Identification of a strong binding site for kinesin on the microtubule using mutant analysis of tubulin
2006; Springer Nature; Volume: 25; Issue: 24 Linguagem: Inglês
10.1038/sj.emboj.7601442
ISSN1460-2075
AutoresSeiichi Uchimura, Yusuke Oguchi, Miho Katsuki, Takeo Usui, Hiroyuki Osada, Jun‐ichi Nikawa, Shin’ichi Ishiwata, Etsuko Muto,
Tópico(s)Photosynthetic Processes and Mechanisms
ResumoArticle23 November 2006free access Identification of a strong binding site for kinesin on the microtubule using mutant analysis of tubulin Seiichi Uchimura Seiichi Uchimura Brain Development Research Group, Brain Science Institute, RIKEN, Wako, Saitama, Japan Department of Bioscience and Bioinformatics, Faculty of Computer Science and Systems Engineering, Kyushu Institute of Technology, Fukuoka, Japan Search for more papers by this author Yusuke Oguchi Yusuke Oguchi Department of Physics, School of Science and Engineering, Waseda University, Tokyo, Japan Search for more papers by this author Miho Katsuki Miho Katsuki Brain Development Research Group, Brain Science Institute, RIKEN, Wako, Saitama, JapanPresent address: Molecular Motors Group, Marie Curie Research Institute, The Chart, Oxted, Surrey RH8 0TE, UK. Search for more papers by this author Takeo Usui Takeo Usui Antibiotics Laboratory, Discovery Research Institute, RIKEN, Wako, Saitama, JapanPresent address: Graduate School of Life and Environmental Sciences, University of Tsukuba, Tsukuba, Ibaraki, Japan Search for more papers by this author Hiroyuki Osada Hiroyuki Osada Antibiotics Laboratory, Discovery Research Institute, RIKEN, Wako, Saitama, Japan Search for more papers by this author Jun-ichi Nikawa Jun-ichi Nikawa Department of Bioscience and Bioinformatics, Faculty of Computer Science and Systems Engineering, Kyushu Institute of Technology, Fukuoka, Japan Search for more papers by this author Shin'ichi Ishiwata Shin'ichi Ishiwata Department of Physics, School of Science and Engineering, Waseda University, Tokyo, Japan Advanced Research Institute for Science and Engineering, Waseda University, Tokyo, Japan Search for more papers by this author Etsuko Muto Corresponding Author Etsuko Muto Brain Development Research Group, Brain Science Institute, RIKEN, Wako, Saitama, Japan Search for more papers by this author Seiichi Uchimura Seiichi Uchimura Brain Development Research Group, Brain Science Institute, RIKEN, Wako, Saitama, Japan Department of Bioscience and Bioinformatics, Faculty of Computer Science and Systems Engineering, Kyushu Institute of Technology, Fukuoka, Japan Search for more papers by this author Yusuke Oguchi Yusuke Oguchi Department of Physics, School of Science and Engineering, Waseda University, Tokyo, Japan Search for more papers by this author Miho Katsuki Miho Katsuki Brain Development Research Group, Brain Science Institute, RIKEN, Wako, Saitama, JapanPresent address: Molecular Motors Group, Marie Curie Research Institute, The Chart, Oxted, Surrey RH8 0TE, UK. Search for more papers by this author Takeo Usui Takeo Usui Antibiotics Laboratory, Discovery Research Institute, RIKEN, Wako, Saitama, JapanPresent address: Graduate School of Life and Environmental Sciences, University of Tsukuba, Tsukuba, Ibaraki, Japan Search for more papers by this author Hiroyuki Osada Hiroyuki Osada Antibiotics Laboratory, Discovery Research Institute, RIKEN, Wako, Saitama, Japan Search for more papers by this author Jun-ichi Nikawa Jun-ichi Nikawa Department of Bioscience and Bioinformatics, Faculty of Computer Science and Systems Engineering, Kyushu Institute of Technology, Fukuoka, Japan Search for more papers by this author Shin'ichi Ishiwata Shin'ichi Ishiwata Department of Physics, School of Science and Engineering, Waseda University, Tokyo, Japan Advanced Research Institute for Science and Engineering, Waseda University, Tokyo, Japan Search for more papers by this author Etsuko Muto Corresponding Author Etsuko Muto Brain Development Research Group, Brain Science Institute, RIKEN, Wako, Saitama, Japan Search for more papers by this author Author Information Seiichi Uchimura1,4,‡, Yusuke Oguchi2,‡, Miho Katsuki1,‡, Takeo Usui3, Hiroyuki Osada3, Jun-ichi Nikawa4, Shin'ichi Ishiwata2,5 and Etsuko Muto 1 1Brain Development Research Group, Brain Science Institute, RIKEN, Wako, Saitama, Japan 2Department of Physics, School of Science and Engineering, Waseda University, Tokyo, Japan 3Antibiotics Laboratory, Discovery Research Institute, RIKEN, Wako, Saitama, Japan 4Department of Bioscience and Bioinformatics, Faculty of Computer Science and Systems Engineering, Kyushu Institute of Technology, Fukuoka, Japan 5Advanced Research Institute for Science and Engineering, Waseda University, Tokyo, Japan ‡These authors contributed equally to this work *Corresponding author. Brain Development Research Group, Brain Science Institute, RIKEN, Hirosawa 2-1, Wako, Saitama 351-0198, Japan. Tel.: +81 48 467 6959; Fax: +81 48 467 7145; E-mail: [email protected] The EMBO Journal (2006)25:5932-5941https://doi.org/10.1038/sj.emboj.7601442 Present address: Molecular Motors Group, Marie Curie Research Institute, The Chart, Oxted, Surrey RH8 0TE, UK. PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The kinesin-binding site on the microtubule has not been identified because of the technical difficulties involved in the mutant analyses of tubulin. Exploiting the budding yeast expression system, we succeeded in replacing the negatively charged residues in the α-helix 12 of β-tubulin with alanine and analyzed their effect on kinesin-microtubule interaction in vitro. The microtubule gliding assay showed that the affinity of the microtubules for kinesin was significantly reduced in E410A, D417A, and E421A, but not in E412A mutant. The unbinding force measurement revealed that in the former three mutants, the kinesin-microtubule interaction in the adenosine 5′-[β,γ-imido]triphosphate state (AMP-PNP state) became less stable when a load was imposed towards the microtubule minus end. In parallel with this decreased stability, the stall force of kinesin was reduced. Our results implicate residues E410, D417, and E421 as crucial for the kinesin-microtubule interaction in the strong binding state, thereby governing the size of kinesin stall force. Introduction Kinesin is a molecular motor involved in many cellular force-generating processes such as organelle transport and chromosome segregation (Vernos and Karsenti, 1996; Goldstein and Yang, 2000). Conventional kinesin contains two identical heavy chains and can move processively along a microtubule more than 1 μm in length without dissociation (Howard et al, 1989; Block et al, 1990). Processive movement of kinesin is explained by the hand-over-hand model, in which the motor maintains continuous contact with the microtubule as a result of alternating head catalysis of ATP (Hackney, 1994; Ma and Taylor, 1997). At each ATP hydrolysis cycle, kinesin makes an 8-nm step (equal to the size of tubulin dimer) towards the microtubule plus end (Hua et al, 1997; Schnitzer and Block, 1997), and this stepping motion is triggered by a conformational change in the ATP-bound head (Rice et al, 1999). Docking of the crystal structure of kinesin into cryoelectron-microscopy maps of kinesin-microtubule complex indicated L8, L11, and α4/L12/α5 in the motor domain of kinesin as the structural key elements for microtubule binding in the presence of AMP-PNP, a non-hydrolyzable ATP analogue mimicking the ATP-bound state (Hirose et al, 1999; Hoenger et al, 2000; Kikkawa et al, 2000; Skiniotis et al, 2004). In this model, L8 may bind to α-helix 12 (H12) in β-tubulin, and L11 extends towards the H11–H12 loop in α-tubulin. α4/L12/α5 is also closely associated with H12 in β-tubulin and possibly with the C-terminus of β-tubulin, which is not defined in the crystal structure. The model indicates that kinesin-microtubule interaction might be mediated by electrostatic interactions. Some of the positively charged amino acids in these contact areas of kinesin potentially interact with the negatively charged surface of the microtubule, which is mainly composed of H12 in β-tubulin. Consistent with this assumption, using alanine-scanning mutagenesis of kinesin, several positively charged residues in L7/8, L11, and α4/L12/α5 have been identified as microtubule-interacting kinesin residues (Woehlke et al, 1997). However, the critical residues on tubulin have not been identified. Proteolytic digestion of the C-terminal region of tubulin led to the reduced processivity of both single- and double-headed kinesin (Okada and Hirokawa, 2000; Thorn et al, 2000; Wang and Sheetz, 2000; Lakämper and Meyhöfer, 2005). The biochemical measurement of Kd of kinesin to both intact and protease-digested microtubule revealed that the negatively charged C-terminus of tubulin may be a binding partner for kinesin in the weak binding state (ADP state), but not in the strong binding state (AMP-PNP state) (Okada and Hirokawa, 2000; Skiniotis et al, 2004; Lakämper and Meyhöfer, 2005). A structural element other than the tubulin C-terminal region may serve as an interface that is specific for the strong binding state; the structural analyses have implicated H12 in β-tubulin as a potential candidate for this (Hirose et al, 1999; Hoenger et al, 2000; Kikkawa et al, 2000; Skiniotis et al, 2004). For elucidating the mechanism of kinesin motility, it is essential to identify the structural elements involved in each chemical state because the physicochemical properties inherent to each chemical state might be critically dependent on the interface configuration and the surface force type that works between the interfaces (Israelachvili, 1992; Okada and Hirokawa, 2000; Skiniotis et al, 2004; Lakämper and Meyhöfer, 2005). Our lack of knowledge about kinesin interface on tubulin is attributed to the technical difficulties involved in mutant analysis of tubulin. Tubulin is a heterodimer composed of α- and β-polypeptides, each requiring a distinct set of chaperons for proper folding. Apart from this complexity, multiple α- and β-tubulin genes are found in most eukaryotic cells, with each subunit undergoing different post-translational modifications (Luduena, 1998). Consequently, it has been difficult to express and purify isotypically pure tubulin in biochemically useful amounts. Here, we have used the budding yeast Saccharomyces cerevisiae to conduct mutant analyses on microtubules. S. cerevisiae contains only two α-tubulin genes (TUB1 and TUB3) and one β-tubulin gene (TUB2); since TUB3 is nonessential, it provides a potential source of isotypically pure tubulin (Bode et al, 2003). Using TUB3-null cells, we succeeded in generating a set of charged-to-alanine point mutations in the sequence coding for H12 in β-tubulin. Each of these mutated tubulins was purified from the cell lysate, polymerized into microtubules, and the effect of mutation was examined in microtubule gliding assay on conventional two-headed kinesin. This showed that the affinity of microtubule for kinesin was reduced in E410A, D417A, and E421A mutants. The measurement of the unbinding force (Kawaguchi and Ishiwata, 2001; Uemura et al, 2002) revealed that in the former three mutants, the kinesin-microtubule interaction in AMP-PNP state became less stable for minus end loading, whereas their interaction in ADP state was unaffected by the mutations. These results indicate that the negatively charged amino-acid residues E410, D417, and E421 in β-tubulin are crucial for the strong binding of kinesin to the microtubule. Hereafter, we have abbreviated the α-helix and β-sheet in the tubulin structures as 'H' and 'S,' corresponding to α and β, respectively, in the kinesin structure, according to the original paper on the structure of microtubule (Nogales et al, 1998). Results Construction, expression, and purification of H12 mutated microtubules To prepare isotypically pure tubulin, we used TUB3-null yeast cells, having only a single α- and β-tubulin gene, TUB1 and TUB2, respectively (see Supplementary Methods and Table SII). As the aim of this study was to examine the motility of kinesin along the mutated microtubules, it was necessary to obtain yeast microtubules that were stable at low concentrations required for in vitro motility assay (∼10 μg/ml). Hence, Taxol-binding ability was introduced into a β-tubulin gene by site-directed mutagenesis at five amino acids (Gupta et al, 2003), and the gene tub2-A19K-T23V-G26D-N227H-Y270F thus obtained was referred to as TUB2tax. The yeast strain expressing TUB1 and TUB2tax was used as the wild-type in the following analyses. To examine the function of β-tubulin H12 in kinesin motility, we generated a set of charged-to-alanine point mutations in the sequence coding for H12 in β-tubulin (Figure 1). Among these four mutants, E410A and D417A were haploid lethal. Therefore, we attempted to isolate these mutated tubulins by expressing two species of tub2tax genes in a strain, one of which, tub2tax-plusE (tub2tax-440GDFGEEEEGEEEEGEEEEGEEEEGEEEEA), contains numerous negatively charged amino acids at the C-terminus and the other, tub2tax-E410A or tub2tax-D417A, which has point mutation in H12; the latter is under the control of the inducible galactose promoter (Burke et al, 1989). The haploid cells were first grown in YPD medium, expressing only tub2tax-plusE. When the growth reached the late-log phase, the cells were transferred to YPG medium containing galactose for inducing the coexpression of tub2tax-E410A (or tub2tax-D417A). The cells were cultured for another 8 h in YPG medium, and then harvested for tubulin purification. The cell lysate contains two species of tubulin dimers, including either Tub2taxp-plusE or Tub2taxp-E410A (or Tub2taxp-D417A) as β-tubulin subunit, but these two species can be separated using their charge differences during purification (see Supplementary Figure S1). For the other two mutants, E412A and E421A, tubulin could be readily purified from the cultured haploid cells as these cells containing only this mutated tubulin were viable under normal growth conditions (YPD medium). Figure 1.Design of the H12 mutants of β-tubulin in Saccharomyces cerevisiae. (A) Sequences of the H12 region are shown with negatively charged residues indicated by blue and the residues substituted by alanines are indicated by red. (B) A ribbon diagram of the tubulin dimer viewed from the side of the microtubule with its minus end to the left (Nogales et al, 1998). Image analysis of the kinesin-microtubule complex revealed that in both nucleotide free and AMP-PNP state, kinesin motor domain is associated in close proximity to H11 (orange), H12 (cyan), and the COOH terminus (undefined in crystal structure) of β-tubulin (Kikkawa et al, 2000; Hoenger et al, 2000). The acidic residues in H12 mutagenized to alanine are indicated in blue. Download figure Download PowerPoint To purify tubulin from the cell lysate, a couple of anion exchange column chromatographies were successively used (Davis et al, 1993), and the crude tubulin fraction eluted from the second column was further purified by polymerization and depolymerization (see Supplementary Methods and Figure S2). This procedure generated approximately 30 μg of assembly-competent tubulin from 6L of culture with purity higher than 95% on SDS gel electrophoresis (Figure 2A). To examine if these mutated tubulins were post-translationally modified at their C-terminus (Luduena, 1998), both α- and β-polypeptides were analyzed by electrospray ionization/ion trap and quadrupole-TOF mass spectrometry. The MS and MS/MS data suggested that the isolated tubulin dimers predominantly contained α- and β-polypeptides lacking any post-translational modifications at the C-terminal region (Supplementary Figure S3). Figure 2.Purity of yeast tubulin and images of polymerized microtubules. (A) SDS-PAGE analysis of purified tubulin. Lane 1, yeast wild-type tubulin; lane 2, E410A; lane 3, E412A; lane 4, D417A; lane 5, E421A; and lane 6, porcine brain tubulin. In each lane, 1 μg of sample was loaded and stained by Coomassie blue. In SDS gel containing Sigma SDS (L-5750), α- and β-polypeptide of porcine brain tubulin were separated more as compared to these peptides purified from yeast cells (Best et al, 1981; Bode et al, 2003). (B) Dark-field images of the microtubules polymerized from yeast wild-type and mutated tubulins in the presence of 1 μM Taxol. Bar=5 μm. Download figure Download PowerPoint When these isolated tubulins (0.5–1.0 mg/ml) were incubated at 30°C in the presence of 1 μM Taxol, all species of tubulins could polymerize into long filaments having length up to ∼20 μm (Figure 2B). In vitro motility assay To examine the influence of the mutations on kinesin-microtubule interaction, these mutated microtubules were first tested in the microtubule gliding assay on the conventional two-headed kinesin. When the density of kinesin on the glass surfaces was >1000 μm−2, both wild-type and mutated microtubules showed smooth gliding movement at similar velocities (Table I). When the direction of the movement was examined using polarity marked microtubules (Tanaka-Takiguchi et al, 1998), the movement of kinesin was plus-end-directed in all cases. However, at kinesin density of ∼100 μm−2, the wild-type and the E412A microtubules showed smooth gliding movement, whereas the E410A, D417A, and E421A microtubules often rotated erratically about a roughly vertical axis, resulting in wobbly gliding motions. This indicates that the latter three mutants may have lower affinity for kinesin as compared to the wild-type. Table 1. Velocity of microtubule movement in the gliding assaya Construct Velocity (μm/s)b n WT 0.75±0.08 120 E410A 0.78±0.06 120 E412A 0.76±0.06 103 D417A 0.73±0.08 123 E421A 0.74±0.10 102 a Gliding assay was performed at a kinesin density of ∼2000/μm2. b Mean±s.e.m. To quantify the affinity of these mutated microtubules for kinesin, we measured the fraction of the microtubules that moved a distance greater than their own lengths, f, over a range of kinesin densities (Figure 3A and B; Howard et al, 1989). Here, to compare between each mutant and wild-type, the length of the microtubules was adjusted to approximately 3 μm in all strains (see Materials and methods). The result showed that in the wild-type, virtually all the microtubules moved more than their own length at kinesin density >300 μm−2, and the fraction of the microtubules that covered distances greater than their own length gradually decreased with the decrease in kinesin density. At kinesin density 1. This result may indicate that either a simultaneous, collective action of multiple motor molecules is required for the stable gliding movement of mutated microtubules (Hancock and Howard, 1998; Shima et al, 2006) or that a run length of a single kinesin along the mutated microtubule is too short to support the gliding movement of microtubule over a distance greater than its own length. To understand which the operative mechanism is, we conducted a single molecule motility assay (Figure 3C); the motility of fluorescently labeled two-headed kinesin HK560-Cy3 (0.29 nM) was examined along these mutated microtubules using total internal reflection fluorescence microscopy (TIRFM). The result showed that whereas kinesin moved processively along the wild-type and E412A microtubules, it scarcely interacted with the E410A and D417A microtubules (Figure 3D). For E421A, kinesin could move along the microtubule, but its processivity was reduced as compared to that of the wild-type. The mean run length of kinesin for the wild-type, E412A, and E421A microtubules was 0.843±0.004 μm, 0.837±0.006 μm, and 0.663±0.004 μm, respectively. These results are consistent with the values of ρ0 and n derived from the fraction curves in Figure 3B. The estimated value of n=3 for E421A was due to the reduced processivity of single kinesin. It is reasonable that only the E410A, D417A, and E421A mutations affected the kinesin-microtubule interaction. Docking experiments of the crystal structure of kinesin into cryoelectron-microscopy maps of kinesin-microtubule complex revealed that H12 of β-tubulin is located on the surface of microtubules and aligned diagonally to the longitudinal axis of protofilament (Hoenger et al, 2000; Kikkawa et al, 2000). In its coiled structure, the residues E410, D417, and E421 are facing kinesin, whereas E412 is positioned on the opposite side, facing the microtubule core. Unbinding force To clarify the chemical state at which the binding affinity of the microtubules for kinesin was modulated in the mutants, we measured the force required to dissociate kinesin from these mutated microtubules (unbinding force) under two nucleotide conditions—in the presence of ADP and AMP-PNP (Kawaguchi and Ishiwata, 2001; Uemura et al, 2002). One-headed kinesin heterodimers were used for the measurement because our previous work demonstrated that the two distinct binding modes (weak and strong), each inherent to the nucleotide condition, can be unambiguously characterized with the single-headed kinesin (Uemura et al, 2002). Using optical tweezers, a polystyrene bead attached with a single-headed kinesin (Kojima et al, 1997) was made to interact with a microtubule in the presence of either 1 mM ADP or 1 mM AMP-PNP. An external load was gradually applied to the kinesin-microtubule complex by moving the stage of the microscope towards the plus end or the minus end of the microtubule until the bead dissociated from it. The unbinding force was calculated by multiplying the magnitude of abrupt bead displacement during detachment with the stiffness of the optical tweezers. The yeast wild-type microtubules showed properties similar to those observed previously for brain microtubules (Figure 4; Kawaguchi and Ishiwata, 2001; Uemura et al, 2002). Unbinding force in the AMP-PNP state was significantly higher than that in the ADP state, and in each nucleotide state, the unbinding force for the minus-end loading was higher than that for the plus-end loading. The dependence on loading direction in the ADP state, however, was not as significant as in the AMP-PNP state. Figure 4.Unbinding force distribution of one-headed kinesin in (A) ADP and (B) AMP-PNP state. An external load was applied towards either the plus end (orange for (A) and red for (B)) or the minus end of the microtubule (light and dark green for (A) and (B), respectively). The stiffness of the trap was 0.038 pN/nm (ADP) and 0.076 pN/nm (AMP-PNP). The average unbinding force (pN) with s.e.m. is shown in each panel. Download figure Download PowerPoint Whereas the unbinding force in ADP state for mutated microtubules was almost similar to that measured for wild-type (Figure 4A), the unbinding force in AMP-PNP state was significantly altered by mutations (Figure 4B). In the presence of AMP-PNP, the unbinding force for minus-end loading was considerably reduced in all three mutants (7.8±0.4, 5.3±0.3, 3.8±0.2, and 3.6±0.2 pN for wild-type, E410A, D417A, and E421A, respectively; mean±s.e.m.), yet the unbinding force for plus-end loading was scarcely affected (5.7±0.3, 5.2±0.3, 6.2±0.4, and 5.6±0.3 pN for wild-type, E410A, D417A, and E421A, respectively). Unexpectedly, the mutations rendered the kinesin-microtubule interaction less stable only for minus-end loading. In the presence of ADP, the unbinding force measured for mutants was fairly similar to that of the wild-type except in the case of E421A, where the asymmetry for loading direction was reversed. The residues E410, D417, and E421 appeared to be crucial for the strong binding of kinesin to the microtubules, and this may underlie the reduced affinity of these mutated microtubules for kinesin during movement (Figure 3). Stall force Previous measurements of the mechanical properties of two-headed kinesin indicated that the kinesin stall force might be determined by (1) the binding affinity of the head to the microtubule in the strongly bound state and (2) the ATP-binding kinetics to the nucleotide-free head (Visscher et al, 1999; Nishiyama et al, 2002; Lakämper and Meyhöfer, 2005; Shao and Gao, 2006). Thus, the kinesin stall force exerted on the mutated microtubules with reduced affinity for kinesin in AMP-PNP state is expected to be smaller than that exerted on the wild-type microtubules. To examine this hypothesis, we next attempted to measure the stall force of conventional two-headed kinesin along these mutated microtubules. When the displacements and forces caused by single kinesin molecules were measured in the presence of 1 mM ATP using optical tweezers (trap stiffness=0.076 pN/nm), the stall force measured for E410A, D417A, and E421A was 3.5±0.1, 3.0±0.1, and 3.1±0.1 pN, respectively (mean±s.e.m.). These values were significantly lower than that measured for the wild-type (5.6±0.2 pN) (Figure 5A and B). Although we observed the processive movement of kinesin only for the wild-type, E412A, and E421A microtubules in the single molecule motility assay (Figure 3D), in the experiment using optical tweezers, a single-kinesin-bound bead is also observed to move along the E410A and D417A microtubules, probably because their initial interaction/successive interaction with the microtubule was enforced by the laser trap. Figure 5.Stall force measurement. (A) Representative tracing of a trapped bead powered by a conventional two-headed kinesin along the wild-type, E410A, D417A, and E421A microtubules, measured at the trap stiffness of 0.076 pN/nm. Light shaded, unfiltered; solid, filtered at 100 Hz. (B) Distribution of the stall force for wild-type and mutated microtubules. The average stall force with s.e.m. is shown in each panel. Total number of events counted are (from top to bottom) 61, 104, 102, and 69, respectively. (C) The stall force plotted against the unbinding force for minus- (left) and plus-end loading (right). The stall force was linearly related
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