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• W2055665417 abstract "Cytoplasmic transport is mediated by a group of molecular motors that typically work in isolation, under conditions where they must move their cargos long distances without dissociating from their tracks. This processive behavior requires specific adaptations of motor enzymology to meet these unique physiologic demands. One of these involves the ability of the two heads of a processive motor to communicate their structural states to each other. In this study, we examine a processive motor from the myosin superfamily myosin V. We have measured the kinetics of nucleotide release, of phosphate release, and of the weak-to-strong transition, as this motor interacts with actin, and we have used these studies to develop a model of how myosin V functions as a transport motor. Surprisingly, both heads release phosphate rapidly upon the initial encounter with an actin filament, suggesting that there is little or no intramolecular strain associated with this step. However, ADP release can be affected by both forward and rearward strain, and under steady-state conditions it is essentially prevented in the lead head until the rear head detaches. Many of these features are remarkably like those underlying the processive movement of kinesin on microtubules, supporting our hypothesis that different molecular motors satisfy the requirement for processive movement in similar ways, regardless of their particular family of origin. Cytoplasmic transport is mediated by a group of molecular motors that typically work in isolation, under conditions where they must move their cargos long distances without dissociating from their tracks. This processive behavior requires specific adaptations of motor enzymology to meet these unique physiologic demands. One of these involves the ability of the two heads of a processive motor to communicate their structural states to each other. In this study, we examine a processive motor from the myosin superfamily myosin V. We have measured the kinetics of nucleotide release, of phosphate release, and of the weak-to-strong transition, as this motor interacts with actin, and we have used these studies to develop a model of how myosin V functions as a transport motor. Surprisingly, both heads release phosphate rapidly upon the initial encounter with an actin filament, suggesting that there is little or no intramolecular strain associated with this step. However, ADP release can be affected by both forward and rearward strain, and under steady-state conditions it is essentially prevented in the lead head until the rear head detaches. Many of these features are remarkably like those underlying the processive movement of kinesin on microtubules, supporting our hypothesis that different molecular motors satisfy the requirement for processive movement in similar ways, regardless of their particular family of origin. Cytoplasmic transport motors are found in both the myosin and kinesin families and typically work in isolation (1Mehta A. J. Cell Sci. 2001; 114: 1981-1998PubMed Google Scholar, 2Goldstein L.S.B. Philp A.V. Annu. Rev. Cell Dev. Biol. 1999; 15: 141-183Crossref PubMed Scopus (217) Google Scholar). This physiology places a unique requirement on them: they must remain attached to their respective tracks through multiple ATPase cycles. This need for processivity demands that, at any given time, at least one of the two motor-containing “heads” remains strongly attached to its track to prevent the entire motor from prematurely detaching. Such coordination requires appropriate kinetics as well as structural communication between the heads to optimize processive movement. Myosin V is the first of the myosin superfamily members shown to be processive (3Forkey J.N. Quinlan M.E. Shaw M.A. Corrie J.E. Goldman Y.E. Nature. 2003; 422: 399-404Crossref PubMed Scopus (383) Google Scholar, 4Yildiz A. Forkey J.N. McKinney S.A. Ha T. Goldman Y.E. Selvin P.R. Science. 2003; 300: 2061-2065Crossref PubMed Scopus (1553) Google Scholar, 5Veigel C. Wang F. Bartoo M.L. Sellers J.R. Molloy J.E. Nat. Cell Biol. 2001; 4: 59-65Crossref Scopus (333) Google Scholar, 35Mehta A.D. Rock R.S. Rief M. Spudich J.A. Mooseker M.S. Cheney R.E. Nature. 1999; 400: 590-593Crossref PubMed Scopus (675) Google Scholar). The mechanism underlying its movement has been studied extensively over the past 4 years, and a general picture of how it works has emerged (6Vale R.D. J. Cell Biol. 2003; 163: 445-450Crossref PubMed Scopus (111) Google Scholar). The motor needs to take 36-nm steps along an actin filament to avoid spiraling around the actin filament. Myosin V accomplishes this by possessing large lever arms made up of six “IQ motifs” and their associated light chains (7Purcell T.J. Morris C. Spudich J.A. Sweeney H.L. Proc. Natl. Acad. Sci. U. S. A. 2002; 99: 14159-14164Crossref PubMed Scopus (146) Google Scholar, 8Espindola F.S. Suter D.M. Partata L.B. Cao T. Wolenski J.S. Cheney R.E. King S.M. Mooseker M.S. Cell Motil. Cytoskeleton. 2000; 47: 269-281Crossref PubMed Google Scholar), and it uses these long lever arms to walk hand over hand along an actin filament for many steps before dissociating. How then is this processive behavior maintained? The processive behavior of myosin V presumably reflects specific adaptations it has made in its enzymology, which in aggregate facilitates its function as a transport motor. One of these adaptations is the identity of the rate-limiting step of the actin-activated ATPase cycle. The rate-limiting step for most isoforms of myosin II is phosphate release, which insures that the motor spends the majority of its ATPase cycle either detached or weakly bound to actin (i.e. low duty ratio). However, for myosin V, the rate-limiting step is ADP release, which allows the myosin to remain strongly bound to actin for the majority of the cycle (i.e. high duty ratio). The result of this difference is that myosin V has a duty ratio of ∼0.9 at high actin concentrations, compared with duty ratios of 0.03-0.04 for smooth and skeletal muscle myosin II (9De La Cruz E.M. Wells A.L. Rosenfeld S.S. Ostap E.M. Sweeney H.L. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 13726-13731Crossref PubMed Scopus (355) Google Scholar). This large duty ratio is a feature that has been thought to be essential for processive, transport motors, because it would reduce the probability of premature dissociation from actin. However, a large duty ratio by itself may not be enough to ensure that myosin V can function properly in its transport role, because we have shown that myosin IIB, a member of the myosin II family designed for tension generation, also has a high duty ratio (10Rosenfeld S.S. Xing J. Chen L.-Q. Sweeney H.L. J. Biol. Chem. 2003; 278: 27449-27455Abstract Full Text Full Text PDF PubMed Scopus (91) Google Scholar). In addition to needing large duty ratios, the two heads of processive motors must also communicate their structural states to each other throughout their mechanochemical cycles. This communication is needed to ensure that the two heads do not weakly bind to their track simultaneously, an event that would lead to premature dissociation and that could have dire physiologic consequences. In the case of myosin V, how this allosteric communication is carried out and how it is coupled to the ATPase cycle remain unclear. Three models have recently emerged (5Veigel C. Wang F. Bartoo M.L. Sellers J.R. Molloy J.E. Nat. Cell Biol. 2001; 4: 59-65Crossref Scopus (333) Google Scholar, 11Rief M. Rock R.S. Mehta A.D. Mooseker M.S. Cheney R.E. Spudich J.A. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 9482-9486Crossref PubMed Scopus (359) Google Scholar, 12De La Cruz E.M. Ostap E.M. Sweeney H.L. J. Biol. Chem. 2001; 276: 32373-32381Abstract Full Text Full Text PDF PubMed Scopus (201) Google Scholar). In the first, the intramolecular strain that develops when both heads are bound to the actin filament mediates this allosteric communication. In this model, rearward strain on the forward head prevents this motor domain from binding ATP and detaching until the strain is relieved by detachment of the trailing head (11Rief M. Rock R.S. Mehta A.D. Mooseker M.S. Cheney R.E. Spudich J.A. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 9482-9486Crossref PubMed Scopus (359) Google Scholar). In the second model, forward strain on the rearward head accelerates its detachment from actin (5Veigel C. Wang F. Bartoo M.L. Sellers J.R. Molloy J.E. Nat. Cell Biol. 2001; 4: 59-65Crossref Scopus (333) Google Scholar). In both of these models, intramolecular strain plays a central role, much as in the case of microtubule motor kinesin (13Rosenfeld S.S. Fordyce P. Jefferson G.M. King P.H. Block S.M. J. Biol. Chem. 2003; 278: 18550-18556Abstract Full Text Full Text PDF PubMed Scopus (158) Google Scholar), where we have shown that strain accelerates motor dissociation from the trailing head and blocks motor dissociation from the leading head. In the third model, intramolecular strain plays no role at all (12De La Cruz E.M. Ostap E.M. Sweeney H.L. J. Biol. Chem. 2001; 276: 32373-32381Abstract Full Text Full Text PDF PubMed Scopus (201) Google Scholar). Rather, strong binding by the rearward head prevents actin binding and phosphate release by the forward head. Processivity in this model would be favored if ATP binding to the rearward head were to lead to rapid and strong binding of the forward head. Thus, it remains unclear whether strain plays a role at all in the processivity of myosin V, and if so, what effect it has on the kinetics of specific transitions in the mechanochemical cycle. Addressing these issues is the focus of this study. Our results show that both forward and rearward strain can affect the kinetics of crucial structural transitions in the myosin V ATPase cycle. However, under steady-state conditions, it is the effect of rearward strain in retarding ADP release from the forward head that provides the critical coordination for processivity. Our results also show that phosphate release is rapid for both heads and occurs before the heads bind strongly to actin. Taken together these data provide the first evidence that phosphate is released immediately upon myosin V binding to actin and that it occurs prior to formation of a strong binding state and development of intramolecular strain. Our results demonstrate that only the release of ADP is strain-sensitive, ensuring that both heads will be strongly bound to actin once the lead head finds an actin-binding site. Thus the degree of processivity of myosin V is ultimately limited by the rate at which a newly detached head can find an actin binding site. This arrangement ensures that both heads will be strongly bound to actin once the lead head finds an actin-binding site. Reagents—The N-methylanthraniloyl derivative of 2′deoxy-ADP was synthesized as described previously (24Hiratsuka T. Biochim. Biophys. Acta. 1983; 742: 496-508Crossref PubMed Scopus (396) Google Scholar). N-1-Pyrenyl iodoacetamide was obtained from Molecular Probes (Portland, OR). Protease inhibitors and chemicals used for buffers were obtained from Sigma. Pre-poured Sephadex G-25 columns (PD10) were obtained from Amersham Biosciences. Proteins—Actin was prepared from rabbit acetone powder, and labeling at cysteine 374 with N-1-pyrenyl iodoacetamide was carried out as described before (14Taylor E.W. J. Biol. Chem. 1991; 266: 294-302Abstract Full Text PDF PubMed Google Scholar). Phosphate-binding protein was purified from Escherichia coli and labeled with MDCC 1The abbreviations used are: MDCC, N-[2-(1-maleimidyl)ethyl]-7-(diethylamino)coumarin-3-carboxamide; mant, N-methylanthraniloyl; 2′dmD, 2′-deoxy-mant-ADP; 2′dmT, 2′-deoxy-mant-ATP; PBP, phosphate-binding protein; S1, monomeric construct of myosin V; HMM; dimeric construct of myosin V. as described (25Brune M. Hunter J.L. Corrie J.E.T. Webb M.R. Biochemistry. 1994; 33: 8262-8271Crossref PubMed Scopus (432) Google Scholar). Recombinant Myosin V Expression and Purification—Chicken myosin V cDNA was expressed in two forms. The constructs were based on the previously described two-headed (HMM-like) myosin construct, myosin V-6IQ HMM (7Purcell T.J. Morris C. Spudich J.A. Sweeney H.L. Proc. Natl. Acad. Sci. U. S. A. 2002; 99: 14159-14164Crossref PubMed Scopus (146) Google Scholar). The construct was used to create either a two-headed (HMM-like) or single-headed (S1-like) construct. This myosin V-6IQ HMM heavy chain was truncated at Glu-1099, to which was added a leucine zipper (GCN4) to ensure dimerization, then followed by a FLAG tag (for purification). For the single-headed (S1-like) construct, the coiled-coil was removed and the heavy chain was truncated at amino acid Lys-910 to create myosin V-6IQ S1. As for the HMM, a FLAG tag was added after Lys-910 to facilitate purification. Recombinant baculoviruses were generated and used for co-expression in SF9 cells with calmodulin and essential light chains (7Purcell T.J. Morris C. Spudich J.A. Sweeney H.L. Proc. Natl. Acad. Sci. U. S. A. 2002; 99: 14159-14164Crossref PubMed Scopus (146) Google Scholar). Using the methodology detailed previously (26Sweeney H.L. Rosenfeld S.S. Brown F Faust L Smith J. Xing J. Stein L.A. Sellers J.R. J. Biol. Chem. 1998; 273: 6262-6270Abstract Full Text Full Text PDF PubMed Scopus (200) Google Scholar), purified myosin V HMM or S1 protein was obtained. Fig. 1 illustrates an SDS-PAGE of a purified myosin V HMM 6IQ preparation and demonstrates that a typical yield has high purity and minimal evidence of proteolysis. Concentrations of recombinant myosin V preparations were determined with the Bio-Rad protein assay. The reference samples were recombinant myosin V HMM and S1 whose concentrations were determined by absorbance, using calculated extinction coefficients of 540,240 m-1 cm-1 for HMM and 129,610 m-1 cm-1 for S1. Complexes of myosin V constructs with 2′dmD were formed by pre-incubating S1 and HMM with a 20-fold molar excess of 2′dmD, followed by gel filtration on pre-poured Sephadex G-25 columns (PD10, Amersham Biosciences) according to the manufacturer's instructions. Fractional labeling of the complexes with 2′dmD was determined by using the extinction coefficient of the fluorescent nucleotide (5700 m-1 cm-1 at 356 nm (24Hiratsuka T. Biochim. Biophys. Acta. 1983; 742: 496-508Crossref PubMed Scopus (396) Google Scholar)) and the measured protein concentration. Ratios of 2′dmD to active site concentration were typically 0.90-0.95. Kinetic Methodologies—Kinetic measurements were made using an Applied Photophysics SX.18MV stopped-flow spectrophotometer with an instrument dead time of 1.2 ms (27Rosenfeld S.S. Xing J. Jefferson G.M. Cheung H.C. King P.H. J. Biol. Chem. 2002; 277: 36731-36739Abstract Full Text Full Text PDF PubMed Scopus (44) Google Scholar). The excitation and emission wavelengths for monitoring pyrene-labeled actin and MDCC-labeled phosphate-binding protein fluorescence have been described (10Rosenfeld S.S. Xing J. Chen L.-Q. Sweeney H.L. J. Biol. Chem. 2003; 278: 27449-27455Abstract Full Text Full Text PDF PubMed Scopus (91) Google Scholar). For studies of 2′dmD and 2′dmD·Pi release, mant fluorescence was monitored by both direct excitation (λex = 356 nm) and by energy transfer from vicinal tryptophan residues (λex = 295 nm), and both methods gave similar results. Our experimental approach was to compare the kinetics of key steps of the myosin V HMM mechanochemical cycle during the first one to two turnovers to those for an S1 construct, because this would allow us to test the role of internal strain in shaping the kinetics of native myosin V. We also examined the kinetics of several of these steps under steady-state conditions to evaluate the physiologic relevance of our findings. Actin Binding in the Presence of ADP—Previous studies have noted that dimeric myosin V constructs can aggregate actin filaments under conditions that favor strong binding (28Walker M.L. Burgess S.A. Sellers J.R. Wang F. Hammer J.A. Trinick J. Knight P.J. Nature. 2000; 405: 804-807Crossref PubMed Scopus (283) Google Scholar, 29Cheney R.E. O'Shea M.K. Heuser J.E. Coelho M.V. Wolenski J.S. Espreafico E.M. Forscher P. Larson R.E. Mooseker M.S. Cell. 1993; 75: 13-23Abstract Full Text PDF PubMed Scopus (380) Google Scholar). This effect is presumed to be due to the presence of extended lever arms, which would allow cross-linking of actin filaments and could interfere with spectroscopic measurements. We have addressed this issue by comparing the kinetics of the light scattering increase produced by mixing myosin V S1·ADP with actin to those using HMM·ADP. Fig. 2A illustrates a typical light scattering transient, produced by mixing 0.8 μm S1 or HMM (active site concentration) with 6 μm actin in the presence of 1 mm ADP. For S1 (red), the transient consisted of a rapid phase (inset, red transient), sometimes accompanied by a low amplitude (<5% total) slow phase. By contrast, mixing with HMM (green) produced two well separated phases. The faster phase (inset, green transient) constituted ∼60% of the total signal amplitude. The slower phase fit a single exponential process (solid curve), showed little actin concentration dependence, and measured 0.03-0.05 s-1. We propose that the fast phase for both the HMM and S1 transients represents strong binding of at least one head to actin, whereas the slow phase seen with HMM is due to cross-linking. The HMM-induced cross-linking could be due to one of two possibilities. First, steric constraints on the lever arm could prevent the second head of HMM from binding strongly to the same actin filament. Because of its limited rotational freedom in the ADP state, the second head would only be able to slowly attach to neighboring actin filaments, producing a cross-linked aggregate. This possibility would predict that mixing HMM with pyrene-labeled actin in the presence of ADP should produce a fluorescence decrease that is the mirror image of the light scattering transient, e.g. two phases of similar amplitude, with the slower phase characterized by a rate constant of 0.03-0.05 s-1. The second possibility is that binding of both heads of HMM to the same actin filament is kinetically favored but leads to a system that is internally strained. In this case, strain could be relieved by release of the forward head (with rate constant of 0.03-0.05 s-1), followed by attachment to a neighboring actin filament. In this scenario, mixing of HMM with pyrene-labeled actin should produce a fast fluorescence decrease that is not followed by a slow phase. Furthermore, this possibility would predict that the amplitude of the fast fluorescence decrease should be the same as that for an equimolar S1 concentration. Fig. 2B illustrates the results of mixing 0.8 μm (active site concentration) S1·ADP (green) or HMM·ADP (red) with 6 μm pyrene-labeled actin (50% labeled). Strong binding to actin quenches the pyrenyl fluorescence, and for both S1 and HMM, this process (inset) is considerably more rapid than cross-linking (Fig. 2A). A detailed analysis of the kinetics of this rapid quenching demonstrates biphasic decays for both HMM and S1 (inset), with apparent second order rate constants of 16-19 μm-1 s-1 and 1-4 μm-1 s-1 (data not shown). More important, the total amplitudes of this pyrene fluorescence transient were nearly identical for equimolar active site concentrations of S1 and HMM preparations (Fig. 2B, inset). Finally, it should be noted that over the time scale where cross-linking occurs (>2 s after mixing with HMM), no further decrease in pyrene fluorescence was seen in the HMM transients. In fact, a small amplitude rising phase was noted, which fit a single exponential process at 0.05-0.07 s-1 (Fig. 2B, solid curve). As indicated by the discussion above, we interpret these results to mean that, after mixing, both heads of HMM bind relatively rapidly to the same actin filament. We predict that this would generate intramolecular strain, which would be relieved by a slow dissociation of the leading head from one actin filament and its subsequent rebinding to a neighboring filament. Labeling actin with pyrene reduces its affinity for myosin (14Taylor E.W. J. Biol. Chem. 1991; 266: 294-302Abstract Full Text PDF PubMed Google Scholar). Hence, the combination of rearward strain on the leading head, which would favor its dissociation from actin, with the increased affinity of an ADP-containing head for unlabeled actin, would together lead to rebinding to unlabeled actin subunits on neighboring filaments. This should lead to an overall reduction in pyrene actin subunits with a strongly bound HMM head, because in this experiment, only 50% of the actin subunits were labeled, and this would be manifested by a low amplitude rise in fluorescence at ∼0.05 s-1. This prediction is confirmed in Fig. 2B. Actin-activated ADP Release—Because ADP release is rate-limiting in the cycle (9De La Cruz E.M. Wells A.L. Rosenfeld S.S. Ostap E.M. Sweeney H.L. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 13726-13731Crossref PubMed Scopus (355) Google Scholar), we investigated the effect of strain on the kinetics of ADP release. This was monitored by mixing a complex of S1·2′dmD or HMM·2′dmD in the stopped flow with actin plus excess nucleotide. Fig. 3A illustrates the fluorescence transients produced by mixing a complex of 2 μm (active site concentration) S1·2′dmD (red) or HMM·2′dmD (green) with 20 μm actin plus 2 mm ADP. For S1, the transient could be fit to a single exponential decay, reflecting dissociation of the bound 2′dmD, and the rate constant of this process varied hyperbolically with actin concentration (Fig. 3A, inset, red circles), defining a maximum rate constant of 15.8 ± 1.6 s-1. This result is very similar to previously reported studies that used mant-ADP, which is a mixture of the 2′ and 3′ isomers (9De La Cruz E.M. Wells A.L. Rosenfeld S.S. Ostap E.M. Sweeney H.L. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 13726-13731Crossref PubMed Scopus (355) Google Scholar). By contrast, the corresponding fluorescence transient for HMM was clearly biphasic, and the two phases had similar amplitudes (Fig. 3B). Only the faster phase showed a dependence of rate constant on actin concentration (Fig. 3A, inset, green boxes), with an extrapolated maximum of 29.5 ± 3.4 s-1: nearly twice as large as the corresponding value for S1. The slower phase (inset, green triangles) averaged ≈0.3-0.4 s-1 and showed little variation with actin concentration. These results are consistent with the model illustrated in Fig. 3C, where the fluorescence of the bound 2′dmD is symbolized by the magenta rays emanating from the actin-bound heads. We propose that forward strain on the rear head accelerates ADP release by a factor of approximately two, whereas rearward strain on the forward head markedly slows ADP release, to ≈0.3 s-1. Release of this strain would occur with a slow dissociation of the forward head (at 0.03-0.05 s-1). This would then be rapidly followed by rebinding to a neighboring actin filament, which would likely occur with an altered geometry, one that would be unlikely to generate strain. We tested this proposal by mixing HMM·2′dmD or S1·2′dmD in the stopped flow with actin plus 2 mm ATP. Because S1 cannot generate internal strain, we would predict that the results with S1 in this experiment should be similar to those for the experiment discussed above. Fig. 4A confirms this. The fluorescence transient for S1 (red) is well fit by a single exponential decay, and the rate constant of the transient shows nearly an identical hyperbolic dependence on actin concentration (inset, red triangles, maximum rate constant 15.2 ± 1.0 s-1). Because we are proposing that forward strain accelerates ADP release from the trailing head, we would also expect that the fluorescence transient for HMM should be biphasic and that the maximum rate for the faster phase should be similar to that in Fig. 3A. The transient for HMM in this experiment (Fig. 4A, green) fits a double exponential decay, and the rate constant of the faster phase shows a hyperbolic dependence on actin concentration, which is very similar to that measured in the presence of ADP (inset, green boxes, maximum rate constant 28.9 ± 5.3 s-1). By contrast, the rate constant of the second phase for the HMM transient in this experiment was considerably faster, with a maximum of 8.8 ± 1.8 s-1 (inset, green triangles). As in the case of Fig. 3, the relative amplitudes of the two phases for HMM were similar (Fig. 4B). These results are consistent with the scheme depicted in Fig. 4C. Although excess ADP would keep HMM strongly bound and under strain, and while strain would inhibit 2′dmD release from the leading head, excess ATP would bind to the trailing head once it had released its 2′dmD. This would dissociate the trailing head, relieve the internal strain (symbolized by a serpentine regulatory domain) and allow 2′dmD to dissociate from the leading head. A maximum rate constant of 8.8 ± 1.8 s-1 (Fig. 4A, inset, green triangles) would be consistent with the model depicted in Fig. 4C if release of 2′dmD from the trailing head were immediately followed by ATP binding and dissociation from actin, allowing 2′dmD to release from the leading head at a rate defined by our studies with S1 (12-15 s-1; Ref. 9De La Cruz E.M. Wells A.L. Rosenfeld S.S. Ostap E.M. Sweeney H.L. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 13726-13731Crossref PubMed Scopus (355) Google Scholar and Fig. 4A). Kinetics of Phosphate and Product Release—We measured the kinetics of actin-activated phosphate release from myosin V S1 and HMM at equal active site concentrations in a sequential mixing experiment. Nucleotide-free S1 or HMM was mixed with a 10-fold molar excess of ATP, the complex was allowed to age for 1 s to allow population of the myosin V·ADP·Pi state, and it was then mixed with varying concentrations of actin plus 2 mm ADP. Phosphate release was monitored with MDCC-labeled phosphate-binding protein (MDCC-PBP), as previously described (10Rosenfeld S.S. Xing J. Chen L.-Q. Sweeney H.L. J. Biol. Chem. 2003; 278: 27449-27455Abstract Full Text Full Text PDF PubMed Scopus (91) Google Scholar), with a MDCC-PBP·myosin concentration ratio of 7:1 after the second mix. Fig. 5 illustrates an example of the resulting fluorescence transients for HMM (red) and S1 (black) at final active site concentrations of 0.5 μm and at a final actin concentration of 5 μm. For both constructs, the fluorescence fit a single exponential process (solid curves in Fig. 5). The S1·HMM amplitude ratio averaged 1.28 ± 0.18 over a range of actin concentrations (Fig. 5, inset). Finally, the rates of phosphate release from both S1 (Fig. 6B, closed red circles) and HMM (Fig. 7B, closed red circles) showed a similar hyperbolic dependence on actin concentration, defining maximum rates of 198 ± 18 s-1 for S1 and 228 ± 32 s-1 for HMM.Fig. 6Kinetics of product release for S1.A, fluorescence transient produced by mixing 4 μm S1·2′dmD·Pi with 20 μm actin plus 2 mm ATP. The resulting fluorescence decrease (jagged red curve) fit a double exponential function (solid black line). Inset: mixing 2 μm S1 with a 20 μm unlabeled ATP, followed by 20 μm actin plus 200 μm 2′dmT produced a monophasic rise in fluorescence. B, plot of rate versus actin concentration for product release reactions. S1 was mixed sequentially with a 10-fold excess of 2′dmT and then with actin plus 2 mm ATP, and the rates of both phases (closed blue boxes and open blue boxes) are plotted versus actin concentration. The rate constant of the faster phase varied hyperbolically with actin concentration with maximum rate of 200 ± 75 s-1, whereas the slower phase showed little actin concentration dependence and ranged between 10 and 14 s-1. The rate constant of the fluorescence rise illustrated in the inset (solid green circles) varied little with actin concentration, and its mean value is 10.2 ± 1.6 s-1. The rate of phosphate release, measured with MDCC-labeled phosphate binding protein, versus actin concentration is also shown (solid red circles), and a hyperbolic fit defines a maximum rate constant of 198 ± 18 s-1. C, fluorescence transients produced by mixing 7.5 μm nucleotide-free myosin V S1 in the stopped flow with 5 μm 2′dmT (red transient) or 2′dmD (green transient). For 2′dmT, an initial rapid rise in fluorescence is followed by a slow decay with rate constant of 0.11 s-1 (solid blue curve), whereas for 2′dmD, only a rapid rising phase is seen. Note that the final voltage for the transient with 2′dmT is nearly identical to that for 2′dmD. D, fluorescence transients produced by mixing 7.5 μm nucleotide-free myosin V S1 in the stopped flow with 37.5 μm 2′dmT (red transient) or 2′dmD (green transient). The amplitude of the transient with 2′dmT is 2.4-fold larger than that for 2′dmD.View Large Image Figure ViewerDownload Hi-res image Download (PPT)Fig. 7Kinetics of product release for HMM.A, fluorescence transient produced by mixing 2 μm HMM·2′dmD·Pi with 20 μm actin plus 2 mm ATP. The resulting fluorescence decrease (jagged red curve) could be fit a three exponential decay (solid black line). Inset: mixing 2 μm HMM with a 200 μm unlabeled ATP, followed by 100 μm actin plus 200 μm 2′dmT produced a biphasic rise in fluorescence with rates of 22.5 and 5.9 s-1. B, a plot of the ratio of the amplitude of the two slower phases versus actin concentration, demonstrating a similarity in the amplitudes of these two phases over a range of actin concentrations. C, the rates of the three phases produced by mixing HMM·2′dmD·Pi with actin plus 2 mm ATP are plotted as a function of actin concentration. The rates of the first two phases (closed blue boxes and open blue boxes) varied hyper" @default.
• W2055665417 created "2016-06-24" @default.
• W2055665417 creator A5002499186 @default.
• W2055665417 creator A5020641571 @default.
• W2055665417 date "2004-09-01" @default.
• W2055665417 modified "2023-10-11" @default.
• W2055665417 title "A Model of Myosin V Processivity" @default.
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