Configuration of the two kinesin motor domains during ATP hydrolysis

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1 Configuration of the two kinesin motor domains during ATP hydrolysis Ana B Asenjo, Natan Krohn & Hernando Sosa To understand the mechanism of kinesin movement we have investigated the relative configuration of the two kinesin motor domains during ATP hydrolysis using fluorescence polarization microscopy of ensemble and single molecules. We found that: (i) in nucleotide states that induce strong microtubule binding, both motor domains are bound to the microtubule with similar orientations; (ii) this orientation is maintained during processive motion in the presence of ATP; (iii) the neck-linker region of the motor domain has distinct configurations for each nucleotide condition tested. Our results fit well with a hand-over-hand type movement mechanism and suggest how the ATPase cycle in the two motor domains is coordinated. We propose that the motor neck-linker domain configuration controls ADP release. Kinesin, a cellular motor protein involved in intracellular transport, uses the energy of ATP hydrolysis to move along microtubules 1. Conventional kinesin has two identical motor domains and is a highly processive motor that takes hundreds of steps before dissociating from the microtubule 2. Processivity is thought to depend on the coordination between the ATPase cycles of both motor domains 3. There is, however, very limited information regarding the relative configurations of the two motors during ATP hydrolysis. Mechanical experiments have indicated that they both can bind to the microtubule 4, but previous structural studies have given conflicting results regarding their relative configuration 5 7. Likely causes for the reported discrepancies are artifacts caused by saturating the microtubule lattice with kinesin molecules and the averaging procedures used to obtain the reported structures. Here we used an alternative approach, fluorescence polarization microscopy 8,9,to investigate this issue. The method does not require saturation of the microtubule lattice and can also give information at the single molecule level on actively moving motor proteins. RESULTS Fluorescent probe locations For this study, five cysteine-light (having only two cysteines) kinesin motor domain constructs were fluorescently labeled at three alternative locations (Fig. 1). The only difference between the constructs was in the location of the unique cysteines targeted for labeling and whether part of the dimerization domain was included or not (see Methods). We called these constructs KM_A, KM_B, KM_C, KD_A and KD_B where M or D denotes whether the construct is monomeric or dimeric and the last letter (A, B, C) indicates the probe location. The monomeric constructs provide information on the configuration of a single motor domain attached to the microtubule. The dimeric constructs, in addition, provide information on the extra motor domain present in the dimer. Two probe locations, A and B (Fig. 1a,b), are on opposite sides of the kinesin motor domain in regions not reported to undergo major conformational changes relative to the rest of the motor. Location C (Fig. 1c) is in the so-called neck-linker region. This region has been reported to undergo a disorder-to-order transition upon ATP binding when the motor is attached to the microtubule 10. We verified that in these kinesin constructs motor activity was not impaired. All the labeled constructs showed microtubule-activated ATPase activity (V max = and ATP s 1 per motor head for KM and KD constructs, respectively). In single molecule fluorescence motility assays, the dimeric constructs KD_A and KD_B moved processively along axonemal microtubules in the presence of ATP with mean run lengths of 0.8 µm (KD_A) and 1.2 µm (KD_B) and speeds of 709 ± 112 nm s 1 (KD_A) and 620 ± 165 nm s 1 (KD_B), respectively. Single motor orientation in the presence of nucleotide analogs We used fluorescence anisotropy to measure the orientation and mobility of the labeled kinesin constructs bound to axonemal microtubules. Four equilibrium conditions were used to investigate different points along the ATPase hydrolysis cycle: (i) in the presence of AMPPNP (adenosine 5-(β,γ-imido)triphosphate, a nonhydrolyzable ATP analog mimicking the ATP bound state); (ii) in the absence of nucleotides (NN, no nucleotides added and in the presence of apyrase); (iii) in the presence of ADP+AlF 4 (mimicking the ADP-Pi state 11 ); (iv) in the presence of ADP. In the AMPPNP, NN and ADP+AlF 4 states, the microtubules were very well decorated with the kinesin constructs. In fluorescent images the microtubules look uniformly bright along their length over a relatively dark background (Fig. 2a). In comparison, in the ADP state the fluorescence intensity of Department of Physiology and Biophysics, Albert Einstein College of Medicine, Bronx, New York 10461, USA. Correspondence should be addressed to H.S. (hsosa@aecom.yu.edu). Published online 14 September 2003; doi: /nsb VOLUME 10 NUMBER 10 OCTOBER 2003 NATURE STRUCTURAL BIOLOGY

2 a b c Figure 1 Probe location and orientation. The bifunctional fluorescence probe BSR was targeted to three alternative locations on the kinesin motor domain by the engineering of two unique cysteines in the motor domain of kinesin (see Methods). The double attachment of the bifunctional probe to the protein reduces probe mobility and orients the transition dipole parallel to the line between the attachment points 8,9,28. (a) Probe position A. (b) Probe position B. (c) Probe position C. The locations of the Cβ carbons of the unique cysteine are indicated by spheres and a line connecting them (A, red; B, blue; C, green). Bound ADP is yellow and the neck-linker region orange. The structures shown correspond to the rat kinesin motor domain crystal structure 15. In A, B and C the motor domain is oriented relative to the microtubule long axis so that the probes at positions A and B have axial angles of 71 and 28, respectively, matching the values calculated experimentally for KM_A and KM_B in the AMPPNP state (Table 1). the microtubules relative to the background was lower (data not shown), indicating a weakly attached state 12. Therefore, we will refer collectively to the AMPPNP, NN and ADP+AlF 4 conditions as the strongly bound (to the microtubule) states. In these states the labeled microtubules show strong anisotropy (Fig. 2a). Emitted fluorescence intensity changes considerably depending on the relative angle between the microtubule long axis and the direction of polarization of the excitation light. This anisotropy was expressed as a linear dichroism ratio 8 (LD), defined as the difference between the fluorescence intensities corresponding to two perpendicular polarization excitations divided by the sum of the intensities (see Methods). For an ensemble of fluorophores with cylindrical symmetry (such as the one formed by labeled kinesin molecules attached to a microtubule), the LD value depends on the orientation of the microtubule relative to the polarization excitation axes, the fluorophore absorption dipole orientation relative to the microtubule (β) and the extent of probe disorder 8,9 (Γ) (Fig. 2b). We measured the LD of many individual kinesin decorated microtubules in random orientations on the microscope stage plane (x-y plane) for each kinesin construct and nucleotide condition. Selected LD versus microtubule angle plots (KM_A, KM_B and KM_C in the absence of nucleotides) are shown (Fig. 2c). The three examples presented (Fig. 2c) illustrate the range of fluorophore dipole orientations relative to the microtubule: near perpendicular (KM_A), near parallel (KM_B) and disordered or near 54.7 (KM_C). The data for each experimental condition can be summarized by calculating the extrapolated LD value when ω = 0, LD 0. LD 0 can have values between 1 and 1 depending on the orientation of the fluorescent absorption dipole relative to the microtubule axis. LD 0 > 0 for near perpendicular orientations (β > 54.7 ). LD 0 < 0 for near parallel orientations (β < 54.7 ) and LD 0 = 0 if β = 54.7 or if the probes are fully disordered 8 (see equation (1) in Methods). In the strongly bound states, the LD 0 values of KM_A are from +0.4 to +0.5 but are closer to 0 (+0.04 to +0.08) in the presence of ADP (Table 1 and Fig. 2d). In the case of KM_B the LD 0 values for the strongly bound states are from 0.5 to 0.6 and also close to 0 for the ADP state ( 0.02 to 0.05). These results confirm a previous study with a monomeric kinesin construct indicating a disordered state in the presence of ADP (LD 0 0) and an ordered configuration for the strongly bound states induced by AMPPNP, NN or ADP+AlF 4 (ref. 9). The previous study was done with the probe in location A only. Thus, our results with the alternate probe location in KM_B indicate that the disordered configuration induced by ADP is not restricted to particular probe locations, but instead involves the whole motor domain as proposed 9. In the strongly bound states, the LD 0 values of the constructs with probes at positions A and B (Table 1 and Fig. 2d,e) have opposite signs, indicating that the fluorophores dipoles are near perpendicular to each other in the two positions. This result agrees well with the relative orientation of the probes in the two locations (70 ) predicted from the kinesin crystal structure (Fig. 1a,b). Therefore, to a first approximation, locations A and B report on the orientation of the whole motor domain as a rigid body. The calculated axial angles (β) for KM_A and KM_B (Table 1) in the strongly bound states orient the motor domain relative to the microtubule as shown in Figure 1. This orientation is very close to the one predicted from cryo-electron microscopy docking models for kinesin motor domains bound to the microtubule Table 1 Summary of ensemble fluorescence anisotropy results Protein Nucleotide LD 0 Avg. dev. n Axial angle (β) KM_A ADP 0.04 (0.08) (57) a NN 0.45 (0.51) (73) AMPPNP 0.36 (0.46) (71) ADP+AlF (0.52) (73) KM_B ADP 0.02 ( 0.05) (53) a NN 0.55 ( 0.63) (28) AMPPNP 0.53 ( 0.62) (28) ADP+AlF ( 0.57) (31) KM_C ADP 0.03 (0.07) (57) a NN 0.02 ( 0.05) (53) a AMPPNP 0.25 ( 0.34) (42) ADP+AlF ( 0.54) (33) KD_A ADP 0.18 (0.30) (65) NN 0.43 (0.51) (73) AMPPNP 0.41 (0.49) (72) ADP+AlF (0.54) (74) KD_B ADP 0.26 ( 0.38) (40) NN 0.53 ( 0.63) (28) AMPPNP 0.56 ( 0.66) (26) ADP+AlF ( 0.66) (26) a Angular values close to 54.7 correspond to LD values near 0. LD values near 0 can be the result of axial angles close to the magic angle 54.7 or the result of angular disorder 8. LD 0, LD when the microtubule is parallel to the x-axis (ω = 0). Axial angle β, axial angle of the probe dipole with the microtubule long axis (Fig. 2b). LD 0 and β average values (outside parentheses) were calculated by a least-square fitting of equation (1) to the LD vs. ω data (Fig. 2c) corresponding to each experimental condition. n, number of data points; avg. dev., average deviation of the experimental data points to the fitted curve. The LD 0 values in parentheses are equal to the LD 0 average value + 1 average deviation with the same sign as the average LD 0. This corrected LD 0 value is used to account for the fact that many non-ideal effects would tend to reduce the LD values from the ideal ones if perfect cylindrical symmetry were preserved in all the axonemes 8,9. The β angles calculated with the corrected LD 0 are also shown in parentheses. NATURE STRUCTURAL BIOLOGY VOLUME 10 NUMBER 10 OCTOBER

3 a b d c Figure 2 Fluorescence polarization anisotropy of axonemal microtubules decorated with many kinesin molecules. (a) Fluorescence anisotropy example of a microtubule decorated with a BSR-labeled kinesin construct (KD_B in the presence of ADP+AlF 4 ). The green arrow in each panel indicates the polarization direction of the excitation light. (b) Definition of angles and axes. Microtubules lay flat on the x-y plane (microscope stage) at a random angle ω with the x-axis. A dipole bound to the microtubule can be described by its axial angle β with the microtubule axis and a mobility cone angle Γ 8. (c) LD versus microtubule angle (ω) for the three probe locations on the monomeric constructs (KM_A, KM_B, KM_C) in the absence of nucleotides (NN). LD is defined as (I 90 I 0 ) / (I 90 + I 0 ) where I x is fluorescence intensity and x is the polarization direction angle of the excitation light. (d) Bar plot of the calculated LD 0 for all monomeric constructs (KM_A, KM_B and KM_C) and nucleotide conditions studied. The average deviation of the data points from the fitted curve is indicated by the error T-bars. (e) Bar plot of the LD 0 corresponding to the dimeric constructs (KD_A and KD_B). e in the presence of AMPPNP 7,10,13,14. Such docking models predict axial angles of for location A and 7 21 for location B. With the orientation of the motor domain as in Figure 1 and the neck linker docked onto the motor domain as in the crystal structure 15 shown, location C would be 13 from the microtubule axis. The discrepancy between this angular value and those estimated for KM_C in the presence of AMPPNP or ADP+AlF 4 (42 and 33, respectively; Table 1) is likely because this part of the motor domain is more mobile 10. Mobility (angular disorder within the time scale of the measurements) results in LD 0 values closer to 0 and calculated β angles closer to In the absence of nucleotides, KM_C shows a very different result when compared with KM_A and KM_B. KM_A and KM_B have high LD 0 values ( ) whereas the corresponding value for KM_C is near 0 ( ) (Table 1 and Fig. 2c,d). Therefore, location C (the neck linker) in the absence of nucleotides is very mobile (or has an angle of 54.7 relative to the microtubule) whereas the rest of the motor domain keeps an orientation relative to the microtubule as shown in Figure 1. In the presence of AMPPNP the LD 0 of KM_C is 0.25, indicating a neck linker more ordered and more parallel to the microtubule. These results are consistent with a proposed transition of the neck linker from disorder (in the absence of nucleotides) to order induced upon ATP binding 10. We also observe an additional conformational change in the ADP+AlF 4 state. In the presence of ADP+AlF 4 the LD 0 value of KM_C is 0.5 indicating an orientation even more parallel to the microtubule axis than in the AMPPNP case. Also, there is a small but significant difference (P < 0.01) between the LD 0 values of the NN and ADP states (Table 1 and Fig. 2d). Our data indicate a continuous transition from the ADP state to the ADP+AlF 4 state, in which the neck linker goes from a disordered, or slightly perpendicular orientation relative to the microtubule, to a more parallel one. Thus, the neck linker is sensitive to all possible nucleotide species in the kinesin active site. Dimer motor orientation in the presence of nucleotide analogs The dimeric constructs (KD_A and KD_B) in the presence of ADP show higher LD 0 values than the monomeric constructs (Table 1 and Fig. 2d,e) indicating that the presence of the partner motor domain in the kinesin dimer reduces disorder and orients the motor domains toward the orientation seen in the other nucleotide states that induce strong microtubule binding. Monomeric and dimeric constructs in the strongly bound states display very similar LD 0 values (Table 1 and Fig. 2d,e). This result shows that the extra motor domain in the kinesin dimer does not introduce a new configuration. Thus, in the strongly 838 VOLUME 10 NUMBER 10 OCTOBER 2003 NATURE STRUCTURAL BIOLOGY

4 Figure 3 Models for kinesin dimer bound to microtubules. In the three models depicted, one motor domain (M1) is oriented relative to the microtubule axis as in Figure 1. Locations A and B are indicated in red and blue. The calculated LD 0 values for probes at locations A or B for each model are indicated. The disordered motor domain M2 is represented in model II with several superimposed orientations (±90 ) of the kinesin motor domain. bound states, both motor domains in the dimer are in a configuration similar to that of the single motor domain in the monomer. For comparison we calculated the combined LD 0 of two motor domain orientations according to three plausible alternative models (Fig. 3). In model I, the two motor domains are oriented relative to each other as in the crystal structure of the kinesin dimer 16 ; in model II, one motor domain is bound to the microtubule whereas the other is disordered (adopting any orientation ±90 relative to the microtubule); and in model III, both motor domains have similar orientations. Model I predicted LD 0 values of 0.19 and 0.12 for probes in locations A and B, respectively (0.14 and 0.12, respectively, if M2 is chosen as the microtubule bound motor domain). Model II predicted LD 0 values of 0.22 and 0.35 for probes in locations A and B, respectively. Model III has both motor domains in the same orientation, so it predicts the same LD 0 values as the monomers, 0.46 and 0.62 for locations A and B, respectively. The experimental values we observe for KD_A and KD_B in the strongly bound states (Table 1) are very close to model III and significantly (P < 0.01) different from models I or II. Therefore, the data indicate that the kinesin dimer, when it is bound to the microtubule, does not have its two domains oriented as in the kinesin dimer crystal structure (model I), nor does it have one domain bound and the other one disordered (model II). Models I and II predict lower LD 0 values (closer to 0) for the dimeric constructs as compared with the monomeric ones. In contrast, we observe very similar values in both cases or slightly higher LD 0 values for the dimeric constructs. Therefore, we conclude that in the strongly bound states induced by AMPPNP, ADP+AlF 4 or NN, both kinesin motor domains are bound to the microtubule in equivalent configurations. Motor orientation during processive movement To further investigate the configuration of the kinesin motor domains along their ATPase cycles, we measured the fluorescence anisotropy of KD_B during processive movement in the presence of ATP. These experiments were done at the single molecule level so that only actively moving molecules were monitored (Fig. 4a). For each moving molecule we determined two LD values corresponding to two sets of perpendicular axes (see Methods) and used these values to calculate the angle of the fluorescence dipoles projected on the microscope stage (x-y plane in Fig. 2b). A kymograph (image formed by stacking the image intensities along a microtubule over time) (Fig. 4b) shows the molecule moving at constant speed along the microtubule (see Supplementary Video 1 a b c d Figure 4 Fluorescence polarization of single kinesin molecules moving processively. (a) Composite image showing a single BSR-labeled kinesin molecule (green) on an Alexa-680-labeled axonemal microtubule (red). Scale bar, 1.5 µm. (b) Kymograph image of a KD_B molecule. The kymograph shows a molecule walking a 3-µm distance with a velocity of 560 nm s 1. The molecule is moving processively as it has gone through 375 ATP cycles (1 ATP per 8 nm displacement ). Vertical scale bar, 1 µm; horizontal scale bar, 2 s. (c) Intensity (arbitrary units) traces for each of the four excitation polarization directions used. (d) Histogram of the angles of the fluorescent dipole projected onto the x-y plane (microscope stage) relative to the microtubule angle. The angles were calculated using two LD values obtained from four fluorescence intensity traces like the ones shown in c. The average mobility factor r (see Methods) for the moving molecules is 0.48 (n = 54). Only molecules with r > 0.2 (n = 47) are included in the angles distribution. The average angle of the distribution is 20. NATURE STRUCTURAL BIOLOGY VOLUME 10 NUMBER 10 OCTOBER

5 Figure 5 Proposed model for kinesin processive translocation. The two kinesin motor domains (heads) are in dark and light yellow, the neck linker in red and the microtubule αβ tubulin dimer in light and dark blue (for further details see text). online). The fluorescence intensity of the molecule changes periodically over time following the alternation of the excitation light polarization direction (see Methods). The intensity modulation indicates that the dipole is well aligned during movement relative to its microtubule track. Traces of the fluorescence light intensity separated according to the four polarization directions of the excitation light (0, 45, 90, 135 ) used are shown in Figure 4c. The molecule shows strong anisotropy, as indicated by the different fluorescence intensities depending on the polarization of the excitation light. Using the four fluorescence signals we calculated the fluorophore dipole angle projected onto the microscope stage. For the particular example shown (Fig. 4a c), the fluorophore dipole has a projected angle relative to the microtubule axis of 0.2. This is the average angle of the probe on this motor during many ATPase cycles as the motor moved 3 µm (1 ATP hydrolysis per 8 nm step ). We determined the projected angles for many molecules to build a histogram with the frequency distribution of the estimated angles (Fig. 4d). A distribution of projected angles with values between 0 and β and maximum at β is expected for probes arranged with cylindrical symmetry, random azimuthal orientations around the symmetry axis and an axial angle equal to β (Fig. 2b). Random errors in the estimation of the projected angles will widen the distribution and shift the peak slightly toward smaller angular values. Using an error in the estimated projected angles of ±12 (see Methods), we estimated that distributions corresponding to β angles of 0 22 would have a peak between 0 and 10. Our measured distribution (Fig. 4d) has a clear peak at 0 10 indicating an axial angle β between 0 and 22. Thus, the average orientation of the kinesin motor domains during processive movement is similar (within 20 ) to the one estimated earlier using ATP analogs that induce strongly bound states (26 28, Table 1). DISCUSSION Configuration of the two kinesin motor domains Our results indicate that in the strongly bound states induced by AMPPNP, NN or ADP+AlF 4 both kinesin motor domains are bound to the microtubule in equivalent configurations. This configuration is also observed during processive movement. Previous mechanical studies have, in fact, indicated double head attachment to the microtubule in the presence of AMPPNP. However, structural evidence has been ambiguous. Electron microscopy data has been interpreted by different groups as indicative of double 7 or single head binding 5,6. Our results support a model in which both motor domains are bound to the microtubule in the presence of AMPPNP. In the same mechanical study mentioned 4, it was reported that in the absence of nucleotides (NN) the unbinding force of kinesin from the microtubule is half than when AMPPNP is present. The authors concluded that only one head is bound in the absence of nucleotides. Having only one head bound in the absence of nucleotides also seems to fit well with the observation that in the absence of ATP, microtubule interaction triggers the release of only one ADP per kinesin dimer 3,20,21. This observation suggests that in the absence of ATP (as in the NN state) one of the two motor domains is prevented from interacting with the microtubule 22,23. In contrast, our results indicate that in the absence of nucleotides the motor domains have a configuration as if both were bound to the microtubule. One possible explanation for the apparent discrepancy is that in our NN experiments, we observe an equilibrium structure in which the two heads are microtubule bound, whereas in the kinetics 3,20,21 and mechanics 4 experiments a relatively short-lived, single-head-bound intermediate is observed. A second possibility is that microtubule binding is not the only event required to accelerate ADP release. It is conceivable that after the two motor domains bind to the microtubule they would have different properties, such as binding strength or the ability to release ADP. Once the two identical motor domains bind to the microtubule, they are no longer equivalent as they must adjust to the translational symmetry of the microtubule. Kinesin structural elements located between motor and dimerization domains, such as the neck linker, would point in opposite directions (Fig. 5). We propose that the position of the neck linker controls ADP release and binding strength. The motor domain, whose neck linker is forced into the docked (forward-pointing) orientation, would trap ADP and would bind more weakly to the microtubule. Our results and previous ones 10 clearly indicate that the neck linker is sensitive to the nucleotide species present in the kinesin active site. Thus, it is conceivable that the position of the neck linker also regulates the affinity of the active sites for different nucleotide species. The proposed relationship between the neck-linker position and nucleotide affinity provides a mechanism to coordinate the ATPase cycle of the two motor domains. A model for processive translocation of kinesin Our results and the ideas previously discussed can be incorporated into the following model for processive kinesin translocation (Fig. 5). A kinesin dimer containing two ADP molecules 3 binds to the microtubule with both motor domains, creating an asymmetry in the necklinker configurations (steps 1 to 2). Only the motor domain with the neck linker pointing backward is able to release ADP. ATP binding to the forward head induces zippering of the neck linker (as previously proposed 10 ) causing detachment of the trailing head that moves 840 VOLUME 10 NUMBER 10 OCTOBER 2003 NATURE STRUCTURAL BIOLOGY

6 quickly to find another tubulin binding site farther ahead. The nowleading head releases ADP after microtubule binding (steps 2 to 3). This movement occurs in short time relative to the whole cycle so that on average the two heads are bound to the microtubule most of the time. ATP is hydrolyzed in the now-trailing head producing further zippering of the neck-linker domain (steps 3 to 4). P i is released from the trailing head and this motor domain becomes weakly attached to the microtubule in the ADP state (steps 4 to 5). State 5 is equivalent to 2 but with the two motor domains swapped and the centroid of the molecule displaced 8 nm to the right (microtubule + direction). In this model both motor domains are bound to the microtubule for most of the ATPase cycle, as our results indicate. This property helps the motor to keep its grip on the microtubule and explains its high degree of processivity. When both motor domains are bound, the neck linkers of each motor domain are pointing in opposite directions. This asymmetry, we propose, causes the two ATP cycles to be out of step as only the motor domain that has the neck linker pointing backward (unzippered) is able to release its ADP. METHODS Proteins and labeling. For this work we made five cysteine-light constructs (KM_A, KM_B, KM_C, KD_A and KD_B) based on the sequence of the ubiquitous human kinesin heavy chain gene (KIF5B). All naturally occurring cysteines were replaced by alanines and two cysteines were introduced at selected positions to allow attachment of a bifunctional thiol-reactive fluorescence probe. Constructs KM_A, KM_B and KM_C comprise the first 349 amino acids of KIF5B, which include only the motor domain. KD_A and KD_B comprise the first 559 amino acids including the motor and part of the dimerization domain. Constructs KM_A and KD_A have unique cysteines at positions 169 and 174, KM_B and KD_B at positions 64 and 71 and KM_C at positions 330 and 335. Proteins were purified by Ni-NTA agarose chromatography and labeled with the bifunctional fluorophore bis-((n-iodoacetyl)piperazinyl) sulfonerhodamine (BSR) (Molecular Probes) as described 8,9. We confirmed that the His-tagged and BSR-labeled constructs have sedimentation coefficients corresponding to monomers or dimers according to their length 24. Kinesin preparations were further purified by microtubule affinity (binding in the presence of AMPPNP and release in the presence of ATP) for the fluorescence microscopy experiments. The presence of the BSR probe crosslinking the two unique cysteines in the constructs was verified by trypsin digestion followed by liquid chromatography and electrospray mass spectrometry 8,9 (LC-MS). Double attachment was also indicated by control experiments in which single cysteine constructs were labeled with the bifunctional BSR probe. These single cysteine constructs have LD 0 values close to 0 regardless of the nucleotide conditions in contrast to the ones with the two cysteines at locations A, B or C. Axonemal microtubules were prepared from sea-urchin sperm as described 8,25. For the single molecule experiments, the axonemal microtubules were labeled with Alexa-Fluor 680 carboxylic acid, succinimydyl ester (Alexa-680, Molecular Probes) following previously described protocols 26. Fluorescence polarization microscopy. Axonemal microtubules were mixed with the BSR-labeled kinesin constructs, placed between two coverslips and observed (within a min period). The experimental solution was 12 mm PIPES (ph 6.8), 2 mm MgCl 2, 1 mm EGTA, 10 mm glucose, 0.1% (v/v) β-mercaptoethanol, 0.1 mg ml 1 catalase, 0.03 mg ml 1 glucose oxidase, 7.5 mg ml 1 BSA. Nucleotides were added depending on the experimental conditions to be tested: AMPPNP, 2 mm AMPPNP; ADP, 2mM ADP; ADP-AlF 4, 4 mm ADP, 2mM AlCl 3, 10 mm KF; no nucleotide, 5 units ml 1 apyrase with no added nucleotide; ATP, 1 mm ATP. Experiments were done at 21 C. The tubulin concentration for the experiments was µm. The concentration of kinesin motor heads was µm in the ensemble experiments and <8 nm in the single molecule ones. Thus, the ratio of tubulin to kinesin heads was at least 3:1. Microtubules decorated with the BSR-labeled kinesin constructs were imaged by wide-field epifluorescence. The experimental setup was essentially as described 8 with minor modifications. Laser light excitation (λ = 532 nm) was linearly polarized in four different transverse directions using an electro-optic modulator (EOM) (Conoptics) and was introduced into an inverted microscope (Nikon Eclipse TE300) through the epifluorescence port. The excitation light polarization axis in the microscope stage plane was changed to 0, 45, 90 and 135 every 100 ms. An image-intensified CCD camera (I-Pentamax, Roper Scientific) was synchronized to the EOM to collect the fluorescence images corresponding to each polarization excitation axis. Alexa-680-labeled axonemal microtubules were imaged using a CY5.5 filter cube set (Chroma) and excitation light from a mercury lamp. Data analysis. Digital movie files were collected from the CCD camera and analyzed using custom-written software. Fluorescence intensities for each polarization excitation direction were measured for microtubule segments or single molecule fluorescence spots 8,9. From the fluorescence intensities two LD values were calculated for each microtubule or molecule according to LD 0 90 = (I 90 I 0 ) / (I 0 + I 90 ) and LD = (I 135 I 45 ) / (I I 45 ), where I denotes fluorescence intensity and the subscript denotes the corresponding excitation light polarization direction. When no subscripts are used in the text, LD 0 90 is intended. The axial angle of the fluorophore relative to the microtubule long axis (β) (see Fig. 2b for angle definitions) in the ensemble experiments was calculated by a least-squares fit of the experimental data to the equation LD 0 90 (Γ,β,ω)= 3cos(2ω)/(1+8/[(3cos 2 β 1)(cosΓ +cos 2 Γ)]) (1) where ω is the microtubule long axis angle in the x-y plane, and Γ is the mobility cone angle 8. We used a cone angle of 32 in the calculations 9. The projected angle on the x-y plane of single moving molecules was calculated using the equation: cos 2 (α) = (1 LD / r) / 2 (2) where α is the projected angle on the x-y plane. This equation derives from the relationship between the intensity of light absorbed, I a,or emitted, I e, and the direction of polarization of the excitation light, I e cos 2 θ, where θ is the angle that the fluorophore absorption dipole makes with the excitation polarization axis 27. A correction factor, r, accounts for probe mobility 8,9 : r 2 = LD LD When calculating the angles we used only molecules with r > 0.2. Equation (2) gives two possible angles, α and 180 α. The ambiguity was solved using LD to calculate cos 2 (α 45) = (1 LD / r) / 2 (3) Only one angle satisfies equations (2) and (3) and this is the angle of the dipole projected on the x-y plane. The standard error in the calculated projected angles using equation (2) was estimated as ±12 by propagating the standard error of the four independently measured intensities (I 0, I 45, I 90, I 135 ) used to calculate LD/r. The angle of the microtubule long axis in the x-y plane (ω) was subtracted from the dipole angle to obtain the relative angle of the dipole with respect to the microtubule. Note: Supplementary information is available on the Nature Structural Biology website. ACKNOWLEDGMENTS We thank G. Rogers, D. Sharp, D. Buster and M. Akabas for discussions and critical reading of the manuscript, H. Deng for mass spectrometry analysis and E. Peterman for experimental advice and discussions. This project was supported by a US National Institutes of Health grant to H.S. COMPETING INTERESTS STATEMENT The authors declare that they have no competing financial interests. Received 24 January; accepted 24 July 2003 Published online at 1. Goldstein, L.S.B. & Philp, A.V. The road less traveled: emerging principles of kinesin motor utilization. Annu. Rev. Cell Dev. Biol. 15, (1999). 2. Howard, J., Hudspeth, A.J. & Vale, R.D. Movement of microtubules by single kinesin molecules. Nature 342, (1989). NATURE STRUCTURAL BIOLOGY VOLUME 10 NUMBER 10 OCTOBER

7 3. Hackney, D.D. Evidence for alternating head catalysis by kinesin during microtubulestimulated ATP hydrolysis. Proc. Natl. Acad. Sci. USA 91, (1994). 4. Kawaguchi, K. & Ishiwata, S. Nucleotide-dependent single- to double-headed binding of kinesin. Science 291, (2001). 5. Hirose, K., Lockhart, A., Cross, R.A. & Amos, L.A. Three-dimensional cryoelectron microscopy of dimeric kinesin and ncd motor domains on microtubules. Proc. Natl. Acad. Sci. USA 93, (1996). 6. Arnal, I., Metoz, F., DeBonis, S. & Wade, R.H. Three-dimensional structure of functional motor proteins on microtubules. Curr. Biol. 6, (1996). 7. Hoenger, A. et al. Image reconstructions of microtubules decorated with monomeric and dimeric kinesins: comparison with X-ray structure and implications for motility. J. Cell Biol. 141, (1998). 8. Peterman, E.J., Sosa, H., Goldstein, L.S. & Moerner, W.E. Polarized fluorescence microscopy of individual and many kinesin motors bound to axonemal microtubules. Biophys. J. 81, (2001). 9. Sosa, H., Peterman, E.J., Moerner, W.E. & Goldstein, L.S. ADP-induced rocking of the kinesin motor domain revealed by single-molecule fluorescence polarization microscopy. Nat. Struct. Biol. 8, (2001). 10. Rice, S. et al. A structural change in the kinesin motor protein that drives motility. Nature 402, (1999). 11. Wittinghofer, A. Signaling mechanistics: aluminum fluoride for molecule of the year. Curr. Biol. 7, R682 R685 (1997). 12. Crevel, I.M., Lockhart, A. & Cross, R.A. Weak and strong states of kinesin and ncd. J. Mol. Biol. 257, (1996). 13. Sosa, H. et al. A model for the microtubule-ncd motor protein complex obtained by cryo-electron microscopy and image analysis. Cell 90, (1997). 14. Kikkawa, M. et al. Switch-based mechanism of kinesin motors. Nature 411, (2001). 15. Sack, S. et al. X-ray structure of motor and neck domains from rat brain kinesin. Biochemistry 36, (1997). 16. Kozielski, F. et al. The crystal structure of dimeric kinesin and implications for microtubule-dependent motility. Cell 91, (1997). 17. Hua, W., Young, E.C., Fleming, M.L. & Gelles, J. Coupling of kinesin steps to ATP hydrolysis. Nature 388, (1997). 18. Coy, D.L., Wagenbach, M. & Howard, J. Kinesin takes one 8-nm step for each ATP that it hydrolyzes. J. Biol. Chem. 274, (1999). 19. Schnitzer, M.J. & Block, S.M. Kinesin hydrolyses one ATP per 8-nm step. Nature 388, (1997). 20. Ma, Y.Z. & Taylor, E.W. Interacting head mechanism of microtubule-kinesin ATPase. J. Biol. Chem. 272, (1997). 21. Gilbert, S.P., Moyer, M.L. & Johnson, K.A. Alternating site mechanism of the kinesin ATPase. Biochemistry 37, (1998). 22. Schief, W.R. & Howard, J. Conformational changes during kinesin motility. Curr. Opin. Cell Biol. 13, (2001). 23. Vale, R.D. & Milligan, R.A. The way things move: looking under the hood of molecular motor proteins. Science 288, (2000). 24. Jiang, W., Stock, M.F., Li, X. & Hackney, D.D. Influence of the kinesin neck domain on dimerization and ATPase kinetics. Biochemistry 37, (1997). 25. Gibbons, I.R. & Fronk, E. A latent adenosine triphosphatase form of dynein 1 from sea urchin sperm flagella. J. Biol. Chem. 254, (1979). 26. Pierce, D.W. & Vale, R.D. Assaying processive movement of kinesin by fluorescence microscopy. Methods Enzymol. 298, (1998). 27. Lakowicz, J.R. Principles of Fluorescence Spectroscopy (Kluwer Academic, Plenum Publishers, New York, 1999). 28. Corrie, J.E.T. et al. Dynamic measurement of myosin light-chain-domain tilt and twist in muscle contraction. Nature 400, (1999). 842 VOLUME 10 NUMBER 10 OCTOBER 2003 NATURE STRUCTURAL BIOLOGY