Complex Formation with Kinesin Motor Domains Affects the Structure of Microtubules

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1 doi: /j.jmb J. Mol. Biol. (2004) 335, Complex Formation with Kinesin Motor Domains Affects the Structure of Microtubules A. Krebs*, K. N. Goldie and A. Hoenger European Molecular Biology Laboratory, Meyerhofstrasse Heidelberg, Germany *Corresponding author Microtubules are highly dynamic components of the cytoskeleton. They are important for cell movement and they are involved in a variety of transport processes together with motor proteins, such as kinesin. The exact mechanism of these transport processes is not known and so far the focus has been on structural changes within the motor domains, but not within the underlying microtubule structure. Here we investigated the interaction between kinesin and tubulin and our experimental data show that microtubules themselves are changing structure during that process. We studied unstained, vitrified samples of microtubules composed of 15 protofilaments using cryo electron microscopy and helical image analysis. 3D maps of plain microtubules and microtubules decorated with kinesin have been reconstructed to, 17 Å resolution. The ab-tubulin dimer could be identified and, according to our data, a- and b-tubulin adopt different conformations in plain microtubules. Significant differences were detected between maps of plain microtubules and microtubule kinesin complexes. Most pronounced is the continuous axial inter-dimer contact in the microtubule kinesin complex, suggesting stabilized protofilaments along the microtubule axis. It seems, that mainly structural changes within a-tubulin are responsible for this observation. Lateral effects are less pronounced. Following our data, we believe, that microtubules play an active role in intracellular transport processes through modulations of their core structure. q 2003 Elsevier Ltd. All rights reserved. Keywords: microtubules; tubulin; kinesin; structure; electron microscopy Introduction Microtubules (MTs) are ubiquitous in the cytoplasm of eucaryotic cells and play important roles in the maintainance of the cells. They interact with a variety of proteins in various transport processes and participate in the formation of complex structures such as the axoneme, the centrosome and the mitotic spindle. MTs are also involved in cell motility and cell morphogenesis. 1 3 The basic unit of the microtubule is the ab-tubulin hetero-dimer. Tubulin-heterodimers form protofilaments, which aggregate laterally into the tubular microtubule-structure. The protofilaments are parallel and axially staggered by about 0.92 nm in a left-handed direction, forming a Abbreviations used: MT, microtubules; MAP, microtubule-associated proteins. address of the corresponding author: krebs@embl-heidelberg.de B-lattice. 4 Thereby the main lateral contacts are usually from a to a and b to b subunits. In microtubules composed of 13 protofilaments (the most common type in vivo) this regular B lattice is interrupted by so-called seams forming A-lattice contacts (i.e. a b subunit contacts between two neighboring protofilaments). 5 7 The protofilament number of microtubules may vary from 9 to 16 8 resulting in microtubules with different configurations and symmetry. 9 Some of them, such as 15-protofilament microtubules can be truly helical. There, the lattice requires a slight correction from the tubular axis, which forces the protofilament into an axially supertwisted conformation to compensate for the additional two protofilaments. In the case of the helical 15-protofilament microtubules this supertwist is right-handed. a- and b-tubulin arrange head to tail and thus form polar microtubules with a highly dynamic plus end and the less active minus end. 10 a- and b-tubulin are very similar, both, in sequence and /$ - see front matter q 2003 Elsevier Ltd. All rights reserved.

2 140 Structural Changes in Microtubules structure. 11,12 Both subunits bind GTP, but only the GTP in b-tubulin will be hydrolyzed upon polymerization. This results in a tubulin dimer within the microtubule, which contains GTP in the a-subunit and GDP in the b-subunit. Structural detail of the ab-tubulin hetero-dimer is known from electron crystallography studies on zincinduced tubulin sheets 11,12 and from single particle analysis of microtubules with 13 protofilaments. 13 These studies lead to the conclusion, that a- and b-tubulin adopt similar structures, both in zincinduced sheets as well as in 13-protofilament microtubules. From the combination of the tubulin crystal structure with a low-resolution envelope of plain helical microtubules we know, that the long C-terminal helices of the tubulin subunits form the crest on the outside of the protofilament making the side, where the motor proteins walk rather smooth. The inside of the microtubule is built up mainly of two big loops forming distinct bumps. 14 Motor proteins of the kinesin family move along this crest and thereby achieve a large variety of transport processes. The structural basis of these transport processes and the exact stepping mechanism are the subject of numerous investigations, but the details are still not completely understood. Thus several models for kinesinmediated transport have been discussed for conventional kinesin. However, so far only the role of the motor proteins and changes in their structure have been investigated and described. None of these studies assume a direct connection to dynamic structural changes within the microtubule upon motor interaction. Thus, a prominent common feature in all these studies is the fact that microtubules are mostly seen as sturdy tracks, with the structures of a- and b-tubulin being similar. An active involvement of microtubules in motor function has never been discussed. But microtubules are very active structures. It is known, that microtubules are highly dynamic, changing rapidly between phases of growth and shrinkage. 10,23 The dynamic behavior seems to be regulated by microtubule-associated proteins (MAPs), 24,25 which act highly specific according to cellular needs. Microtubule dynamics is thought to be related to the nucleotides present in the tubulin subunits 23,26 (GTP or GDP) and therefore structural changes in the microtubule according to cellular needs are likely. Tubulin alone adopts a curved conformation as it depolymerizes, 27,28 which reflects the favored conformation for GDP-tubulin released from the constraints of the microtubule lattice. Curved tubulin dimers are also observed in magnesium-induced GDP-tubulin ring structures. 29 Even the microtubule ends themselves have different properties, and their structures change significantly between growing and shrinking phases. 30 All these studies suggest structurally active microtubules with structural changes serving as basis for the dynamic behavior of microtubules. There is no obvious reason why microtubules should be able to change their structure as basis of dynamic behavior, whereas in connection with motor proteins no changes are present. Therefore we wondered, if the dynamic behavior is related to motor binding? And if so, are structural changes present when motor proteins are bound to the microtubules? To address this question we studied unstained, vitrified samples of plain microtubules and microtubules complexed with the motor protein kinesin from Neurospora crassa using cryo-electron microscopy and helical 3D image reconstruction. We focused exclusively on microtubules composed of 15 protofilaments and reconstructed 3D maps to,17 Å. We find different conformations for a- and b-tubulin in plain microtubules, which enables us to assign the subunits not only in microtubule kinesin complexes but also in plain microtubules. Upon motor binding we detect major conformational changes within the tubulin portion of the microtubule, suggesting that microtubules indeed do react actively to motor binding. With our current data we can demonstrate that kinesin has a stabilizing effect on microtubules and protofilaments and we think, that this effect may be of great importance within the cellular environment. Results and Discussion Image processing All 3D maps presented in this study were calculated from helical 15-protofilament microtubules. The symmetry of these tubes can be described by a helical selection rule of 257/137 (asymmetric units/helical turns). 31 This selection rule is the product of a convolution of a short-pitched, lefthanded helical path with a helical pitch of 16 nm, (following tubulin dimers laterally) and the righthanded supertwist of protofilaments. Our reconstructions are averages of individual datasets, of which each is composed of about 2 3 helical repeats. One data set includes about asymmetric units (tubulin dimers). This number was chosen as a compromise between keeping the length of tubular fragments as short as possible (to eliminate areas with distortions) and generating useful phase and amplitude data for alignment and averaging. One of the key issues in this work was identifying the ab-tubulin boundary. This we attempted, because we found alternating differences along the protofilaments already in filtered maps from single data sets (Figure 1C). Figure 1 illustrates which area in a Fourier transform (D, F, H) is related to certain features in back transformed maps (C, E, G). The micrograph in A shows approximately one helical repeat and its corresponding diffraction pattern is shown in B. Diffraction of such an image typically reveals three visible layer

3 Structural Changes in Microtubules 141 Figure 1. The crucial image information related to the structural definition of the ab-tubulin dimer outline is located in a cluster of layer lines around 1/8 nm (layer lines around n; l ¼ 22,17) and beyond. n ¼ Bessel order, l ¼ layer line number. Diffraction data (B) obtained from an image of the length of one helical repeat (A) show, that this information is rather weak. Images C, E and G are backtransformations of Fourierfiltered diffraction data including only the layer line data marked with yellow lines in D, F, and H. The back-transformed images, and in particular the insets in C and G show clearly that the fenestrations appear different along the protofilament axis. C, Less stringent filtering; G, more stringent filtering reduced to the 1/8 nm cluster (including n; l ¼ 22,17), two strong layer lines at the 1/4 nm cluster (n; l ¼ 24, 34 and n; l ¼ 11,35) and the first strong layer line (n; l ¼ 15,1). If the information around the 1/8 nm cluster is absent (E: including information only from three layer lines; n; l ¼ 15,1; n; l ¼ 24,34; n; l ¼ 11,35), the structure shows holes of similar size repeating every 4 nm along the protofilament axis. Thus the information, which is contained in the 1/8 nm cluster, is crucial for the correct reproduction of the structure. I, Comparison of the collapsed power spectrum of an average of 50 data sets (black) with one from a selected piece of a native microtubule (gray). The diffraction pattern of a single piece of the microtubule shows mainly three strong layer lines (n; l ¼ 15,1; n; l ¼ 24,34; n; l ¼ 11,35, compare with B, F). Averaging of,50 individual data sets by our method described here, however, shows that information on weak layer lines (e.g. the layer lines in the 1/8 nm cluster, in particular n; l ¼ 22,17) can be extracted as well. The relative intensities given for the data of the single piece are multiplied by ten with respect to the averaged data. lines: The first layer line with the Bessel order 15 ðn; l ¼ 15; 1Þ gives information about the protofilament supertwist. Layer line 34 (Bessel order n ¼ 24; layer line number l ¼ 34) gives information on the axial 4 nm repeat and the next layer line (Bessel order n ¼ 11; layer line number l ¼ 35) represents a convolution of the axial 4 nm repeat as in layer line 34 with the protofilament supertwist. The

4 142 Structural Changes in Microtubules intensities of all other layer lines are only marginally above the noise level and therefore virtually invisible. The back transformation of these three strong layer lines alone produces supertwist information and information on the 4 nm repeat (F and E) and clearly the holes in E are of similar size 4 nm apart (one tubulin subunit has a dimension of roughly 4 nm in the microtubule lattice). Thus, distinct features originating from different a- and b-tubulin conformations cannot be determined by using information from the three strong layer lines only. This information can only be retrieved from the cluster of layer lines positioned around the 1/8 nm repeat (e.g. layer line n; l ¼ 22,17; see C/D and G/H). The map in G was produced from three strong layer lines plus layer lines around n; l ¼ 22,17. We clearly see differences, which alternate along the protofilament axis. The holes are still 4 nm apart, but now they are of alternating different size. Therefore, in this case, holes with similar size repeat every 8 nm (note that one tubulin-dimer has a length of about 8 nm). Thus, it is information in particular which has to be retrieved accurately, to be able to outline the ab-tubulin boundary. In addition, as we detect alternating differences already in filtered maps from single images (C) we are sure, that a- and b-tubulin adopt different conformations in plain microtubules. We want to point out, that meaningful information on the 1/8 nm cluster is weak. Therefore we demonstrate in Figure 1I that this information is real and can be amplified in the averaging process, whereas background noise diminishes (e.g. see layer line n; l ¼ 22,17). We used these data to identify differences between the tubulin subunits in the plain microtubule. Averaging For an averaged map about 80 data sets (near and far sides) were selected. All our images were corrected for the effects of the contrast transfer function (CTF) prior to averaging. The individual data sets were shifted against a preselected reference and their phase residuals were calculated. In our calculations we followed the standard helical processing procedures but adapted some of the procedures to make sure that the dimer outline information will not be lost. We integrated a reference based refinement procedure, in which the reference was iteratively improved. The reference used for alignment of the individual data sets is built purely from raw data, eliminating the need for modeled data and any model-bias. Alignment of data sets to this, initially crude reference lead to an average structure, which itself could then be used as an improved reference. Successional rounds of refinements produced the final data set. The strategy for averaging was the following: an initial average was calculated from six excellent images against a reference taken from one single image among that group. In this average only three strong layer lines, namely layer line 1 (n; l ¼ 15,1; supertwist information), and layer lines 34 and 35 (n; l ¼ 2 4,34; n; l ¼ 11,35; 4 nm cluster ¼ axial monomer repeat) were included (compare Figure 1F). This procedure was chosen because it resulted initially in the best possible agreement of these particularly strong layer lines (phase residual ¼ 118) without obstruction from noisy data around the 8 nm cluster. Of course, now this first average reveals a merge between a- and b-tubulin, randomly shifted against each other and either aligned correctly or shifted by 4 nm along the protofilament axis. To regain the data necessary for a correct outline of the tubulin dimer we went back and inspected all individual layer lines (amplitudes and phases) of all individual images and compared them to each other with respect to their (coincidental) alignment. Consequently some of the individual datasets were shifted correctly and some were not. Inspecting the phase values on the predicted positions of layer lines around the 8 nm cluster, and layer line 17 (n; l ¼ 22,17) in particular, allowed eliminating the wrongly shifted ones (mis-alignment with phases of the reference) from the correctly shifted data (correct alignment with phases of reference). This data was then taken to generate a new refined reference, which then included layer line data from both, the 8 nm and the 4 nm clusters. This procedure was cycled several times with different groups of images in an iterative way and resulted finally in the data shown in Figure 2. The mean phase residual of all data sets included in the final merge was 358 and the applied phase residual limit of 408 excluded 20 data sets from averaging in the final rounds. It became evident, that the first rounds of refinement, establishing accurate data for layer lines around n; l ¼ 2 2,17 proved critical for further averaging. Final layer line data from selected layer lines are shown in Figure 2A (plain microtubule) and B (microtubule kinesin complex, using the monomeric construct of the kinesin from the fungus N. crassa) together with individual amplitude and phase data for all data sets included in the final merges after correction of the effects of the contrast transfer function. In general, phase data from strong layer lines (e.g. n; l ¼ 15,1; n; l ¼ 24,34; n; l ¼ 11,35) were in perfect agreement over areas with high amplitudes. The signal, mainly responsible for the alternating details ðn; l ¼ 22; 17Þ is weak but consistently present (Figure 2A). The clustering of phase data from individual data sets (red box), apparent at regions with visible amplitudes, shows that correct alignment was achieved. Averaging the motor kinesin complex was much easier, because in that case layer line n; l ¼ 22; 17 is much stronger due to the motor protein, which is bound to every b-tubulin subunit (Figure 2B). The resolution of our maps, determined by calculating the Fourier shell correlation of two independent sub averages, was determined to be,17 Å (Figure 3, Table 1).

5 Structural Changes in Microtubules 143 Figure 2. Selected layer-line amplitudes and phases corrected for the effects of the contrast transfer function of the final average of plain microtubules (A) compared with the final average of the microtubule kinesin complex (B). The most obvious differences are in the cluster of layer lines at 1/8 nm (e.g. n; l ¼ 22,17), which are highly amplified by the presence of one motor unit per tubulin dimer. In the boxes individual amplitude and phase data corrected for the effects of the contrast transfer function are shown for selected layer lines. We want to point special attention to the clustering of the phases on layer line n; l ¼ 22,17 (red boxes). In both cases individual phase data agree well with the averaged phase, although the amplitudes are much stronger in the microtubule motor complex due to the motor binding. Comparison of the phases of different layer lines from data of the plain microtubule (top) with the corresponding data of the microtubule kinesin complex (bottom) shows that they agree well. Important phase data are outlined in blue ðn; l ¼ 15; 1Þ; red ðn; l ¼ 22; 17Þ and yellow ðn; l ¼ 24; 34Þ and show the perfect alignment of phases. Assignment of alpha and beta tubulin subunit in plain microtubules First the polarity of the maps was checked. The polarity can be determined according to the slew of the tubulin subunits, which is slightly counterclockwise when viewing the projected map from its plus end towards its minus end. 6,14,32 Then we aligned the map of the plain microtubule to one of the microtubule motor-complex (see

6 144 Structural Changes in Microtubules Figure 3. Estimation of the resolution of the map of the plain microtubule (blue) and the microtubule kinesin complex (yellow). Fourier shell correlation (FSC) and differential phase residual (DPH) was calculated between two independent halves of both data sets. The black line represents the expected value of the Fourier shell correlation with background noise, corresponding to the usual 3s curve. The FSC correlation for the plain microtubule data set drops to 0.5 at a value of 0.056, corresponding to an estimated resolution of 17.8 Å, and the FSC of the microtubule kinesin complex drops to 0.5 at 0.057, indicating a resolution of about 17.5 Å. Details of data analysis are given in Table 1. alignment of phases in the colored boxes in Figure 2). It was established from previous studies that kinesin motors interact predominantly with b-tubulin, and that the motor locates at the outer rim of the plus end. 33,34 Hence, the a-tubulin subunit is exposed at the minus end. 35 Other independent studies state, that the main mass of the kinesin head is associated with the tubulin subunit closer to the plus end of the microtubule. 32 This agrees well with data revealing that the exchangeable GTP-site in b-tubulin is exposed at the plus end of the microtubule. 36 Hence it seems reasonable to correlate the motor binding site to the b-subunit at the plus end of the microtubule map. Accordingly, correct alignment of the map of the microtubule kinesin complex to the map of the plain microtubule was the way to go for a correct assignment of a- and b-tubulin in the map of the plain microtubule. The mathematical alignment of the two maps, however, is not straightforward. The features outlining the ab-dimer are weak in both maps, and in the microtubule kinesin complex the b-subunit is labeled by the motor density, whereas this label is missing in the plain microtubule. When the map of the plain microtubule is used as reference to align the map of the microtubule motor complex two possible origins for the microtubule motor complex are given by the calculations. The two possible origins were 4 nm apart and correlated with opposite polarity of the maps. Cross-correlation values of these two possibilities were very close (compare cross-correlation values of and 0.763) and therefore a mathematical determination was not possible in that case. However, as the polarity in both maps is unambiguous according to other studies (outlined above, see Refs. 6,14,32), the result with the wrong polarity could be ruled out. When we used the map of the microtubule motor complex as initial reference to align the data sets of the map of the plain microtubule we got better results. In that case, the accurate phases from layer line n; l ¼ 22; 17 helped determine the origin of the map of the plain microtubule. The procedure was carried out with different sub averages and checked crosswise. The resulting alignments were consistent in all attempts. To confirm our alignment, phase data in Figure 2 should be compared. Especially the phases of plain microtubules at n; l ¼ 22; 17 (red box, top) should be compared to the corresponding phase data of the microtubule kinesin complex (red box, bottom) as well as the data from n; l ¼ 15,1 and n; l ¼ 24,34 (blue and yellow boxes, respectively). If one of the maps would be shifted by 4 nm along the protofilament axis (according to one tubulin subunit), the phase data would be shifted by Such a shift would destroy the perfect alignment of the two data sets. Hence, this confirms that our alignment is correct. Electron densities in plain microtubules and microtubule kinesin complexes Surface rendered representations of the plain microtubule (Figures 4B and C) and of the microtubule kinesin complex (Figure 4F and G) point out the great similarity of both structures and the most prominent features of microtubule maps, the fenestrations, 4 are clearly visible. Table 1. Data analysis Parameters Plain microtubules Microtubule nk355 complexes Number of microtubules Number of data sets a Number of asymmetrical units b 35,000 42,000 Defocus range (nm) Resolution cutoff c (Å) a Number of independent near and far sides included in the final average. b Number of tubulin dimers included in the final average. c The resolution of the data sets was determined by Fourier shell correllation (see Figure 3).

7 Structural Changes in Microtubules 145 Figure 4. Electron micrographs (A and E) and 3D reconstructed surface rendered maps of the plain microtubule (upper half) and the microtubule kinesin complex (lower half). Electron density maps of the MTs are shown from the exterior (the side where the motor proteins walk, B and F), as view from the central channel (C and G) and as side view along one protofilament axis (D and H). Tubulin heterodimers are arranged head to tail along the protofilaments leading to low-density areas between neighboring protofilaments, clearly visible as holes (pointed out with dark blue arrows). These holes are of alternating different size in plain microtubules (B and C), implying that a- and b-tubulin contribute unequally to the overall structure. Contacts between two subunits of one heterodimer appear broader than inter-dimer contacts. This results in smaller fenestrations between two neighboring dimers and larger ones (lower density) where four dimers meet. D, A clear distinction between neighboring tubulin heterodimers in the high-density core (pink) is observed, as the connection is broken along the protofilament. In addition, the core of the a-tubulin subunit appears weaker (white arrow) in comparison to the core of b-tubulin. These are indications, that the tubulin subunits adopt different conformations in plain microtubules. In the map of the decorated microtubule (F and G) the holes are more similar. H, In higher contour levels (pink) the central mass of the protofilaments appears regular. M, points out the density of the bound kinesin motor.

8 146 Structural Changes in Microtubules The fenestrations or holes (blue arrows) are the low-density areas between neighboring protofilaments, which are a result of the architecture of the microtubules of roughly globular tubulin subunits. However, these low-density areas are at the same time a very visible indication of conformational differences between plain microtubules and microtubules decorated with kinesin. Clearly the size of the holes varies in the map of the plain microtubule (blue), while they appear almost identical in motor-decorated maps (yellow). These differences are one of the most obvious indications of different tubulin conformations in our maps. Other alternating differences along the microtubule axis can be seen on closer inspection of Figure 4B (white arrows). Unlike previous reconstructions this 3D map of plain microtubules reveals structural details exhibiting a clear 8 nm repeat with details never described before at that level of resolution. The most obvious features along the protofilament axis are the low-density areas between two adjacent protofilaments (blue arrows). The holes themselves are 4 nm apart but as they alternate in size, holes with similar size are repeating every 8 nm. Interestingly these low-density areas appear of equal size every 4 nm in motor-decorated maps (Figure 4F and G; discussed below). Thus, in the plain microtubule we observe a varying density distribution along the protofilaments, outlining a tubulin dimer (the a and b tubulin subunits belonging to one dimer are indicated). The varying density distribution is not only reflected in unequal low-density areas but also in an uneven highdensity core of the map, which becomes obvious using higher contour levels (Figure 4D). The pink high-density core of the plain microtubule is broken along the protofilament, revealing more density between the tubulin subunits belonging to one dimer than between tubulin subunits of adjacent dimers. Variations in contacts between tubulin subunits along the protofilament axis in plain microtubules have been suggested, 14 but only here we provide experimental evidence for their existence. In addition we find, that the high-density core of plain microtubules varies from a- to b-tubulin, because the core of the a-tubulin subunit appears smaller than the core of b-tubulin in Figure 4D (white arrows). This shows that the tubulin subunits adopt different structures in plain microtubules and it may indicate increased structural flexibility of a-tubulin. So far, differences between subunits have only been observed in data obtained from negatively stained samples but have not been seen in other cryo-em maps of plain microtubules. 13,14 The map of the microtubule motor complex, on the other hand, seems to be much more even regarding fenestrations (Figure 4F and G) and high-density core (pink in H). The densities for a- and b-tubulin are roughly equal and the highdensity core is not broken along the protofilament axis. All these qualitative observations could result from a strengthening of inter-dimer contacts along the protofilaments upon motor binding. This result matches well with that of our observations of microtubules and protofilaments in solution, shown in Figure 5A and B: only in the presence of motors we do find elongated protofilaments and small protofilament bundles. In preparations of plain microtubules stabilized single protofilaments have not been observed and protofilaments disassemble rapidly into small oligomers or curved structures (negative stain image in Figure 5C, tubulin oligomers in the background). These results show that a taxolstabilized microtubule does not splay and fall apart as protofilaments but as tubulin oligomers. The behavior of plain microtubules has been investigated in detail and it was shown that in general microtubules disassemble primarily into oligomers or highly curved structures. 27,40 What we observe with kinesin-decorated microtubules is different. In that case the protofilaments are stabilized over a long distance with low curvature and such a stabilizing effect can only be observed with motor decorated microtubules. Single stabilized protofilaments are always observed, even if excess tubulin is separated from the sample prior to decoration. Without motors, we observe tubulin oligomers and microtubules only but never do we find stabilized protofilaments. This raises the question of the origin of the protofilaments: weather they are newly polymerized protofilaments or if they are the result of separated microtubules because of a lateral destabilizing effect of the motors. According to our data, statistically significant conformational changes within the tubulin portion lead to a more stable protofilament along its axis. This conclusion arises from t-test analysis of the maps, which is shown in detail in Figure 6. However, from a qualitative point of view, a lateral destabilization of neighboring protofilaments might also occur. This is supported by the observation of larger and more regular low-density areas in the motor-decorated map (Figure 4F and G) in comparison with the low-density areas of the map of the plain microtubule (Figure 4B and C). In the statistical analysis however, these differences did not show up with a high enough significance level. This might be due to the fact, that the resolution we achieve in this study is not high enough and that the observed changes are small and therefore hard to detect. Consequently we interpret stabilization along the protofilament axis as the major effect whereas lateral destabilizing effects seem to be much smaller order and we can only speculate that such effects occur. Conformational changes upon complex formation with kinesin motor domains Quantitative analysis of the maps reveals

9 Structural Changes in Microtubules 147 Figure 5. Cryo-electron microscopy images directly illustrate the increase of axial protofilament stability in the presence of kinesin motors (A and B). C, Plain microtubules, shown here in negative stain disassemble in small oligomers but do not form the elongated protofilament bundles as observed for microtubule motor complexes shown in A and B. All preparations were washed twice after polymerization. significant changes upon motor binding within the tubulin portion; these areas of major changes are displayed as magenta and red volumes in Figure 6. It is important to state, that small changes, such as amino acid movements or movements of small parts of the proteins are unlikely to be detected with our resolution of 17 Å. Therefore, the observations we are describing are of much larger order. In Figure 6 we are referring to positive densities at locations of increased mass upon motor binding (red in A, E G), and vice versa for negative densities (magenta in A, E G). Consequently, the density corresponding to the kinesin motor domain itself shows up positive (red in E) and significant structural changes within the tubulin portion may be either positive or negative. Most pronounced is an increase in mass near the center of a-tubulin towards the binding interface to the next tubulin-heterodimer (A, F and G). At the outer rim of both subunits a mass decrease is observed (E G) with the changes in b-tubulin being weaker. Thus, surprisingly, the most striking differences are detected within a-tubulin rather than b-tubulin, although b-tubulin is the primary motor binding site of Neurospora kinesin. 33 The conformational changes within a-tubulin are indicative for a rearrangement and stabilization, leading to an increased density near the center of Figure 6. Effects of kinesin binding illustrated on a single protofilament in side view (A) and in contour-sections cut perpendicular to the microtubule axis (B G). Electron densities are displayed in blue (plain microtubule) and yellow (microtubule kinesin complex) with red and magenta volumes (major changes upon motor binding). Most obvious is the increase of density within a-tubulin (red volumes in A, F and G). The location of this difference corresponds nicely to the observed density variations of the high-density core (Figure 1D), and is, we believe, the origin for the even high density core observed in the microtubule kinesin complex (Figure 1H). A slight decrease of density (magenta) is detected at the outer rim of both subunits (A, F and G). E, b-tubulin does not show internal changes. The density on the outside (red) refers to the density of the motor protein (M).

10 148 Structural Changes in Microtubules the subunit. This comes together with a slight rotation towards the left of the outer rim of the microtubule when viewed from the plus end (F and G) leading to the continuous high-density core, which we observe in microtubule kinesin complexes (Figure 4H). The main response of the b-tubulin to motor binding is most likely located on the interface of tubulin and motor. However, this is difficult to interpret as it coincides with the position of the difference peak related directly to the motor density (Figure 6E). We used taxolstabilised microtubules throughout this study and it is clear that taxol itself stabilizes microtubules, but much less vigorously than a complete decoration with motors does. This becomes obvious by looking at the raw-data images in Figure 5. We therefore believe, that the conclusions drawn here are not affected by taxol stabilization. A refined model of a dynamic microtubule For a more detailed analysis at near-atomic resolution we modeled the tubulin dimer crystal structure 12 (1JFF.pdb) as a rigid body into our maps in accordance with the orientation given by Nogales et al. 14 Since the differences between a- and b-tubulin observed here are not directly interpretable with the crystal structure, we refined the overall positions of the a- and b-tubulin subunits independently using the program package URO. 41 In Figure 7, in our tubulin-model of the plain microtubule we point out the locations of major changes upon motor binding. According to our model, the major density increase upon motor decoration in the a-tubulin is located near the end of helix 7 (helix 7 is colored black) and less pronounced at the beginning of helix 10 (red volumes). The origin of this difference density may either be an increased rotational flexibility of the a-subunit in plain microtubules, increased structural flexibility within the molecule, or a combination of both. Towards the outer rim of the microtubule the negative density is positioned near the end of helix 4 and the beginning of the loop B3-H3 (magenta). This peak can easily be caused by a slight clockwise axial rotation of the a-subunit upon motor binding (when viewed from the minus-end). A similar, but smaller peak is also observed in the b-subunit at that position. In principle it might be possible that the C terminus of the b-subunit is responsible for the signal but at the resolution achieved, we cannot state that with sufficient certainty. Microtubule dynamics is related to the structure of the subunits A huge number of biochemical and functional studies is available, pointing out the possibility for different tubulin conformations within Figure 7. Stereo view of the proposed model of the basic unit in plain microtubules seen from the exterior (outside of the microtubule). Areas affected by motor binding are red (density increase upon motor-binding) and magenta (density decrease) and the map of the plain microtubule is shown in gray. Green: a-tubulin with GTP bound, helix 7 is drawn in black, blue: b-tubulin with GDP bound, yellow: Taxol. In our modeling rotational and translational refinement of a- and b-tubulin were allowed independently; initially the crystal structure by Löwe et al. 12 was used.

11 Structural Changes in Microtubules 149 microtubules. The effect of nucleotide hydrolysis and the timing of its occurrence in b-tubulin has been subject of several investigations. 23,26,42,43 After an initial phase, a- and b-tubulin contain different nucleotides, GTP in a-tubulin and GDP in b-tubulin. GTP-b-tubulin remains at the ends of microtubules forming a GTP-cap. 44,45 This cap is believed to prevent depolymerization (forcing the GDP-tubulins within the microtubule in a straight conformation), and upon GTP-hydrolyzation protofilaments disassemble laterally and curl outwards 27,28,40 adopting their favored curved (GDP-tubulin) conformation. Thus, a conformational change within the tubulin subunits is assumed to explain microtubule dynamics. Interestingly, motor-decorated protofilaments also curve in the same direction, but remain stable and maintain a much larger radius (see Figure 5A and B). In addition, it has been shown that kinetochores in vitro can differentiate between GTP and GDP-tubulin within a microtubule 46 and it has been proposed that this is the mechanism by which they preferentially attach near the plus end. These results demonstrate, that proteins exist that can distinguish the GTP conformation of the microtubule lattice. Other, similar proteins are known, which localize to the microtubule plus end giving the appearance of tracking. 47 Furthermore, a bent tubulin configuration can be induced with Op18/Stathmin. 48,49 Very important in this context is the dynamics of the microtubule plus ends themselves. 30 By attaching the (plus) ends of microtubules to cellular structures, these structures are pushed around the cell according to the growing and shrinking of the microtubules. It is thought, that the different properties of the microtubule end are associated with specific end structures on which end-binding proteins can assemble to modulate the dynamic properties of microtubules. Furthermore, KinI Kinesins, microtubuledepolymerizing machines, have been shown to depolymerize microtubules by bending the underlying protofilaments. 50 Hence, there are a variety of different examples suggesting that different tubulin conformations in the cell are not only possible but also likely. Furthermore, different conformations are likely to be of physiological relevance, not only for growing and shrinking of microtubules, but also, as we show in this study, for microtubule motor interactions. According to our data, it is mainly a-tubulin that reacts upon motor binding. The structural change in this subunit, which we believe is a combination of an overall rotation and rearrangement, leads to a stabilization of the protofilament along the microtubules axis (Figure 8) thus stabilizing the track of the motor transporting its cargo. We should add, that lateral destabilizing effects might occur in conjunction, however, these effects, if present, are much smaller. One may speculate that the configuration found here in plain microtubules reflects that of a GDP-tubulin dimer, while that of motor decorated microtubules may more likely reflect a configuration of a GTP tubulin dimer. In our procedure we prepare microtubules well in advance prior to motor decoration. Hence, we can assume that upon motor decoration most of our microtubules already adapted a GDP-tubulin state, and the reversal to a GTP-like tubulin configuration is directly mediated by the motor interaction. It would certainly make sense for a molecular motor to stabilize the road it is traveling without having to change the nucleotide state. This however, would require that cooperative changes are traveling along the protofilament ahead of the motor. We do see effects as shown in Figure 5 already at sub-stoichiometric motor concentrations (ratio tubulin dimers/motors ¼ 2/1), thus cooperative effects might possibly occur. However, the test of the hypothesis, that conformational changes occur ahead of the motor (once bound to b-tubulin) towards the plus-end, would require microtubules, which are half-way decorated. The microtubules would need to be fully decorated towards the plus end, whereas towards the minus end no decoration should occur. This we have never been able to observe directly. If the motor concentration is decreased we observe partially decorated microtubules together with stabilized protofilaments. Helical reconstruction then leads to an averaged structure between the decorated and the undecorated state. Very interesting in this context is the question, whether Ncd (a minus end directed non-processive motor) would induce similar changes towards the minus end. So far, no structural changes inside the tubulin portion were observed with that motor. 51 This, however, might be due to the fact, that the resolution achieved in that study was not sufficient to detect such changes. In addition, at that time no structure of the plain microtubule with a and b-tubulin distinguishable was available. Nevertheless, we believe that similar structural changes might occur there as well. Different processivity and directionality is manifested in the neck-linker region and not in the binding region. Therefore we assume, that plus and minus directed motors have similar binding effects. Two different mechanisms may be present in the microtubules, allowing microtubules to react to environmental changes. One way, which uses GTP-hydrolysis and another one, in which microtubules interact with MAP s and motors. What we observe here appears to affect mostly the a-subunit, and not b-tubulin. Hence, it seems likely, that conformational changes induced by motor proteins are of a different kind as they are independent from nucleotide hydrolysis, and motor proteins predominantly affect the structure of a-tubulin, whereas GTP mainly affects the conformation of b-tubulin. However, since both, GTP and motor proteins bind predominantly to b-tubulin structural effects on a-tubulin may in any case be caused by allosteric or cooperative effects, which in both cases have a similar outcome:

12 150 Structural Changes in Microtubules Figure 8. Effects of motor binding on plain microtubules; schematic model. Initially an uneven high-density core with different conformations of a- and b-tubulin is present in plain microtubules. Although the motor itself binds to b-tubulin, 33 on the structural level mainly the a-subunit seems to be affected through its binding. A shift and rotation in combination with a rearrangement of a-tubulin is the key for the even high-density core, suggesting stabilized protofilaments in microtubule kinesin complexes. increasing the binding strength between two dimers. In any case, the molecular mechanisms behind protofilament stabilization, either through GTP-hydrolysis, or through motor domain interactions, could be related but not identical, as indicated by the different degrees of outward bending in protofilaments upon GTP-hydrolysis or motor domain interaction. Cooperation between two separate mechanisms may constitute a system responsible for the fine-tuning of microtubule stabilization according to cellular conditions. Conclusive remarks Our study indicates that microtubule dynamics goes beyond that of dynamic instability, and that their role in microtubule-based motility is more than just providing a sturdy track. We have new structural data that reveal the following: (A) It is possible to distinguish between the tubulin subunits in the microtubule lattice. (B) Microtubules themselves show distinct structural changes upon binding of motor head domains. (C) Most striking is the rearrangement of density within the a-tubulin subunit. (D) While there seem to be minor effects on lateral protofilament contacts, the most obviously affected contacts are the axial interdimer contacts, which become broader upon motor binding. The changes observed here may mimic a structural change as found with GTP hydrolysis in b-tubulin (the GTP-conformation is known to stabilize the caps in microtubules). In addition, microtubules appear to react to environmental changes with a second mechanism. In this one, the a-subunit is responsible for overall stability of protofilaments and microtubules, thereby reacting directly to motor binding by increasing axial tubulin dimer contacts. This change may induce a lesser outward bent into protofilaments as observed on plain microtubules upon depolymerization. The b-subunit, on the other hand, may be more directly involved in microtubule dynamic instability and changes the tubulin dimer conformation upon nucleotide hydrolysis, e.g. by inducing a strong outward bent into depolymerizing protofilaments. 27

13 Structural Changes in Microtubules 151 Materials and Methods Microtubules were polymerized in 80 mm Pipes, ph 6.8, 2 mm MgCl 2, 0.5 mm EDTA at a concentration of 2.5 mg/ml in the presence of 5% (v/v) DMSO, 20 mm taxol and 2 mm GTP at 37 8C for 20 minutes. Polymerized tubulin was washed twice by centrifugation in a centrifuge (Eppendorf Inc. Madison, WI), and resuspended in GTP-free buffer. Decoration of microtubules with Neurospora kinesin 355 was performed in solution at a final motor concentration of,20 mm with a 1:3 ratio of tubulin dimers/motor in the presence of 2 mm AMP-PMP. Samples were incubated for two minutes, adsorbed to holy carbon grids for one minute and quick-frozen in liquid ethane. Cryo-electron microscopy was performed on a CM200 FEG electron microscope (FEI-Company, Eindhoven, NL). Micrographs were recorded at a magnification of 38,000 on Kodak SO-63 films under extreme low dose conditions in KODAK D19. Images were taken at underfocus values ranging from 21.5 to 23.0 mm. Micrographs were digitized using a step size of 14 mm on a Zeiss SCAI scanner. Image analysis For helical analysis images with microtubules, which exhibit a Moiré, pattern corresponding to a 15 protofilament/2-start helical microtubule were used. Computational analysis was carried out mainly using routines from the program package SUPRIM, 52 PHOELIX, 53 the Brandeis helical package 54 and the program package SPIDER (Health Research Inc., Albany, NY). Briefly, tubes of up to 1.5 mm lengths were divided in portions of about mm lengths, computationally straightened and amplitudes and phases were extracted. In order to be able to deal with smaller parts of tubes and hence with much weaker diffraction pattern several routines of the standard software 52,53 were adapted for our purposes. Layer line amplitudes and phases were corrected for the modulations of the contrast transfer function, out of plane tilt and shift. Suitable references were determined and averaged maps were calculated. References were iteratively refined starting from a crude reference including data exclusively from strong layer lines of only a few selected data sets in the first round. The number of data sets in the average was increased step by step as was the number and extent of layer line data included in the calculation of the references used in the subsequent steps of refinement. Iterative rounds of refinement with improved references then lead to the final data set. Refinement was performed until the phase residuals between individual data sets and reference reached a stable value. Different approaches were tested using different reference data to align the individual data sets. During averaging data sets were checked for their orientation in the average calculating the individual phase errors for both directions, e.g. data sets may be up or down compared to the reference data. Only data sets with a sufficient difference in phase residual were included in further rounds of averaging and data sets, which showed different orientations in different rounds of refinement, were excluded. Different independent sub-averages were calculated with and without correction of the contrast transfer function and checked for consistency. Different averages, starting from different references with different refinement procedures, were calculated and checked for consistency. An estimate of the resolution was calculated by comparison of two independent sub averages by Fourier shell correlation using the program package SPIDER (Health Research Inc., Albany, NY). Statistical analysis and modeling The significance of density differences between 3D maps of native microtubules and decorated ones was assessed using statistical difference mapping and students t-tests 55 within the Brandeis package 54 and Phoelix. 53 Only significance levels of higher than 99% are discussed in the manuscript. In our docking the refined crystal structure of the tubulin dimers in zinc sheets was used 12 according to the overall position given in Nogales et al. 14 Fitting of the model electron densities into our EM reconstructions was performed using the program package URO. 41 The methodology is similar to the molecular replacement technique, adapted to take into account the phase information and the symmetry imposed during the EM reconstruction. Here we took advantage of the possibility to directly implement the helical symmetry of a 15-protofilament microtubule (selection rule 254/157) in the refinement process. The calculations were performed in real space and the normalized quadratic misfit of the correlation coefficient measured the quality of the fit. Data from 400 Å to 17 Å were used in the minimization process. Several optimization cycles were performed until only minimal shifts in the coordinates were observed. Initially the tubulin hetero dimer was fit, then the dimer was split and the two subunits were refined independently. Electron density maps were displayed in O 56 and VOLVIS (Research Foundation of the State University of New York, 1993). Ribbon diagrams were designed using Bobscript 57 and Ribbons (Ribbons 3.02 M. Carson, UAB/ CMC). Simulated maps were calculated using CCP4. 58 Acknowledgements This work was supported by the Austrian Academy of Sciences (APART 673 to A.K.). We thank Professor Eckhard Mandelkow and Dr Young Hwa Song for the Neurospora crassa kinesin motor domain. We are very thankful to Javier Borge for help in the usage of the program URO. References 1. Nogales, E. (2001). Structural insight into microtubule function. Annu. Rev. Biophys. Biomol. Struct. 30, Amos, L. A. (2000). Focusing-in on microtubules. Curr. Opin. Struct. Biol. 10, Hyams, J. S. & Lloyd, C. S. (1994). Microtubules, Wiley Liss, New York. 4. Amos, L. & Klug, A. (1974). Arrangement of subunits in flagellar microtubules. J. Cell Sci. 14, Mandelkow, E. M. & Mandelkow, E. (1985). Unstained microtubules studied by cryo-electron microscopy: substructure, supertwist and disassembly. J. Mol. Biol. 181, Sosa, H. & Milligan, R. A. (1996). Three-dimensional structure of ncd-decorated microtubules obtained by a back-projection method. J. Mol. Biol. 260,