on how much of the alpha-helical tail was included in the construct. Since the human clone of kinesin used in these studies is highly homologous to that from Drosophila, it seemed likely that homologous constructs from this clone would behave similarly. Two constructs were generated from the human clone: K413, containing the first 413 residues from the amino terminus and corresponding to residues 1-401 in the Drosophila sequence; and K332, containing the first 332 residues from the amino terminus and corresponding to residues 1-340 in the Drosophila sequence. Sedimentation velocity studies of K413 at concentrations geq 2 µM reveal an s(w) value of 4.86. This is very similar to the corresponding value for the Drosophila 1-401 construct, which behaves largely as a dimer (Correia et al., 1995). Equilibrium sedimentation studies^2 establish that at 24.6 °C, the dissociation constant for dimerization of K413bulletADP is 0.738 ± 0.043 µM. Studies with K413bulletADPbulletBeF, K413bulletADPbulletAlF, and K413bulletADPbulletP(i) yield very similar values.^2 Thus, over the concentration range used in this study (50-500 nM), the fraction of K413 which is dimeric varies from <5% to 50%. By contrast, sedimentation velocity studies of K332 reveal an s(w) value of 3.32, which, although slightly larger than the corresponding value of the Drosophila homologue (Correia et al., 1995), is within the range of constructs which in that study behaved as monomers below 4 µM. The monomeric nature of K332 is also supported by its elution behavior on Sephacryl S-300 and on Superose 12 FPLC (data not shown), as well as by equilibrium sedimentation studies of this construct to be reported elsewhere.^2

Titration of the active site of K413 was accomplished by measuring the equilibrium constant for binding of the fluorescent nucleotide derivatives mant ADP and mant AMPPNP, using an equilibrium dialysis technique. Scatchard plot analysis of the data (Fig. 1) demonstrates that K413 binds mant ADP with one class of binding site, characterized by a dissociation constant of 0.67 µM and stoichiometry of 0.80 mol of mant ADP/mol of head. By contrast, the dissociation constant of mant AMPPNP, at 7.1 µM, is approximately 10-fold greater, and the stoichiometry is 1.3 mol/mol of head. The effect of EDTA treatment on K413 was investigated by examining the stoichiometry of mant ADP binding under conditions where it was present in large molar excess over K413 active sites, in the absence of EDTA. This revealed a range of stoichiometries of 0.87-0.95 over five separate measurements (data not shown). Thus, EDTA treatment does appear to reduce the binding stoichiometry by approximately 8-19%. This percentage reduction in the binding stoichiometry after EDTA treatment is very similar to that measured previously, using [^3H]ATP binding (Ma and Taylor, 1995b).


Figure 1: Binding of mant AMPPNP (closed circles) and mant ADP (closed boxes) to nucleotide-free K413. K413 was made nucleotide-free as described (Sadhu and Taylor, 1992) and mixed with a range of concentration of mant nucleotide. Free nucleotide was measured by an equilibrium dialysis method. Scatchard analysis of the data reveals that binding of mant ADP to K413 is characterized by a dissociation constant of 0.67 µM and a stoichiometry of 0.80; for mant AMPPNP, the corresponding values are 7.1 µM and 1.30.



Steady-state Kinetics of the Kinesin ATPase

The steady-state ATPase kinetics of K413 and K332 were examined in the presence of 0.5 mM ATP, which is a concentration that is approximately 25-35-fold greater than the K for the corresponding Drosophila constructs. The steady-state ATPase rates for K413 and K332 at this ATP concentration were 0.003 and 0.007 s. The values of k for the corresponding Drosophila constructs are 0.01 and 0.029 s, respectively (Gilbert and Johnson, 1993; Huang and Hackney, 1994).

As in the case of the Drosophila constructs, the K413 and K332 MgATPase activities are markedly activated by microtubules. At 50 nM K413, the construct is essentially entirely monomeric, and data in Fig. 2A fit conventional Michaelis-Menten kinetics, defining values of K and k of 3.5 µM and 35.1 s, respectively. Increasing the K413 concentration to 300 nM generated data that needed to be fit to . This defined values of K and k for dimeric K413 of 0.54 µM and 8.25 s. Fig. 2B demonstrates the corresponding data for K332, which defines values of K and k of 26.0 µM and 43.6 s.


Figure 2: ATPase rate versus tubulin dimer concentration for kinesin constructs. Panel A, K413 at 300 nM (open squares), 50 nM (open triangles), and 300 nM in the presence of 1 mM BeSO(4) + 5 mM NaF (open circles). Data for K413 at 50 nM fit Michaelis-Menten kinetics with k = 35.1 s and K = 3.5 µM. Data for K413 ± beryllium fluoride at 300 nM were fit to (see ``Experimental Procedures''), which corrects for reversible dimerization of this construct. Values of were determined from sedimentation equilibrium studies (Footnote 2). This reveals k = 8.25 s and K = 0.54 µM for dimeric K413 and k = 0.83 s and K = 0.38 µM for dimeric K413 + beryllium fluoride. Panel B, corresponding data for K332 in the absence (closed diamonds) or presence of 1 mM AlNO(3) + 5 mM NaF (open squares), 1 mM BeSO(4) + 5 mM NaF (open circles), and 0.5 mM sodium vanadate (open triangles). Fitting to Michaelis-Menten kinetics reveals the following: k = 43.6 s and K = 26.0 µM for K332; k = 7.2 s and K = 4.0 µM for K332 + beryllium fluoride; k = 0.08 s and K = 3.7 µM for K332 + aluminum fluoride; and k = 2.9 s and K = 5.8 µM for K332 + sodium vanadate. Panel C, double-reciprocal plot of data from panel B for K332 in the absence (open squares) and presence (open circles) of 1 mM BeSO(4) + 5 mM NaF. This reveals parallel curves, as expected for uncompetitive inhibition, and defines a value of K of 41 µM.



The kinetics of the microtubule-activated ATPase activity showed a marked ionic strength dependence. Raising the ionic strength from 50 mM potassium acetate to 150 mM KCl raised the value of K for K413 to 80 µM and decreased the value of k to 4 s (data not shown). Corresponding data for K332 were 97 µM and 10 s (data not shown).

Dissociation Constant of K413 and K332bulletADP for Microtubules

Previous kinetic studies on intact brain kinesin and on truncated constructs (Hackney, 1988; Hackney et al., 1989; Sadhu and Taylor, 1992; Gilbert and Johnson, 1994) indicated that the rate-limiting step in the microtubule kinesin ATPase cycle was ADP release, implying that kinesinbulletADP would be the predominant species under steady-state conditions. To test this with the human construct, the affinities of tritium-labeled K413 and K332 for microtubules were measured in the presence of 1 mM MgADP, using an Airfuge assay. Previous studies with a similar human kinesin construct have shown that labeling with N-succinimidyl [2,3-^3H]propionate has no effect on the microtubule-activated ATPase or on the kinetics of mant ATP binding (Ma and Taylor, 1995a). At a concentration of 50 nM, K413 is >95% monomeric.^2 Binding to microtubules in the presence of ADP and at this concentration could be described by a hyperbolic binding isotherm, defining a value of K(d) of 7.8 µM and a maximum degree of binding of 0.80 ± 0.09 (Table 1). Increasing the concentration of K413 to 250 nM increases the fraction of dimer to 40% and enhances the relative affinity. Fitting to reveals values of 1/K(1) and 1/K(2) of 0.5 and 7.1 µM, respectively (Table 1). Data for K332 could be fit to a hyperbolic binding isotherm and are depicted in Fig. 3B, which reveals a value of K(d) of 20.8 µM and a maximum degree of binding of 0.78 ± 0.05. For K332, values of K(d) are very similar to the corresponding values of K discussed above. As in the case of K, binding affinity showed a strong ionic strength dependence, increasing to a value of 139 µM for K332 at 150 mM KCl (data not shown).



Complex Formation with Beryllium Fluoride and Aluminum Fluoride

Beryllium fluoride and aluminum fluoride are known to form ternary complexes with myosin, actin, and other NTPases when nucleotide is bound in the active site (Goodno, 1979; Combeau and Carlier, 1988, 1989; Maruta et al., 1993; Phan and Reisler, 1992; Phan et al., 1993). These transition metals act as uncompetitive inhibitors of the myosin ATPase by virtue of their tight binding to the myosinbulletADP state. Recent crystallographic data suggest that myosinbulletADPbulletBeF(n) mimics the myosinbulletATP state, whereas myosinbulletADPbulletAlF(4)andmyosinbullet ADPbulletVnmimicaprehydrolyticmyosin-nucleotidetransitionstate (Fisheret al., 1994). To establish if these metals had similar effects on the kinesin ATPase, K413 was incubated in the presence of 30 µM MgADP with 1 mM BeSO(4) + 5 mM NaF for 10 min, 1 h, 2 h, 4 h, and 6 h. Taxol-stabilized microtubules were then added to 20 µM, and MgATP was added to 0.5 mM to initiate the ATPase reaction. In the absence of added beryllium fluoride, the ATPase rate at this microtubule concentration is 11.1 s. Within 10 min of incubation, this had decreased to 1.12 s, and by 1 h of incubation, had decreased further to 0.56 s. More prolonged incubation did not reduce the ATPase any further, and this result indicates therefore that complex formation is complete within 1 h of incubation.

Beryllium and aluminum fluoride, like vanadate, reduce the affinity of myosinbulletADP for actin, since myosinbulletADPbulletberyllium fluoride and myosinbulletADPbulletaluminum fluoride mimic the weakly bound prehydrolytic myosin states (Fisher et al., 1994). The value of the actin-activated k is reduced in these complexes because of the tight binding of the metals to the active site (Maruta et al., 1993). Thus, if kinesin follows the same behavior, it would be expected that beryllium fluoride and aluminum fluoride should decrease the value of k and increase the value of K for the microtubule-activated kinesin ATPase. As Fig. 2B demonstrates, both transition metals reduce not only k but also K for K332, implying that the kinesinbullet ADPbulletBeF and kinesinbulletADPbulletAlF states are more strongly bound than kinesinbulletADP. Experiments performed in the presence of 0.5 mM sodium vanadate also demonstrate that this transition metal reduces both k and K (Fig. 2B and Table 1). Corresponding data for K413 in the presence of beryllium fluoride are shown in Fig. 2A. A double-reciprocal plot of the microtubule-activated K332 ATPase rate versus microtubule concentration is shown in Fig. 2C for samples in the absence and presence of 1 mM BeSO(4) + 5 mM NaF. This demonstrates essentially parallel curves, as expected for uncompetitive inhibition, and defines a value of K(i) for beryllium fluoride of 41 µM. Corresponding values of K(i) for aluminum fluoride and vanadate are 1.7 and 35 µM, respectively. Thus, at aluminum and beryllium concentrations of 1 mM, essentially all of the active sites of K413bulletADP and K332bulletADP have transition metal bound.

The decrease in the value of K implies that the transition metals increase the affinity of kinesinbulletADP for microtubules. This was confirmed by using tritium-labeled K413 and K332 in an Airfuge binding assay. Data are summarized in Fig. 3A for K413 at 250 nM and Fig. 3B for K332 and in Table 1. Dissociation constants for K332bulletADP are 5.6 µM in the presence of beryllium fluoride and 1.4 µM in the presence of aluminum fluoride, and both values are close to the corresponding values of K (Table 1). Binding data for K332 in the presence of 1 mM AMPPNP are nearly superimposable on those for aluminum fluoride (Fig. 3B and Table 1). Fitting of binding data for K413bulletADPbulletberyllium fluoride to reveals values of 1/K(1) and 1/K(2) of 0.6 and 0.4 µM, respectively (Table 1).

Effect of Inorganic Phosphate on the Microtubule Affinity of KinesinbulletADP

Inorganic phosphate binds to a number of NTPases, including actin, tubulin, and myosin, with millimolar affinity (Carlier and Pantaloni, 1988; Rickard and Sheterline, 1988; Schlistra et al., 1991; Carlier et al., 1988; Warshaw et al., 1991; Yamakawa and Goldman, 1991; Iwamoto, 1995). In vitro motility studies of actomyosin indicate that phosphate binds to the myosinbulletADP intermediate state with an apparent dissociation constant of 9.5 mM (Warshaw et al., 1991) and inhibits motility by reducing the affinity of myosin for actin. If beryllium fluoride and aluminum fluoride induce formation of a kinesinbulletADPbulletP(i) transition state, then it would be expected that millimolar phosphate should increase the affinity of kinesin for microtubules. Conversely, if phosphate reduces the affinity, this would suggest that these transition metals instead mimic a strong binding state. To test this, tritium-labeled K413 at a concentration of 400 nM was mixed with a range of microtubule concentrations in the presence of 1 mM ADP and increasing phosphate concentrations (1-15 mM). Data are summarized in Fig. 4. Binding data could be fit adequately to a hyperbola, even though nearly 50% of the K413 is dimeric under these conditions. Fitting the values of K(a) into reveals that phosphate binds to K413bulletADP with an apparent dissociation constant of 0.8 mM. The reduction in apparent affinity could be explained by assuming that only one head of the K413bulletADPbulletP(i) dimer can attach to the microtubule and does so with a dissociation constant of 17.0 µM, a value that is nearly identical to that for the monomeric K332bulletADP (Table 1). This result suggests that millimolar concentrations of inorganic phosphate should have little effect on the binding of K332bulletADP to microtubules. This is confirmed by the data in Fig. 3B and Table 1, which reveal that the affinity of K332bulletADP for microtubules is unaffected by the presence of inorganic phosphate at a concentration as high as 10 mM.


Figure 4: Effect of phosphate concentration on K413bulletADP microtubule affinity. Data were fit to (see ``Experimental Procedures'') to determine the dissociation constants of K413bulletADPbulletP(i) for microtubules (1/K(3)) and of phosphate for K413bulletADP (1/K(2)), and this reveals 1/K(3) = 17.0 µM and 1/K(2) = 0.8 mM.




DISCUSSION

This study utilized two constructs derived from a human homologue of the Drosophila kinesin heavy chain. K413, containing residues 1-413, is homologous to Drosophila 1-401, a construct that sedimentation equilibrium studies have revealed is largely dimeric (Correia et al., 1995). Similar conclusions have been reached with Drosophila 1-392 (Huang et al., 1994). The sedimentation velocity behavior of K413 is consistent with its capacity to dimerize, although sedimentation equilibrium studies indicate a dimerization dissociation constant that is approximately 10-fold larger.^2 This larger dimerization constant may reflect intrinsic differences between the human and Drosophila sequences or may be due to the steric and/or ionic effects of the hexahistidine sequence at the extreme carboxyl terminus of the human construct, which is used in affinity purification. By contrast, K332 is homologous to the monomeric Drosophila construct containing residues 1-340 (Huang and Hackney, 1994; Correia et al., 1995), and this is consistent with its sedimentation velocity behavior as well as its elution behavior on Sephacryl S-300 and Superose 12 FPLC.^2 Both K332 and K413 possess an intrinsic MgATPase activity that is very low. For both, microtubules enhance this ATPase activity several thousandfold.

A striking feature of the results from microtubule-activated ATPase studies is that the value of k for monomeric K413 is nearly 5-fold larger than that for dimeric K413 and is only 19% less than that for the monomeric construct K332. This finding resolves an apparent discrepancy between the values of k reported for Drosophila constructs that have been presumed to be dimeric. The value of k for dimeric K413 is close to that reported for experiments with Drosophila K401 at a kinesin concentration of 500 nM (10.9 s; Gilbert and Johnson(1993)). Equilibrium sedimentation studies have shown that at this concentration, approximately 90% of the construct is dimeric (Correia et al., 1995). More recent studies from same laboratory reported a value of k of 20 s (Gilbert et al., 1995). However, in this study, the K401 concentration was lower, at 200 nM. At this concentration, approximately 18-20% of the K401 would be monomeric (Correia et al., 1995). predicts that an overall rate of 20 s could be observed if k for dimeric K401 is 10.9 s, and k for monomeric K401 is approximately 55 s, which is consistent with our own results. Other experiments on a Drosophila construct that is 9 residues shorter measured a value of k of 47 s, which is very similar to that for K332 and only 1.3-fold larger than the value of k for monomeric K413 (Hackney, 1994a). In this study, the concentration of the kinesin construct used in the ATPase assay was 8.4 nM. It is reasonable to assume that the association constant for dimerization of this construct should be similar to or perhaps less than that for Drosophila K401, and given this, the published data would indicate that this construct would be >75% monomeric (Correia et al., 1995). Thus, our data clearly indicate that dimerization of kinesin markedly inhibits the degree of microtubule activation.

An increase in enzymatic activity with increasing degrees of truncation has been reported not only for kinesin, but also for other microtubule motors, such as ncd (Stewart et al., 1993; Huang and Hackney, 1994; Huang et al., 1994; Chandra et al., 1993). One possible explanation for this inhibition is that the conformational changes that occur during the power stroke of the kinesin ATPase cycle may be linked to a rate-limiting rotation of one head relative to the other, which would be mediated through the dimerization segment contained within the carboxyl-terminal 81 residues of K413 (Fig. 5). Loss of dimerization, either by dilution of K413 or by truncation of the dimerization segment, would lower the energy barrier for this conformational change and accelerate this rate-limiting step.


Figure 5: Structural model of the kinesin:microtubule mechanochemical cycle. Panel A, model of the K332bulletmicrotubule interactions. The tubulin dimer is depicted as the arrow-shaped structure, and its binding site for kinesin is depicted as the vertically shaded box. The kinesin monomer is depicted as the comma-shaped structure. In the presence of ADP ± P(i) (MKD/MKDbulletP), kinesin assumes a weak binding conformation, symbolized by the rounded shape of the microtubule-associated end of the molecule. When ATP occupies the active site (MKT), kinesin assumes a strong binding conformation, symbolized by the squared-off shape of the microtubule-associated end of the molecule. Panel B, model of the K413bulletmicrotubule interactions. In this figure, the two kinesin heads are shaded differently to distinguish them. In addition, the dimerization segment, containing portions of the alpha-helical tail segments of each monomer, is shaded separately. In the absence of nucleotide, or when ATP occupies the active site (MKT, MK), binding to the microtubule demonstrates positive cooperativity, presumably because of a stabilizing of the orientation of the second kinesin head (cross-hatched) by attachment of the first head (dotted) to the microtubule binding site (vertically shaded box). Hydrolysis of ATP to ADP+P(i) (MKDbulletP) leads to a weakening of the affinity of the cross-hatched head of the kinesin dimer. This is postulated to lead to a change in flexibility of the dimerization segment, which allows a rotation of the second (dotted) head and prevents this head from attaching to the microtubule. Dissociation of phosphate (MKD) allows the unattached (dotted) head to reattach weakly to the next available downstream binding site. This is presumed to stabilize the interaction of the other (cross-hatched) head, whose microtubule affinity is thereby enhanced. Dissociation of ADP from the active sites enhances the affinity of the attached heads to that seen in the presence of ATP and regenerates a fully attached state.



Active site titrations with mant ADP and mant AMPPNP give the expected result of one class of binding site with an approximately 1:1 stoichiometry. The affinity of mant ADP for K413, determined in this study by equilibrium methods, is approximately 2 orders of magnitude lower than that measured with the complete heavy chain-light chain complex (Hackney, 1988; Sadhu and Taylor, 1992). The large difference in affinities between intact kinesin and the truncated constructs may reflect the trapping of nucleotide in the active site of intact kinesin when it assumes the folded conformation (see above; Hackney(1992)). In contrast to myosin, which binds AMPPNP with higher affinity than ADP (Trybus and Taylor, 1982), kinesin binds with the reverse order of affinities: the dissociation constant of mant AMPPNP binding to K413 is approximately 10 times larger than that for ADP (Fig. 1). This argues that kinesin interacts with nucleotide in a manner distinct from that for myosin.

The results of microtubule binding studies with the monomeric construct K332 will be discussed first, as their interpretation is more straightforward. Binding studies in the presence of 1 mM MgADP yielded dissociation constants that are close to the corresponding values of K. The relationship between the midpoint of the ATPase titration and the dissociation constant was explicitly developed in a previous study of the actosubfragment 1 ATPase mechanism (Rosenfeld and Taylor, 1984), and the four-state model derived in that study can be applied to the case of kinesin as well. This model, as applied to kinesin, assumes that the kinesinbulletATP and kinesinbulletADPbulletP(i) states are in rapid equilibrium with microtubules, an assumption that is supported by recent studies (Gilbert et al., 1995). According to this model, the relation between steady-state velocity and microtubule concentration is

where k and k are the weighted forward and reverse rate constants, respectively, for the nucleotide hydrolysis step, and K(a) is the association constant of kinesinbulletADPbulletP(i) for the microtubule. For both kinesin and the microtubule-kinesin complex, k k (Ma and Taylor, 1995a, 1995b; Gilbert and Johnson, 1994), which leads to

where K = 1/K(a). Since the model predicts that the intrinsic affinities of kinesinbulletADP and kinesinbulletADPbulletP(i) are essentially equal, this leads to the prediction that for K332bulletADP, K K(d), which is supported by the data in Table 1.

Experiments with K332bulletADP + beryllium fluoride and aluminum fluoride support the contention that kinesin interacts with nucleotide in a manner distinct from that for myosin. Both transition metals were found to enhance the affinity of K332 for microtubules. In the case of aluminum fluoride, the enhancement in affinity was the same as that seen with AMPPNP. This contrasts markedly with the case of myosins, where these transition metals, as well as inorganic phosphate, reduce the affinity of myosin for actin, as expected with the assignment of the myosinbulletATP and myosinbulletADPbulletP(i) states as weak binding (Goodno and Taylor, 1982; Yamakawa and Goldman, 1991; Phan and Reisler, 1992). Previous studies of kinesin with aluminum fluoride and ATPS in an in vitro motility assay suggested that kinesin has reversed the cycle, that kinesinbulletADP is weak binding and kinesinbulletATP and kinesinbulletADPbulletP(i) are strong binding (Romberg and Vale, 1993). Interpretation of these results and of our own, however, depends on an unambiguous assignment of the conformation for kinesinbulletADPbulletaluminum fluoride and kinesinbulletADPbulletberyllium fluoride. The results of our study indicate that aluminum and beryllium fluoride mimic prehydrolytic kinesin-nucleotide states, which is supported by crystallographic data from myosin (Fisher et al., 1994) and that these states are strongly binding, the opposite of what is seen in myosin.

Binding data for K332 can be interpreted by a relatively simple model, in which monomeric kinesin can attach to the tubulin dimer in one of two conformations: a strong binding state that releases approximately -8.0 kcal/mol of binding free energy, and a weak binding state that releases approximately -6.4 kcal/mol (Fig. 5A). The data in Table 1and Fig. 3B can then be explained by assuming that K332bulletADP assumes a weak conformation, whereas K332bulletAMPPNP and K332bulletADPbullet aluminum fluoride assume the strong conformation.

Interpretation of the binding studies with K413 is more complicated because of the reversible dimerization that occurs with this species^2 and the need to account for one-site versus two-site binding of the dimer. The model defined by gave a superior fit to the data (^2 = 0.011 for K413bulletADP and K413bulletADPbulletBeF) than that for a simple hyperbolic isotherm (^2 = 0.10). However, initial attempts to fit the data to led to large standard errors for 1/K(1) and 1/K(2) because of the high degree of correlation between these parameters. This is consistent with the model described by , since the values of K(1) and K(2) are related to each other (see ``Experimental Procedures''; Tanford(1961)). Improved fitting required use of the Moore-Penrose inverse and led to values of K(1) and K(2) which had acceptable errors. It is proposed that each head of the K413 dimer can assume a strong or a weak conformation. Furthermore, it is presumed that monomeric K413bulletADP, like K332bulletADP, attaches to the microtubule via a weak interaction only. We propose that each head of K413bulletATP, mimicked by K413bulletADP + beryllium fluoride or aluminum fluoride, is in a strong conformation. If these two heads behave in an identical and independent manner, then K(2) = (K(1))/4 (Tanford, 1961). Fitting of binding data, however, demonstrates that K(2) = 1.5(K(1)). This implies that binding of the second head of the K413bulletATP dimer demonstrates positive cooperativity and releases an additional -1.1 kcal/mol of interaction-free energy. Although the structural basis for this cooperative interaction was not determined from this study, one possibility is that the binding of the first head stabilizes the orientation of the second head so that its binding is more favorable. Hydrolysis of ATP to ADP+P(i) is proposed to do two things: it alters the conformation of both heads (to weak binding) as well as their relative orientation, making it sterically impossible for the second head to attach to the microtubule at all. Release of phosphate, to generate a kinesinbulletADP dimer, would allow the second head to reattach. Results of fitting to indicate that for K413bulletADP, K(2) = (K(1))/14.2, which is nearly four times smaller than predicted from a noninteracting system. These data can be interpreted to mean that phosphate release has no effect on the intrinsic microtubule affinity of kinesinbulletADP; rather, that weak attachment of the second head (dotted in Fig. 5B) stabilizes the interaction of the first head (cross-hatched in Fig. 5B) with the microtubule and enhances its affinity. This model thus predicts that in the absence of a second head, binding of kinesinbulletADP should be weak and that phosphate should have no effect. The data with K332 and monomeric K413 support this ( Table 1and Fig. 5A). Furthermore, it is proposed that the strong reattachment of one of the K413bulletADP heads to the microtubule generates force and accelerates release of its bound nucleotide. This is consistent with the finding that K K(1) for K413bulletADP and explains why microtubules accelerate the release of bound ADP from only one head of dimeric kinesin constructs (Hackney, 1994b).

This model is also consistent with the relative affinities of ADP versus AMPPNP for K413 (Fig. 1). Differences in the energetics of microtubule binding between K413bulletATP (mimicked by K413bulletAMPPNP or K413bulletADPbulletberyllium fluoride) and K413bulletADP should be reflected in reciprocal differences in the energetics of binding of AMPPNP versus ADP if the conformational changes between K413bulletADP and K413bulletATP are driven by nucleotide binding. Calculation from the nucleotide binding data reveals a difference of 2.8 kcal/mol, which is close to the value calculated from microtubule affinities, of 2.3 kcal/mol. Finally, the effect of phosphate on K413bulletADP is not due to a destabilizing of microtubules, as some studies have shown that phosphate enhances microtubule stability, with an apparent dissociation constant of 25 mM (Carlier et al., 1988; Schlistra et al., 1991), whereas others see no effect of inorganic phosphate over this concentration range (Caplow et al., 1989).

These results therefore predict that dissociation of the microtubule-kinesin complex during the ATPase cycle should occur upon formation of the weakly bound kinesinbulletADPbulletP(i) complex, as suggested by one of two models proposed on the basis of kinetic studies (Gilbert et al., 1995). This study provided a kinetic explanation for processivity through the relatively slow rate of dissociation and the rapid rate of rebinding of the weakly bound kinesin intermediate. Our results demonstrate that the most weakly bound K413 intermediate (K413bulletADPbulletP(i)) has a dissociation constant for microtubules which is still significantly lower than the local concentration of tubulin intracellularly (Gilbert et al., 1995). Thus, our data indicate that at least one head of the kinesin dimer would likely remain attached to the microtubule at all times in the ATPase cycle. It also predicts that there is a large free energy change associated with hydrolysis of ATP in the kinesin-microtubule complex. This free energy change presumably drives the power stroke and if so, should be reflected in a significant conformational change in the kinesin molecule. As a subsequent study will demonstrate, this can be detected through fluorescent anisotropy decay studies on labeled kinesin preparations.^2

The model presented here is also consistent with several recent structural models of the kinesin-microtubule and ncd-microtubule complex. Hydrolysis of ATP would lead to dissociation of one head and pivoting of the attached head toward the adjacent tubulin subunit. This would enable the unattached head of the kinesin dimer, presumably containing ADP, to attach to the next site along the microtubule protofilament. Since the kinesin head is smaller than the tubulin heterodimer, attachment of the second head could only occur in one direction because of simple steric considerations. This model is similar to those proposed previously (Hirose et al., 1995; Hackney, 1994b), except that it assigns ATP hydrolysis and the production of kinesinbulletADPbulletP(i) as the step that leads to detachment and pivoting of the kinesin molecule. Directionality in this model could be explained by assuming that the more weakly attached of the two kinesinbulletATP heads is the one that detaches after hydrolysis and that for kinesin, this is the head which is closer to the(-) end. Thus, directionality becomes an intrinsic consequence of the polarity of the microtubule lattice. Reversal of directionality, as is seen in ncd, could then simply result from reversing the affinities of the two heads in the ATP-bound state.


FOOTNOTES

*
This work was supported by Grants NS01500 and NS31096 from the NINDS, National Institutes of Health (to S. S. R.), by Grant AR31239 from the NIAMS, National Institutes of Health (to H. C. C.), and by Grant HSF BIR9216150 (to J. J. C.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Dept. of Neurology, University of Alabama at Birmingham, UAB Station, 510 Medical Education Bldg., Birmingham, AL 35294-0007.

(^1)
The abbreviations used are: ATPS, adenosine 5`-O-(thiotriphosphate); AMPPNP, adenosine 5`-(beta,-imidotriphosphate); mant ADP, N-methylanthraniloyl ADP; DTT, dithiothreitol; K332, bacterially expressed human kinesin construct containing residues 1-332; K413, bacterially expressed human kinesin construct containing residues 1-413; PIPES, 1,4-piperazinediethanesulfonic acid.

(^2)
S. S. Rosenfeld, B. Rener, J. J. Correia, M. S. Mayo, and H. C. Cheung, manuscript in preparation.


ACKNOWLEDGEMENTS

We thank Dr. Ron Vale (University of California, San Francisco) for the generous gift of a clone of human kinesin, and Sylvia and David MacPherson of the Protein Expression Core Facility of the UAB AIDS Center (supported by Grant P30 AI27767 from the National Institutes of Health) for help in the production of the K413 and K332 constructs. We also thank Dr. J. N. Whitaker (Department of Neurology, UAB) for continued support.


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