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• W2000315199 abstract "A variety of models have recently emerged to explain how the molecular motor kinesin is able to maintain processive movement for over 100 steps. Although these models differ in significant features, they all predict that kinesin's catalytic domains intermittently separate from each other as the motor takes 8-nm steps along the microtubule. Furthermore, at some point in this process, one molecule of ATP is hydrolyzed per step. However, exactly when hydrolysis and product release occur in relation to this forward step have not been established. Furthermore, the rate at which this separation occurs as well as the speed of motor stepping onto and release from the microtubule have not been measured. In the absence of this information, it is difficult to critically evaluate competing models of kinesin function. We have addressed this issue by developing spectroscopic probes whose fluorescence is sensitive to motor-motor separation or microtubule binding. The kinetics of these fluorescence changes allow us to directly measure how fast kinesin steps onto and releases from the microtubule and provide insight into how processive movement is maintained by this motor. A variety of models have recently emerged to explain how the molecular motor kinesin is able to maintain processive movement for over 100 steps. Although these models differ in significant features, they all predict that kinesin's catalytic domains intermittently separate from each other as the motor takes 8-nm steps along the microtubule. Furthermore, at some point in this process, one molecule of ATP is hydrolyzed per step. However, exactly when hydrolysis and product release occur in relation to this forward step have not been established. Furthermore, the rate at which this separation occurs as well as the speed of motor stepping onto and release from the microtubule have not been measured. In the absence of this information, it is difficult to critically evaluate competing models of kinesin function. We have addressed this issue by developing spectroscopic probes whose fluorescence is sensitive to motor-motor separation or microtubule binding. The kinetics of these fluorescence changes allow us to directly measure how fast kinesin steps onto and releases from the microtubule and provide insight into how processive movement is maintained by this motor. cysteine-light mutant 5′-adenylyl-β,γ-imidodiphosphate 2′-deoxy mant-ATP fluorescence resonance energy transfer 5-IAEDANS TMR, tetramethyl rhodamine maleimide Several features of kinesin's mechanochemistry contribute to its ability to move long distances on the microtubule without dissociating. These include its high duty ratio and ability to hydrolyze multiple ATP molecules per force productive encounter-features that enhance the probability that a motor will remain attached even under load (1Hancock W.O. Howard J. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 13147-13152Crossref PubMed Scopus (185) Google Scholar, 2Ma Y.-Z. Taylor E.W. J. Biol. Chem. 1997; 272: 717-723Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar, 3Vale R.D. Milligan R.A. Science. 2000; 288: 88-95Crossref PubMed Scopus (1247) Google Scholar, 4Jiang W Stock M.F., Li, X. Hackney D.D. J. Biol. Chem. 1997; 272: 7626-7632Abstract Full Text Full Text PDF PubMed Scopus (75) Google Scholar, 5Schnitzer M.J. Visscher K. Block S.M. Nat. Cell Biol. 2000; 2: 718-723Crossref PubMed Scopus (470) Google Scholar, 6Visscher K. Schnitzer M.J. Block S.M. Nature. 1999; 400: 184-189Crossref PubMed Scopus (845) Google Scholar). However, processive movement also requires two heads, and coordination between these two motor domains is essential if kinesin is to move in an orderly fashion along its track (7Crevel I. Carter N. Schliwa M. Cross R. EMBO J. 1999; 18: 5863-5872Crossref PubMed Scopus (59) Google Scholar, 8Young E.C. Mahtani H.K. Gelles J. Biochem. 1998; 37: 3467-3479Crossref Scopus (52) Google Scholar). Two models have recently appeared to explain how the motor domains work together to bring about processive movement. In the hand-over-hand model (1Hancock W.O. Howard J. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 13147-13152Crossref PubMed Scopus (185) Google Scholar, 9Vugmeyster Y. Berliner E. Gelles J. Biochemistry. 1998; 37: 747-757Crossref PubMed Scopus (19) Google Scholar), each head alternates between a forward and rearward orientation on the microtubule, whereas in the inchworm model (10Hua W. Chung J. Gelles J. Science. 2002; 295: 844-848Crossref PubMed Scopus (175) Google Scholar), the forward and rearward heads retain their relative positions throughout the mechanochemical cycle. The hand-over-hand model proposes that ATP binding docks the neck linker of the attached head, and this swings the tethered head forward toward the next microtubule β subunit 8 nm away (11Coy D.L. Wagenbach M. Howard J. J. Biol. Chem. 1999; 276: 3667-3671Abstract Full Text Full Text PDF Scopus (302) Google Scholar, 12Schief W.R. Howard J. Curr. Opin. Cell Biol. 2001; 13: 19-28Crossref PubMed Scopus (110) Google Scholar). ATP hydrolysis on the rear head and ADP dissociation from the newly attached forward head follow, and they lead to dissociation of the rearward head from the microtubule. The net effect is hydrolysis of one ATP molecule per 8-nm step in the plus end direction (11Coy D.L. Wagenbach M. Howard J. J. Biol. Chem. 1999; 276: 3667-3671Abstract Full Text Full Text PDF Scopus (302) Google Scholar, 13Gilbert S.P. Moyer M.L. Johnson K.A. Biochemistry. 1998; 37: 792-799Crossref PubMed Scopus (154) Google Scholar). In the inchworm model, ATP binding, hydrolysis, and dissociation of ADP are associated with alternating separation and association of the two heads, only one of which is enzymatically active. However, precisely where nucleotide binding and hydrolysis fit into the overall scheme has not been determined (10Hua W. Chung J. Gelles J. Science. 2002; 295: 844-848Crossref PubMed Scopus (175) Google Scholar). Despite their differences, both models agree on several key features. First, ATP induces the two heads to separate by 8 nm, as one of them steps forward in the “+” direction. Second, at some point in the cycle, ATP is hydrolyzed, and phosphate is released. Third, ATP binding and hydrolysis changes the affinity of one of the kinesin motor domains for the microtubule, from “strong binding” to “weak binding,” and this strong to weak transition is associated with forward movement. Finally, the cycle is completed when the trailing head dissociates from the microtubule and rejoins its partner. However, several key questions remain unanswered by these models. What is the nature of the conformational change that produces the forward step? What enzymatic process actually produces this 8 nm separation, ATP binding or ATP hydrolysis? When and how quickly does the trailing head dissociate from the microtubule? How is the mechanical strain that is placed on the neck linkers by attachment of both heads relieved? Determining the speed and timing of kinesin's forward step and trailing head release are therefore crucial in evaluating each of the competing models and may help in their refinement as well as in the testing of their validity. In our previous study (14Rosenfeld S.S. Jefferson G.M. King P.H. J. Biol. Chem. 2001; 276: 40167-40174Abstract Full Text Full Text PDF PubMed Scopus (69) Google Scholar), we demonstrated that a fluorescent probe located on the neck linker of a kinesin monomeric construct could be used to measure in real time the reorientation and docking of the neck linker and to identify the step in the enzymatic cycle when this occurs. In this study, we have extended this approach to a dimeric kinesin construct to measure the kinetics of neck linker-neck linker and neck linker-microtubule separation. These measurements provide constraints that should allow us to critically examine current models of kinesin processivity. Fluorescent probes were obtained from Molecular Probes (Eugene, OR). Synthesis of mant nucleotides was carried out as described (15Hiratsuka T. Biochem. Biophys. Acta. 1983; 742: 496-508Crossref PubMed Scopus (396) Google Scholar). Our goal was to generate a kinesin construct of 413 residues that contained one reactive cysteine at position 333 and in which tryptophan 340 was mutated to phenylalanine. Using a plasmid encoding wild type human K413 (16Rosenfeld S.S. Correia J.J. Xing J. Rener B. Cheung H.C. J. Biol. Chem. 1996; 271: 30212-30222Abstract Full Text Full Text PDF PubMed Scopus (21) Google Scholar), the following PCR primers were used to amplify the native sequence: upstream primer, 5′-AAGTTGCATGTGTCTAGATATACATATGGCGGACCTGGCC-3′; downstream primer, 5′-AAGTTGCATGTGCTCGAGTAAAAAATTTCCTATAACTCCAAT-3′. The PCR product was digested with PstI and XhoI to generate a fragment, which was ligated into pBSK, and the resulting product was linearized with PstI and XbaI. A PCR fragment encoding the N-terminal 349 residues of a cysteine-light mutant of kinesin (kindly provided by Dr. Ron Vale, UCSF) was amplified with the upstream primer described above and with the following downstream primer, 5′-GCATGTGCTCGAGTTCTTTTTCTTTTTCATACTTCTTTTTGAACTGTTCTGCAGTTAAC-3′. The PCR product was digested with XbaI and PstI and ligated into the linearized pBSK plasmid described above to generate the CLM1-(1–413) construct, the cysteine-light mutant of K413. The following primers were then used to PCR-amplify the C-terminal 78 residues from CLM-(1–413): upstream primer, 5′-AGGAGTTAACTGCAGAACAGTTCAAAAAGAAGTATGAAAAAG-3′; downstream primer, 5′-AAGTTGCATGTGCTCGAGTTAAAAATTTCCTATAACTCCAAT-3′. The PCR product and CLM-(1–413) were digested with PstI andXhoI followed by ligation of the PCR product into CLM-(1–413). The mutated kinesin sequence was then excised with Nde and XhoI and ligated into pET21A for protein expression. K413W340F was labeled with 1,5-IAEDANS or tetramethyl rhodamine 5-maleimide (TMR) by incubation of a 10-fold molar excess of label over active sites in 100 mmKCl, 25 mm Hepes, 2 mm MgCl2, 1 mm EGTA, 100 μm Tris(carboxyethyl) phosphine, and 100 μm ATP, pH 7.5 for 3 h at room temperature. Excess label was removed by gel filtration or dialysis through Centriprep 10 concentrators (Amicon). Stoichiometry of labeling was 0.85–0.95 mol/mol of active site for both probes. Lifetime and anisotropy decay measurements were made with a pulsed fluorimeter as previously described (39Xing J. Wriggers W. Jefferson G.M. Stein R. Cheung H.C. Rosenfeld S.S. J. Biol. Chem. 2000; 275: 35413-35423Abstract Full Text Full Text PDF PubMed Scopus (34) Google Scholar). Transient kinetic measurements were made in an Applied Photophysics SX.18 MV stopped flow spectrometer with instrument dead time of 1.2 ms. Complexes of kinesin and microtubules were formed with a 5–10-fold molar excess of microtubules over active sites. ADP was added to 3 μm to ensure that the tethered kinesin head of a kinesin·microtubule complex contained ADP in the active site. Dansyl fluorescence was generated by exciting at 295 nm (for tryptophan energy transfer), and the emission was monitored through a 500-nm broad bandpass filter with a transmission half-width of 20 nm (Omega Optical). Rhodamine fluorescence was monitored by exciting at 520 nm and monitoring emission through a 590-nm cutoff filter. Fluorescence of mant ATP was monitored by exciting at 278 nm. At this wavelength, the mant fluorophor is excited by fluorescence energy transfer from vicinal tyrosine residues of kinesin (17Cheng J-Q Jiang W. Hackney D.D. Biochemistry. 1998; 37: 5288-5295Crossref PubMed Scopus (43) Google Scholar). We have found that for dimeric kinesin constructs, the mant fluorescence emission at this exciting wavelength is not affected by the presence of microtubules (data not shown). The neck linker of kinesin is a region in the C terminus of the motor domain that plays a key role in mechanochemical transduction by this motor (3Vale R.D. Milligan R.A. Science. 2000; 288: 88-95Crossref PubMed Scopus (1247) Google Scholar, 18Vale R.D. Case R. Sablin E. Hart C. Fletterick R. Phil. Trans. Royal Soc. B. 2000; 355: 449-457Crossref PubMed Scopus (33) Google Scholar, 19Rice S. Lin A.W. Safer D. Hart C.L. Naber N. Carragher B.O. Cain S.M. Pechatnikova E. Wilson-Kubalek E.M. Whittaker M. Pate E. Cooke R. Taylor E.W. Vale R. Nature. 1999; 402: 778-784Crossref PubMed Scopus (650) Google Scholar). Our intent was to attach fluorescent probes via a cysteine residue at position 333, within the neck linker sequence. In order to do this we engineered a kinesin dimer devoid of reactive cysteines and with a cysteine mutation at position 333. Furthermore, since we intended to perform FRET studies between microtubule tryptophans and an AEDANS probe at position 333 of kinesin, we mutated tryptophan 340 to phenylalanine. The kinesin mutant thus generated, referred to as K413W340F, demonstrated no energy transfer between the remaining tryptophan residues at positions 360 and 368 and an AEDANS probe at position 333. This is to be expected, given the 35–40 Å distance separating these residues (20Kozielski F. Sack S. Marx A. Thormahlen M. Schonbrunn E. Biou V. Thompson A. Mandelkow E.-M. Mandelkow E. Cell. 1997; 91: 985-994Abstract Full Text Full Text PDF PubMed Scopus (361) Google Scholar) and the value of Ro for this donor-acceptor pair (∼20 Å, Ref. 14Rosenfeld S.S. Jefferson G.M. King P.H. J. Biol. Chem. 2001; 276: 40167-40174Abstract Full Text Full Text PDF PubMed Scopus (69) Google Scholar). Fig.1 depicts the wild type kinesin dimeric structure derived from Rattus norvegicus (20Kozielski F. Sack S. Marx A. Thormahlen M. Schonbrunn E. Biou V. Thompson A. Mandelkow E.-M. Mandelkow E. Cell. 1997; 91: 985-994Abstract Full Text Full Text PDF PubMed Scopus (361) Google Scholar). In this figure, structures relevant to our study are depicted in the space-filling format, including the bound nucleotide (magenta), the neck linker (yellow), and leucine 335 (in red and equivalent to valine 333 in the human sequence). K413W340F has a microtubule-activated ATPase activity, which at 10 mm KCl has values of 19.0 ± 1.9 s−1 and 0.25 ± 0.10 μm for k cat andK 0.5,MT, respectively. Labeling cysteine 333 with either 1,5-IAEDANS or TMR did not appreciably alter these values (data not shown). Although the value of k cat is quite close to values reported for similar Drosophilaconstructs, the ratio ofk cat/K 0.5,MT (referred to as k bi(ATPase)) is ∼4-fold smaller (4Jiang W Stock M.F., Li, X. Hackney D.D. J. Biol. Chem. 1997; 272: 7626-7632Abstract Full Text Full Text PDF PubMed Scopus (75) Google Scholar, 21Moyer M.L. Gilbert S.P. Johnson K.A. Biochemistry. 1998; 37: 800-813Crossref PubMed Scopus (127) Google Scholar). We measured the kinetics of the weak-to-strong transition in this construct by mixing in the stopped flow a complex of 2′-deoxy mant-ADP·K413W340F with microtubules and 1 mm ATP and monitoring nucleotide release (2Ma Y.-Z. Taylor E.W. J. Biol. Chem. 1997; 272: 717-723Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar, 13Gilbert S.P. Moyer M.L. Johnson K.A. Biochemistry. 1998; 37: 792-799Crossref PubMed Scopus (154) Google Scholar, 22Ma Y.Z. Taylor E.W. J. Biol. Chem. 1997; 272: 724-730Abstract Full Text Full Text PDF PubMed Scopus (166) Google Scholar). The resulting fluorescence decay demonstrated a linear dependence on microtubule concentration, defining an apparent second order rate constant,k bi(ADP) of 1.04 ± 0.07 μm−1 s−1 (data not shown). The ratiok bi(ATPase)/k bi(ADP) is a measure of the number of ATP molecules hydrolyzed per processive run. Its value is 76, which compares to the previously reported value of ∼100 for native kinesin constructs of comparable size (4Jiang W Stock M.F., Li, X. Hackney D.D. J. Biol. Chem. 1997; 272: 7626-7632Abstract Full Text Full Text PDF PubMed Scopus (75) Google Scholar). Thus, although our mutagenesis of K413W340F did alter the affinity of a kinesin·ADP state for microtubules (23Rosenfeld S.S. Correia J.J. Mayo M.S. Rener B. Cheung H.C. J. Biol. Chem. 1996; 271: 9473-9482Abstract Full Text Full Text PDF PubMed Scopus (49) Google Scholar), it did not appear to have appreciably affected its processivity. We first sought to label the K413W340F dimer at position 333 with an optical probe whose fluorescence would be sensitive to the distance between the two neck linkers of the motor. Previous reports had demonstrated that the fluorescent probe TMR dimerizes under favorable conditions (24Packard B.Z. Toptygin D.D. Komoriya A. Brand L Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 11640-11645Crossref PubMed Scopus (137) Google Scholar, 25Hamman B.D. Oleinikov A.V. Jokhadze G.G. Bochkariov D.E. Traut R.R. Jameson D.M. J. Biol. Chem. 1996; 271: 7568-7573Abstract Full Text Full Text PDF PubMed Scopus (55) Google Scholar, 26Geoghegan K.F. Rosner P.J. Hoth L.R. Bioconjugate Chem. 2000; 11: 71-77Crossref PubMed Scopus (32) Google Scholar). We therefore reasoned that labeling dimeric kinesin at position 333 with TMR might produce a motor whose rhodamine probes would alternately dimerize and separate during each mechanochemical cycle, as the motor steps along on the microtubule. Monomeric rhodamine demonstrates a peak in its absorption spectrum at 555 nm, a shoulder at 518 nm, and a 518/555 nm ratio of 0.5 (24Packard B.Z. Toptygin D.D. Komoriya A. Brand L Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 11640-11645Crossref PubMed Scopus (137) Google Scholar, 25Hamman B.D. Oleinikov A.V. Jokhadze G.G. Bochkariov D.E. Traut R.R. Jameson D.M. J. Biol. Chem. 1996; 271: 7568-7573Abstract Full Text Full Text PDF PubMed Scopus (55) Google Scholar, 26Geoghegan K.F. Rosner P.J. Hoth L.R. Bioconjugate Chem. 2000; 11: 71-77Crossref PubMed Scopus (32) Google Scholar). Dimerization strongly quenches the rhodamine fluorescence emission. It also produces a blue shift in the absorption spectrum, with two discrete peaks at 555 and 518 nm and with an increase in the 518/555 nm ratio to 1.4 (24Packard B.Z. Toptygin D.D. Komoriya A. Brand L Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 11640-11645Crossref PubMed Scopus (137) Google Scholar, 25Hamman B.D. Oleinikov A.V. Jokhadze G.G. Bochkariov D.E. Traut R.R. Jameson D.M. J. Biol. Chem. 1996; 271: 7568-7573Abstract Full Text Full Text PDF PubMed Scopus (55) Google Scholar, 26Geoghegan K.F. Rosner P.J. Hoth L.R. Bioconjugate Chem. 2000; 11: 71-77Crossref PubMed Scopus (32) Google Scholar, 27del Monte F. Levy D. J. Phys. Chem. B. 1999; 103: 8080-8086Crossref Scopus (44) Google Scholar, 28del Monte F. Mackenzie J.D. Levy D. Langmuir. 2000; 16: 7377-7382Crossref Scopus (135) Google Scholar). Fig. 2 Ashows that the absorption spectrum of TMR-labeled K413W340F (dotted spectrum) has the features characteristic of rhodamine dimer. Addition of guanidine hydrochloride to 6 menhances the 555 nm absorbance and reduces the 518 peak (solid spectrum), as expected since unfolding of K413W340F would separate the rhodamine probes. The fluorescence emission spectrum of TMR-labeled K413W340F (Fig. 2 B, dotted spectrum) is highly quenched, and addition of 6 m guanidine hydrochloride enhances fluorescence over 10-fold (Fig. 2 B, solid spectrum). In the absence of added nucleotide, K413W340F binds to microtubules via one of its two motor domains (29Kawaguchi K. Ishiwata S.-I. Science. 2001; 291: 667-669Crossref PubMed Scopus (108) Google Scholar, 33Uemura S. Kawaguchi K. Yajima J. Edamatsu M. Toyoshima Y.Y. Ishiwata S.-I. Proc. Natl. Acad. Sci. U. S. A. 2002; 99: 5977-5981Crossref PubMed Scopus (106) Google Scholar). The absorption spectrum of a complex of rhodamine K413W340F + microtubules under rigor conditions demonstrated a reduction in the 518/555 nm ratio (1.3 versus1.5 for TMR-labeled K413W340F). This was reduced to 1.2 in the presence of ATP under conditions that support processive movement (10 mm KCl) and is illustrated in Fig. 2 C as the dashed spectrum. In the presence of 1 mm AMPPNP, both heads of kinesin attach strongly to the microtubule (29Kawaguchi K. Ishiwata S.-I. Science. 2001; 291: 667-669Crossref PubMed Scopus (108) Google Scholar). This is reflected in Fig. 2 C by a further reduction of the 518/555 nm ratio to 1.1 (solid spectrum). This data suggests that under conditions that favor processive movement, an appreciable fraction of kinesin molecules are attached to the microtubule via both heads. Our spectroscopic results allowed us to make testable predictions about the time-dependent fluorescence emission of rhodamine-labeled K413W340F as it moves along the microtubule. The first of these is illustrated in Fig. 3 A. Kinesin enters its mechanochemical cycle with one head strongly attached to the microtubule. The fluorescence of this species should be relatively quenched. ATP binding to the attached head should swing the tethered head forward to attach to the next microtubule-binding site, pulling the two heads apart by 8 nm. We would therefore predict that mixing a complex of rhodamine-labeled kinesin·microtubules with ATP should produce an initial fluorescence rise, followed by a fall as dissociation of the trailing head allows the neck linkers to spring back together. The red and green transients marked 10 mmKCl and 100 mmKCl in Fig.3 B confirm this, and the inset in the figure demonstrates the rising phase over a shorter time scale. At 100 mm KCl, the fluorescence at the completion of this transient is lower than that at the start. This would be expected, since high ionic strength reduces kinesin's affinity for the microtubule, and dissociation of the trailing head should be followed by dissociation of the motor (2Ma Y.-Z. Taylor E.W. J. Biol. Chem. 1997; 272: 717-723Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar, 3Vale R.D. Milligan R.A. Science. 2000; 288: 88-95Crossref PubMed Scopus (1247) Google Scholar,30Gilbert S.P. Webb M.R. Brune M. Johnson K.A. Nature. 1995; 373: 671-676Crossref PubMed Scopus (248) Google Scholar). By contrast, at 10 mm KCl, the final fluorescence is ∼50% greater than the starting fluorescence, which implies that during a processive run, the motors partition between singly and doubly attached species. Fig. 3 B also shows that repeating this experiment in the absence of microtubules (blue transient markedno microtubules) produces no change in rhodamine emission. Our second prediction is that the rate of the rising phase should vary hyperbolically with ATP concentration, and should be no faster than the rate of ATP binding. Fig. 4 (open boxes) confirms this and defines a maximum rate of 763 ± 84 s−1 and apparent dissociation constant of 98 ± 24 μm in 100 mm KCl. Similar results were seen at low ionic strength (data not shown). We measured the rate of 2′-deoxy mant-ATP binding to the attached head by monitoring FRET from kinesin tyrosine residues to the mant fluorophor (17Cheng J-Q Jiang W. Hackney D.D. Biochemistry. 1998; 37: 5288-5295Crossref PubMed Scopus (43) Google Scholar). Fig. 4 shows the rate of mant nucleotide binding (closed triangles) was consistently as fast or faster than the rate of the rising phase of the rhodamine transient. It varied linearly with nucleotide concentration, defining an apparent second order rate constant of 4.9 ± 1.1 μm−1 s−1 with an extrapolated dissociation rate constant of 73 ± 24 s−1. This dissociation rate is similar to prior measurements on native kinesin constructs (2Ma Y.-Z. Taylor E.W. J. Biol. Chem. 1997; 272: 717-723Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar, 21Moyer M.L. Gilbert S.P. Johnson K.A. Biochemistry. 1998; 37: 800-813Crossref PubMed Scopus (127) Google Scholar, 22Ma Y.Z. Taylor E.W. J. Biol. Chem. 1997; 272: 724-730Abstract Full Text Full Text PDF PubMed Scopus (166) Google Scholar). Once kinesin has stepped forward, its trailing head dissociates from the microtubule. This should allow the neck linkers to spring back together, reform rhodamine dimers, and quench the rhodamine fluorescence. At 100 mm KCl, the rate of this phase should match the rate at which kinesin dissociates from the microtubule, since the microtubule affinity of K413W340F·ADP is reduced, and dissociation is rapid. Our third prediction is that the rate of the decrease in rhodamine fluorescence at high ionic strength should match the rate at which ATP dissociates kinesin from the microtubule. Fig.5 confirms this. The rate of ATP-induced dissociation at 100 mm KCl, as measured by turbidity (closed circles) is close to that of the falling phase of the rhodamine transient at this ionic strength (closed triangles), with an extrapolated maximum rate of ∼50–60 s−1. At 10 mm KCl, the falling phase of the rhodamine transient fit a double exponential decay, and the faster phase constituted 75–80% of the total signal amplitude. This phase showed an ATP concentration dependence essentially identical to that at 100 mm KCl (Fig. 5, open boxes). The slower phase showed little ATP concentration dependence, and its rate ranged between 1.1–2.4 s−1. At this ionic strength, little dissociation could be detected with turbidity over the time course of the rhodamine transient (data not shown). This strongly suggests that the rate of ATP-induced decrease in rhodamine fluorescence is controlled by (and therefore is a measure of) how fast the trailing head can dissociate from the microtubule. Our fourth prediction, illustrated in Fig.6 A, is that mixing rhodamine-labeled K413W340F + microtubules with ADP should produce a small decrease in fluorescence, as the partially separated neck linkers reapproximate with microtubule dissociation. The orange fluorescence transient depicted in Fig. 6 C (ADP) confirms this. A small amplitude, single exponential decay at 1 mmADP and 100 mm KCl (final concentrations) was observed, corresponding to a 4% decrease in fluorescence intensity, with a rate constant of 128 ± 21 s−1 (n = 10). This rate is ∼7–8 fold faster than the corresponding process with native kinesin, and is consistent with the lower affinity of the K413W340F·ADP for the microtubule (31Ma Y.-Z. Taylor E.W. Biochemistry. 1995; 34: 13233-13241Crossref PubMed Scopus (71) Google Scholar, 32Hackney D.D. Biochemistry. 2002; 41: 4437-4446Crossref PubMed Scopus (39) Google Scholar). AMPPNP binding causes both heads of kinesin to attach to the microtubule (29Kawaguchi K. Ishiwata S.-I. Science. 2001; 291: 667-669Crossref PubMed Scopus (108) Google Scholar, 33Uemura S. Kawaguchi K. Yajima J. Edamatsu M. Toyoshima Y.Y. Ishiwata S.-I. Proc. Natl. Acad. Sci. U. S. A. 2002; 99: 5977-5981Crossref PubMed Scopus (106) Google Scholar). Our fifth prediction, illustrated in Fig.6 B, is that mixing with AMPPNP should produce a fluorescence transient consisting only of a rising phase. Furthermore, the initial fluorescence intensity should be the same as that for mixing with ADP or ATP, because in each case the starting conditions are the same. The red transient marked AMPPNP in Fig. 6 C confirms these predictions. Mixing with 1 mm AMPPNP and 100 mm KCl (final concentrations) produced a biphasic transient with rates of 48 ± 19 s−1 and 1.6 ± 1.1 s−1 (n = 10), and the amplitude of the faster phase constituted 40% of the total amplitude. The rate of the faster phase is similar to the previously reported rate of AMPPNP-accelerated ADP release (13Gilbert S.P. Moyer M.L. Johnson K.A. Biochemistry. 1998; 37: 792-799Crossref PubMed Scopus (154) Google Scholar). However, the presence of a second, slower phase suggests that attachment of the second head to the microtubule may occur in two steps. The green transient in Fig.6 C was produced by mixing with 1 mm ATP and 100 mm KCl (final concentrations). Fitting to two exponential terms, as per Fig. 3, reveals that the extrapolated amplitude of the rising phase is 87% of the total signal amplitude for AMPPNP. Several further predictions are also supported by our results. Mixing unlabeled K413W340F + microtubules with ATP produces no signal beyond that of buffer alone (data not shown), which establishes that the signal change seen in Fig. 3 B is not due to light scattering. Furthermore, repeating these experiments with a rhodamine labeled monomeric kinesin (K349) + microtubules produced no fluorescence change (data not shown). Our conclusions from the rhodamine probe would be supported if a different probe, sensitive to neck linker association with the microtubule, could provide a direct measure of the rate of trailing head dissociation that agreed with the rhodamine data. For this purpose, we utilized an AEDANS probe on the kinesin neck linker. In our previous study, we demonstrated that an AEDANS probe attached in the neck linker of a monomeric kinesin construct could be excited by energy transfer from the microtubule tryptophan residues (14Rosenfeld S.S. Jefferson G.M. King P.H. J. Biol. Chem. 2001; 276: 40167-40174Abstract Full Text Full Text PDF PubMed Scopus (69) Google Scholar), and we established that this probe could monitor kinesin-microtubule association. We labeled K413W340F at position 333 on the neck linker with AEDANS, monitored microtubule binding by FRET, and found that binding produced a 21% fluorescence enhancement in the absence of nucleotide (data not shown). In our previous study, we also had shown that binding of ATP to an AEDANS-kinesin·microtubule complex reduced the FRET efficiency of the AEDANS probe (14Rosenfeld S.S. Jefferson G.M. King P.H. J. Biol. Chem. 2001; 276: 40167-40174Abstract Full Text Full Text PDF PubMed Scopus (69) Google Scholar). This, plus the rapid rate at which a doubly attached intermediate would be formed (>750 s−1, Fig. 4), means that mixing with ATP would likely produce a low amplitude rising phase that may not be observable in the stopped flow. This would then be followed by a fluorescence decay as the trailing head releases from the microtubule (Fig. 7 A). Furthermore, at 10 mm KCl, the amplitude of this decay would be expected to be smaller than at 100 mm KCl, sin" @default.
• W2000315199 created "2016-06-24" @default.
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• W2000315199 creator A5020641571 @default.
• W2000315199 creator A5060459957 @default.
• W2000315199 creator A5074695853 @default.
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• W2000315199 date "2002-09-01" @default.
• W2000315199 modified "2023-09-27" @default.
• W2000315199 title "Measuring Kinesin's First Step" @default.
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