journal of structural biology - uw-madison...eg5 is a highly conserved homotetrameric kinesin with...

Journal of Structural Biology 170 (2010) 257–265

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Journal of Structural Biology

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Cryo-electron tomography of microtubule–kinesin motor complexes

Julia Cope a, Susan Gilbert b, Ivan Rayment c, David Mastronarde a, Andreas Hoenger a,*a The Boulder Laboratory for 3-D Microscopy of Cells, University of Colorado at Boulder, Department of Molecular, Cellular, and Developmental Biology, Boulder, CO 80309-0347, USAb Rensselaer Polytechnic Institute, Biology Department—JRSC 1W14, 110 8th Street Troy, NY 12180-3590, USAc Department of Biochemistry, University of Wisconsin, Madison, WI 53706, USA

a r t i c l e i n f o

Article history:Received 3 November 2009Accepted 3 December 2009Available online 16 December 2009

Keywords:Cryo-electron microscopyCryo-electron tomographyKinesinMicrotubulesHelical 3-D reconstructionEg5Kar3Vik1

1047-8477/$ - see front matter � 2009 Elsevier Inc. Adoi:10.1016/j.jsb.2009.12.004

* Corresponding author. Fax: +1 303 735 0770.E-mail address: [email protected] (A

a b s t r a c t

Microtubules complexed with molecular motors of the kinesin family or non-motor microtubule associ-ated proteins (MAPs) such as tau or EB1 have been the subject of cryo-electron microcopy based 3-Dstudies for several years. Most of these studies that targeted complexes with intact microtubules havebeen carried out by helical 3-D reconstruction, while few were analyzed by single particle approachesor from 2-D crystalline arrays. Helical reconstruction of microtubule–MAP or motor complexes has beenextremely successful but by definition, all helical 3-D reconstruction attempts require perfectly helicalassemblies, which presents a serious limitation and confines the attempts to 15- or 16-protofilamentmicrotubules, microtubule configurations that are very rare in nature. The rise of cryo-electron tomogra-phy within the last few years has now opened a new avenue towards solving 3-D structures of microtu-bule–MAP complexes that do not form helical assemblies, most importantly for the subject here, allmicrotubules that exhibit a lattice seam. In addition, not all motor domains or MAPs decorate the micro-tubule surface regularly enough to match the underlying microtubule lattice, or they adopt conforma-tions that deviate from helical symmetry. Here we demonstrate the power and limitation of cryo-electron tomography using two kinesin motor domains, the monomeric Eg5 motor domain, and the hete-rodimeric Kar3Vik1 motor. We show here that tomography does not exclude the possibility of post-tomo-graphic averaging when identical sub-volumes can be extracted from tomograms and in both cases wewere able to reconstruct 3-D maps of conformations that are not possible to obtain using helical or otheraveraging-based methods.

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1. Introduction

Until now, most cryo-electron microscopy 3-D data on microtu-bules and microtubule–kinesin complexes have been produced byso-called averaging-based methods, mainly helical averaging (DeR-osier and Moore, 1970; for tubulin examples, see Sosa et al., 1997;Beuron and Hoenger, 2001; Wendt et al., 2002; Skiniotis et al.,2003; Kikkawa et al., 2000; Hirose et al., 2006; Sindelar and Down-ing, 2007), but also by single particle approaches (Li et al., 2002)and 2-D crystallography (e.g. to solve the structure of the ab-tubu-lin dimer itself: Nogales et al., 1998). Common to all these cases, 2-D projections of identical particles at various angles are collected,averaged and back-projected into 3-D space. Though this couldbe done in real space, it is often carried out in Fourier space wherephases and amplitudes can be assessed separately. The mostimportant prerequisites for averaging-based 3-D reconstructionsare (A) the averaged particle must be identical in shape and confor-mation, and (B) their projection angles relative to each other (so-called Euler angles) must be determined accurately. If this is

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. Hoenger).

achievable, averaging-based methods are very powerful for eluci-dating molecular structures and may yield close to atomic detail(tubulin: Nogales et al., 1998).

In most biological applications, however, many structures,even small macromolecular assemblies are often too flexibleand dynamic for averaging procedures. For these cases, the cur-rently most promising alternative to averaging-based 3-D recon-struction methods is cryo-electron tomography. Initially,tomographic 3-D reconstruction does not rely on averaging andtherefore can be applied to any specimen that can be imagedin an electron microscope, no matter how complex its structuremay be (reviewed in Lucic et al., 2005; Hoenger and McIntosh,2009). However, once a tomogram is completed, sub-volumescontaining identical particles may be extracted and further pro-cessed by averaging (Walz et al., 1997; Taylor et al., 2007; För-ster and Hegerl, 2007; Bartesaghi et al., 2008). To this end, wehave developed our own software called PEET (Particle Estima-tion for Electron Tomography: Mastronarde et al., in preparation,for example, see Nicastro et al., 2006). Here, we illustrate theadvantages and disadvantages of both helical reconstructionand volume averaging methods for microtubule–kinesin motordomain complexes.

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258 J. Cope et al. / Journal of Structural Biology 170 (2010) 257–265

The kinesin superfamily comprises a large and diverse group ofmolecular motors that interact with microtubules to perform awide variety of essential functions in the cell including roles inintracellular transport and cell division (reviewed in Hirokawaet al., 2009). Mitotic kinesins are a group of kinesins that are re-quired for cell division, playing a crucial role in assembly, mainte-nance and function of the mitotic spindle (reviewed in Wittmannet al., 2001). As such, they now constitute a new focus for studieson anti-cancer drugs and therapies targeted specifically to kinesinmotor function, either directly towards the motor domains (e.g.Monastrol: Maliga et al., 2002, 2006; Cochran et al., 2004, 2005;Crevel et al., 2004; Yan et al., 2004; Krzysiak et al., 2006), or toother functional parts such as cargo domains and receptors. Thisreport focuses on the motor domains of two structurally very dif-ferent kinesins with important roles in mitosis: the plus-end direc-ted kinesin-5 Eg5 (Kashina et al., 1996; X-ray structure: Turneret al., 2001) as a monomeric motor domain, and the minus-end di-rected kinesin-14 Kar3 (Meluh and Rose, 1990) as a functional het-erodimer with Vik1 (Manning et al., 1999; Barrett et al., 2000).

Eg5 is a highly conserved homotetrameric kinesin with anessential role in formation of the bipolar microtubule spindle andseparation of the chromosomes during mitosis (Blangy et al.,1995). Due to its bipolar structure, it is currently believed thatEg5 motors translocate to the plus-ends of microtubules at the on-set of mitosis and begin to organize microtubules by cross-linkingleading to the formation of the stable bipolar spindle seen in meta-phase (Kapitein et al., 2005). Once the spindle is in place, Eg5 pro-vides structural support and contributes to poleward flux bysliding the microtubules apart towards the centrosomes (Valentineet al., 2006). Structural studies are of interest to elucidate themechanical properties of this homotetrameric kinesin and its modeof action in spindle maintenance.

Kar3Vik1 is an unusual heterodimeric kinesin found in Saccha-romyces cerevisiae. Both components of the heterodimer have beenindividually crystallized and solved to about 2.5 Å resolution(Kar3: Gulick et al., 1998; Vik1: Allingham et al., 2007). Kar3Vik1is unusual because it has only one motor domain, Kar3, whoselocalization and activity is regulated through its association witha non-motor protein, Vik1 (Manning et al., 1999). Kar3Vik1 is ac-tive during vegetative growth where it localizes predominantlyat the spindle poles and is thought to cross-link microtubules, con-tributing to mitotic spindle assembly and stabilization (Manningand Snyder, 2000; Gardner et al., 2008). Interestingly, the recentlysolved crystal structure of the C-terminal globular domain of Vik1revealed that it has a fold remarkably similar to a kinesin motordomain, but does not have an active site for ATP hydrolysis (Alling-ham et al., 2007). Despite being unable to hydrolyze ATP, Vik1binds tightly to microtubules in vitro supporting a model whereboth Kar3 and Vik1 interact with the microtubule during its walk-ing cycle. Structural studies on Kar3Vik1 thus aim to identify a no-vel kinesin–microtubule interaction and may also provide insightinto the mechanisms of movement of other heterodimeric kinesinsthat have thus far been difficult to study (Scholey et al., 1996; DeMarco et al., 2001; Woehlke and Schliwa, 2007).

Tomography is not a new method for electron microscopy, butrecent technical advances on hardware and software have sincestrongly boosted its popularity (Frank, 2006). In particular, tomog-raphy on vitrified specimens is now realistic due to sensitive detec-tors and automated low-dose recording software. Here, we focusexclusively on cryo-electron tomography since only frozen-hy-drated specimens have the structural preservation required formolecular analysis. By using cryo-electron tomography we couldresolve molecular details in kinesin–microtubule complexes thatwould be impossible to image in 3-D by helical averaging. How-ever, even by volume averaging approaches the interpretablemolecular details of cryo-tomography compare to the lower-reso-

lution end of helically averaged maps. Hence, both methods arevery complementary. Helical averaging wins the resolution racebut sometimes loses important details due to averaging and sym-metry enforcement.

2. Materials and methods

2.1. Expression and purification of Eg5 and Kar3Vik1

2.1.1. Eg5 monomerThe monomeric motor domain of Eg5 was expressed and puri-

fied as described by Cochran et al. (2004).

2.1.2. Kar3Vik1 heterodimerHeterodimers of Kar3Vik1 were expressed and purified as de-

scribed by Allingham et al. (2006).

2.2. Microtubule polymerization

Microtubules were polymerized in vitro from 45 lM bovinetubulin (Cytoskeleton, Inc., Denver, CO) with BRB80 (80 mM PIPES,pH 6.8, 1 mM MgCl2, 1 mM EGTA) in the presence of 1 mM GTP,10 lM paclitaxel and 8% (v/v) DMSO for 30 min at 37 �C, and al-lowed to stabilize overnight at room temperature.

2.3. Sample preparation for cryo-electron microscopy

Eg5–microtubule complexes were formed in solution in the pres-ence of 2 mM AMP–PNP (Sigma–Aldrich Corp., St Louis, MO), at afinal tubulin concentration of 4.5 lM and a final Eg5 concentrationof 18 lm (see also Krzysiak et al., 2006). Complexes were adsorbedto holey-carbon C-flat grids (Protochips, Inc., Raleigh, NC) for 45 sand subsequently plunge-frozen into liquid ethane using a home-made plunge-freezing device.

Kar3Vik1–microtubule complexes were formed directly on holey-carbon C-flat grids to prevent bundling of the microtubules. Micro-tubules at a concentration of 4.5 lM were adsorbed onto grids for45 s and excess liquid was blotted away. Kar3Vik1 at concentra-tions ranging from 3.5 lM to 4.5 lM was incubated with themicrotubules for 2 min in the presence of 2 mM AMP–PNP andplunge-frozen as described above. Colloidal gold (10 nm) wasadded to all samples before plunging for use as fiducial markers.

2.4. Electron microscopy data collection

Samples were transferred under liquid nitrogen to a Gatan-626cryo-holder (Gatan, Inc., Pleasanton, CA). Cryo-electron microscopydata was collected on an FEI Tecnai F20 FEG transmission electronmicroscope (FEI Company, Eindhoven, The Netherlands) operatingat 200 kV. Images and tilt series were collected at a nominal mag-nification of 29 000� and a defocus of �2.5 lm (Kar3Vik1–micro-tubules complexes) or �6 lm (Eg5–microtubule complexes).Single frame images (Fig. 2A) were taken with an electron doseof 15 electrons/Å2. Tilt series consisting of about 80 low-doseimages, were collected by tilting the specimen between �60� and+60� (Figs. 1A, 3A, and 4A), or �70� and +70� (Fig. 5A), imagingat 1.5� increments. Images at higher tilts were recorded with ahigher dose to compensate for the increase in sample thickness.SerialEM software (Mastronarde, 2005) was used to automate thedata acquisition and minimize exposure of the specimen to theelectron beam. The total dose for each tilt series was about100 electrons/Å2. Images were recorded binned by two on a4K � 4K Gatan Ultrascan 895 CCD camera (Gatan, Inc.). With thiscamera, at a nominal microscope magnification of 29 000� theresulting pixel size corresponds to 7.6 Å on the specimen.

Fig. 1. Comparison between cryo-electron tomography and helical 3-D reconstruction of frozen-hydrated microtubules decorated with monomeric kinesin (Eg5) motordomains. (A) A 4.5 nm slice from a tomogram of microtubules decorated with monomeric Eg5 motors. Microtubules which are slightly tilted to the image plane expose asequence of their top, lumen, and bottom regions as they cut through the slice. The B-lattice pattern emphasized by the motor decoration shows a different angle along theBessel �2 helix according to the sketch. Since this is a left-handed helix, red lines show the angular orientation of the helical path according to the top view (towards theobserver), while the green lines indicate the course of the helix at the bottom of the microtubule. Occasionally a lattice seam, typical for non-helical microtubules with B-lattices, can be seen directly. The inset magnifies the framed area and reveals a bottom view of the seam, here made visible by the motor decoration. The seam is visible due tothe shift of one protofilament by 4 nm in the axial direction relative to the other (interruptions in green lines). (B) Cross-section through a 9 nm projection of a volumeaverage obtained by PEET combining 218 particles along one microtubule. This microtubule is composed of 14-protofilaments, which can be clearly seen despite the obviousloss of resolution along the z-axis (here top to bottom) due to anisotropy caused by the missing wedge of data. (C) The surface rendering of the average clearly shows thehelical path of the microtubule as well as the lattice seam typical for a 14-protofilament microtubule. (D–F) Helical averaging of microtubules is only possible if they are trulyhelical. This is the case for a 15-protofilament microtubule (red frame in (F)) but not for the more common 13-protofilament microtubules (yellow frame in (F)). Fourierfiltering (D and E) of the microtubules marked by frames reveals strikingly different 2-D projection patterns. 13-Protofilament microtubules (D) are perfectly aligned with thetubular axis while 15- (E–I), or 16-protofilament microtubules (Figs. 5 and 6) reveals a super-twisted arrangement. The supertwist is visible in the 3-D reconstruction in (F) asa deviation of the protofilaments (dotted green line) from the microtubule axis (solid green line). Kinesin motor decoration reveals a Fourier filtered image as shown in (G).The helical diffraction pattern (H) shows the helical layerline pattern where the layerlines around Bessel �2 (axial motor–tubulin dimer repeat) and �4 (axial a–b–a–b-tubulin repeat) form a cluster of 3–4 strong layerlines due to the convolution of the �2 and �4 helices with the protofilament supertwist. (I) The 3-D reconstruction revealsthe added kinesin heads as yellow densities, marking each ab-tubulin dimer.

J. Cope et al. / Journal of Structural Biology 170 (2010) 257–265 259

2.5. Tomogram reconstruction and sub-tomogram averaging

Tomograms were reconstructed from tilt series data byweighted back-projection using the IMOD software package (Kre-mer et al., 1996). Colloidal gold beads (10 nm) were tracked asfiducial markers to align the stack of tilted images.

To select sub-tomograms for averaging motor-decorated micro-tubules (Figs. 1, 3, 5, and 6), the center of the microtubule and thetrajectory along one of its protofilaments were modeled by handevery 15–30 nm. A program was run to fill in model points every8 nm along the microtubule axis (representing the length of anab-tubulin dimer repeat), and to determine the twist of the micro-tubule to produce a set of initial orientations for an alignmentsearch in the first round of averaging. The PEET software packagewas used to align and average repeating sub-volumes.

To average the Kar3Vik1 heterodimers in between two microtu-bules (Fig. 4), sub-volumes each containing one Kar3Vik1 ‘bridge’were modeled by hand and input into PEET for alignment andaveraging.

Isosurface rendering of the sub-volume averages was carriedout using IMOD (Figs. 5 and 6) or UCSF Chimera (Figs. 1, 3, and4) (Pettersen et al., 2004). A low pass filter was applied to all aver-ages to reduce noise before rendering.

3. Results and discussion

3.1. Cryo-electron tomography versus helical 3-D reconstruction onkinesin–microtubule complexes

The first structural analyses of microtubules date back to Amosand Klug (1974) who also produced the first 3-D reconstruction oftubulin protofilaments and model of a microtubule (Amos and Ba-ker, 1979) from zinc-induced 2-D crystalline tubulin sheets. How-ever, 3-D cryo-EM on microtubules picked up only after thedevelopments of cryo-EM methods (Dubochet et al., 1988) andthe discovery that microtubules composed of 15-protofilamentsmay form helical polymers (Fig. 1E–I; Wade and Chrétien, 1993;

Fig. 2. Cryo-electron microscopy of microtubules decorated with Kar3Vik1 heterodimers. (A) Frozen-hydrated microtubules decorated with a dimeric Kar3Vik1 constructreveals obvious cooperative microtubule-binding properties. The cooperative binding creates a heterogeneous decoration pattern where some microtubules show fulldecoration on one side while the opposing side is free of motor decoration (red frame). (Insets in A) Crystal structures of the Vik1 motor-homology domain (left), and the Kar3motor domain (right) are shown for comparison. Some structural similarities such as the central b-sheet surrounded by three a helices on each side are obvious. However,despite the similarity, Vik1 lacks a nucleotide-binding site. (B) Magnified and contrast-enhanced region of the frame in (A). The picture contrast has been inverted andboosted by an ‘‘embossing” filter to emphasize the visibility of the tubulin–Kar3Vik1 dimer complexes. Since the motors have been flushed with AMP–PNP, we believe thatthe domain in contact with the microtubule surface is the Kar3-MD while the tethered domain is Vik1-MHD. However, this remains to be determined experimentally.

Fig. 3. Dissection of microtubules partially decorated with Kar3Vik1 by cryo-electron tomography. (A) A 9 nm slice through a tomogram of microtubules decorated withKar3Vik1 heterodimers reveals features similar to those discussed in Fig. 1. Kar3Vik1 decorates microtubules in a cooperative fashion leaving empty patches besides fullydecorated areas. (B) Three individual 4.5 nm sections through the top, center, and bottom of the tomogram of the microtubule boxed in (A). The inset in the center panelshows an end-on view of the microtubule. While the bottom region shows strong striations every 8 nm corresponding to a fully motor-decorated surface (green lines) the topregion reveals empty microtubule protofilaments running axially. A striking advantage of tomographic 3-D reconstruction is observed in the center panel in that small singleevents such as individual missing motors (red circles) are revealed with much better clarity than in 2-D projections. These events would be lost after helical averaging due tothe symmetry constraints. (C) An isosurface representation from a PEET average of 146 particles selected from the microtubule in (B) gives a clear view of how the top of themicrotubule is free of motor decoration while the bottom and sides of the microtubule are completely decorated by KarVik1.


Arnal et al., 1996; Beuron and Hoenger, 2001) while most otherscarry a so-called lattice seam (Fig. 1A and C; Mandelkow et al.,1986; Kikkawa et al., 1994; Sandblad et al., 2006) that interruptshelical symmetry. The second boost came from the X-ray crystal

structures of monomeric and dimeric ncd (Sablin et al., 1996,1998) as well as monomeric (Kull et al., 1996) and dimeric kine-sin-1 (Kozielski et al., 1997). With these data at hand, moleculardocking became popular for kinesin–microtubule structures (Sosa

Fig. 4. Cryo-electron tomography of Kar3Vik1 dimers connecting between adjacent microtubules. (A) Eighteen nanometers x–z (top) and x–y (bottom) slices through atomogram showing the cross-section through two adjacent microtubules (top) and the Kar3Vik1 motors extending toward each other between the two microtubules(bottom). (B) An 18 nm projection and surface rendering of an average obtained by PEET from 44 sub-volumes selected from the tomogram in (A). The bridging motors appearto bend towards each other in a manner different from the regular radially extending conformations shown in Figs. 3 and 5. Since this is a highly artificial in vitro situation,the biological significance of these bridging motors is not clear. Rather, this exemplifies the way we are able to use cryo-electron tomography and subsequent volumeaveraging to analyze structural details that would not be possible by other averaging methods.

Fig. 5. Cryo-electron tomography and subsequent volume averaging of fully decorated Kar3Vik1–microtubule complexes. (A) Three nanometers thick tomographic x–y sliceof a microtubule decorated with Kar3Vik1 overlaid with the averaged volume shown in (B)–(E) and Fig. 6A and B. (B–D) Four nanometers x–y slices through the top and centerof an average of 99 particles selected from the tomogram in (A) according to the planes indicated in (E). The center slice in (D) cuts along the missing wedge, noticeable asreduced resolution compared to the x–y slice above. The central x–y slice (C) through the microtubule clearly shows densities corresponding to the a- and b-subunits oftubulin and to the two globular domains of Kar3 and Vik1 extending out from the microtubule. (E and F) End-on view (x–z) of the 3-D map before (E) and after rotationalaveraging over all protofilaments (F). The rotationally averaged map now looks indistinguishable from a helical reconstruction map at the same resolution. While the missingwedge effect is clearly visible before rotational averaging as the strong densities and smearing out along the z-axis, rotational averaging eliminates the missing wedgecompletely.


Fig. 6. Rotational averaging around the tubular axis effectively eliminates the missing wedge. (A and B) Surface renderings of the averaged 3-D map are shown in Fig. 5A–E.The cross-section reveals the number of protofilaments of this microtubule to be 16, which produces a perfectly helical microtubule (Bessel order�2) without seams (see (B)).(D and E) Surface rendered 3-D map after rotational averaging of the map in (A) (see also Fig. 5F). The missing wedge effect is eliminated. By rotationally averaging over all 16-protofilaments the asymmetric unit changes from axial slices along the microtubule axis to one ab-tubulin–motor domain complex. Hence, the theoretical maximum numberof asymmetric units included in the average is increased from 99 to 1584 (16 � 99). However, our average is based on the 1242 particles with the best correlation to thereference because this subset gave the highest resolution. (C and F) For Fourier-shell correlation calculations, the datasets were split in half on a random basis and the twohalves were correlated against each other. Fourier-shell correlation graphs obtained from the maps in A (C) and D (F) reveal resolution limits of 3.8 nm and 3.2 nm,respectively, based on the 50% correlation criterion. For FSC calculations a dataset is split in half on a random basis and correlated against each other. Accordingly the numberof asymmetric units in each group is maximally 49 before rotational averaging, and 621 afterwards.


et al., 1997) and many other macromolecular assemblies afterautomated procedures became available (Volkmann and Hanein,1999; Wriggers et al., 1999). Today the best resolved 3-D data onkinesin–microtubule complexes were obtained by helical averag-ing and show 8–9 Å detail (Hirose et al., 2006; Sindelar and Down-ing, 2007; Bodey et al., 2009).

The major advantages of helical reconstruction of microtubule–MAP complexes can be summarized as follows:

(A) Helical averaging produces low-noise data with isotropicresolution, sometimes to near-atomic detail. The averagingpower is very high as a large number of unit cells (50,000and more) can be included rapidly, yielding an excellent sig-nal to noise ratio. Diffraction (Fig. 1G) and Fourier filtering(Fig. 1D, E, and G) allow rapid assessments of the dataquality.

(B) Raw-data are obtained from single, untilted projectionswhich strongly facilitates corrections for the contrast trans-fer function (CTF) and allows the application of a high elec-tron dose (�40–100 electrons/Å2; Fig. 1F) at each imagesince each individual specimen is only recorded once.

The major constraints of helical averaging are:

(A) By definition, helical 3-D reconstruction requires a perfectlyhelical assembly (Fig. 1E–I) that is often either highly artifi-cial (e.g. in the case of kinesin–microtubule complexes themotor to tubulin ratio is much higher than under in vivoconditions (Fig. 1I)), does not reproduce native conforma-tions, or is simply impossible to obtain (structures too flex-ible and dynamic).

(B) Single molecular events such as lattice seams (Fig. 1A and C)or missing individual particles (Fig. 3B) cannot be observedin 3-D. If these events occur, they will be lost by being aver-

aged over all other events during the 3-D reconstructionprocess, potentially interfering with the accuracy of theresulting 3-D maps.

The emerging alternative method to study macromolecularassemblies without the constraints associated with helical recon-struction is cryo-electron tomography (Figs. 1A, 3A, B, 4A, and5A). As illustrated in Fig. 1A, kinesin–microtubule complexes canbe reconstructed in 3-D independent of the polymer compositionand can be analyzed for structural detail (such as the lattice seamor handedness of the helical path at distinct locations) that wouldnot be observable by a helical averaging approach. By focusing onthin slices through the tomogram, single motor domains, and theirempty spots when missing can be observed (Fig. 3B). Such singleevents would be difficult to detect on regular cryo-micrographsthat superimpose all the densities along the projection. 3-D datafrom cooperative decoration events (Figs. 2A and 3B) or bridgingmotor domains between two adjacent microtubules (Fig. 4) cannotbe obtained by helical averaging since they do not follow any heli-cal symmetry.

However, although no averaging is done with the raw-data and/or in the tomographic 3-D reconstruction procedure, averagingmay still be applied as a post-tomographic procedure, furtherenhancing the signal to noise ratio for particles which can be iden-tified within a tomogram at various orientations, but which areotherwise structurally reproducible enough to be averaged as en-tire volumes (Walz et al., 1997; Taylor et al., 2007; Förster and He-gerl, 2007; Bartesaghi et al., 2008). This is much morecomputationally demanding and only 10–15 years ago would nothave been realistic without the Linux clusters or multi-core com-puters that we use today. We are developing a dedicated softwaresuite for averaging volumes selected from tomograms called PEET(Particle Estimation for Electron Tomography: Mastronarde et al.,in preparation, for example, see Nicastro et al., 2006). With this


software we produced the averages presented in Figs. 1B, C, 3C, 4B,and 5A–F. Each average requires between 10 and 20 CPU hourssplit up over 35 processors on our Linux cluster. Typically the num-ber of particles that go into an average is much smaller than for ahelical approach. This reduces the averaging power, but 3-D parti-cles contain more structural information than 2-D projections,which facilitates accurate alignments.

The major advantages of tomographic reconstruction of micro-tubule–MAP complexes can be summarized as follows:

(A) Cryo-tomographic 3-D reconstruction is applicable to anyrandom structure and requires neither symmetry nor anystructurally reproducible sub-particles (unless post-tomo-graphic averaging should be applied). Cryo-tomographyworks well on plunge-frozen isolated macromolecularassemblies (as presented here) as well as on vitrified sec-tions (Norlén et al., 2009).

(B) Extraction of thin slices from tomograms reveals relativelynoise-free data that omits the convolution of projected datawith lower and upper structures (see Figs. 3B and 5B, C; theresult is somewhat similar to a confocal recording in lightmicroscopy).

(C) Single molecular events may be observed and analyzed (e.g.missing particles in an otherwise regular array; Fig. 3B).

(D) Post-tomographic data processing such as sub-volume aver-aging is facilitated by starting off with 3-D data rather than2-D projections and if successful, yields high signal to noise3-D data (see Figs. 1B, C, 3C, 4B, and 5B–F).

The major constraints of tomographic 3-D reconstruction are:

(A) Cryo-tomograms are composed of tilt series (typically 80–100 projections) recorded on the very same object. The res-olution in tomograms depends on the accuracy of alignmentand back-projection of each projection. Deviations from theassumed tilt angles, focus variations, and technical limita-tions such as bending support films reduce the resolutionin the final tomograms.

(B) The design of electron microscope stages and grid holdergeometries limit the recording range of tilt series to approx-imately �70� and +70� (rather than a full 180� rotation),leaving a missing wedge of data that results in anisotropicresolution (see Figs. 1B, 3C, 4B, 5E, and 6A and B) that is usu-ally lower than what is achieved with a helical approach.

(C) Cryo-tomograms have to be collected under strict low-doseconditions and therefore individual projections are extre-mely noisy, which may interfere with the alignment andback-projection process.

(D) While not impossible, CTF correction on tilt series isdemanding due to the various focus values in images fromtilted specimens (see Xiong et al., 2009).

3.2. Cryo-EM on Kar3Vik1–microtubule complexes reveals a highlycooperative binding pattern and a one-head-down and one-head-updimer-binding configuration

Kar3Vik1 is an unusual kinesin in the sense that it comprises atypical kinesin-14 motor domain (Kar3-MD) heterodimerized witha non-motor protein, the Vik1 motor-homology domain (Vik1-MHD) by a coiled-coil interaction. Conventional cryo-electronmicroscopy reveals a Kar3Vik1 binding property that is dominatedby a cooperative process (Fig. 2A). This can be seen directly as un-even decoration along microtubules where one side of some micro-tubules is completely free of motor decoration while the other sideappears fully decorated. Fig. 3 shows a cryo-tomographic 3-D

reconstruction of a partially decorated microtubule. Slices throughthe microtubule framed in Fig. 3A reveal the top portion to be voidof motors, while the bottom portion shows full decoration. Thecenter part reveals some missing individual motors along other-wise fully decorated protofilaments (circles in Fig. 3B).

Due to the partial decoration in the framed area in Fig. 2A andthe optimal side-view of the motors, the overall shape of the di-meric Kar3Vik1 construct is clearly visualized. Fig. 2B shows a con-trast-enhanced magnification of the framed region in Fig. 2A. Here,the three domains of the tubulin dimer, the Kar3-MD, and theVik1-MHD are well separated. The tomographic analysis shownin Fig. 5B confirms this finding. The motors have been flushed withAMP–PNP, a non-hydrolyzable ATP analog used to mimic an ATPbinding state. Under these conditions we found isolated Kar3-MDto be strongly bound to microtubules while Vik1-MHD’s boundwith much lower affinity. Accordingly, we assume that the domainin contact with the microtubule surface is the Kar3-MD while thetethered domain is Vik1-MHD. However, this remains to be deter-mined experimentally.

Both features of Kar3Vik1, the cooperative binding process(Wendt et al., 2002), and the binding conformation with one-head-down, one-head-up is highly reminiscent of dimeric ncd con-structs (Sosa et al., 1997; Hirose et al., 1998; Wendt et al., 2002;Endres et al., 2006) suggesting a common pattern for minus-enddirected motors and the members of the kinesin-14 family. Otherdimeric kinesin MD’s prefer to bind with both heads simulta-neously to the microtubule surface and do not display the strongcooperativity seen here and with dimeric ncd (kinesin-1: Hoengeret al., 1998, 2000; Eg5: Krzysiak et al., 2006).

3.3. The power of tomography in combination with volume averaging

Although tomographic 3-D reconstruction does not rely on anykind of averaging procedure, post-tomographic averaging consti-tutes a powerful tool to further enhance the signal to noise ratioin some parts of the tomogram. However, as in every averaging ap-proach, the structures subjected to averaging have to be identicalelse corrupted 3-D maps prone to misinterpretations and false con-clusions will be obtained. The advantage of post-tomographic aver-aging lies in the fact that 3-D structures are generally more easilyassessed than 2-D projections and often show more recognizablefeatures. The difficulty with post-tomographic averaging, however,comes from the missing wedge of data that reduces the resolutionparticularly in the Z-direction and the resulting elongation of struc-tures in Z potentially interferes with the 3-D alignment process.Our data on volume averaging presented here has been exclusivelyproduced by our software suite PEET (initial procedure describedin Nicastro et al., 2006; Mastronarde et al., in preparation). Themathematical and computational details of the averaging processwith PEET will be published elsewhere (Mastronarde et al., inpreparation).

In the case of kinesin–microtubule complexes the tomogram al-lows us to carefully inspect a microtubule for lattice defects andother irregularities. Once approved, we divide the microtubule intooverlapping axial segments about 130 nm long, at intervals of8 nm, making sure to include a full ab-tubulin repeat. While thelattice seams interrupt any helical symmetry, the sequence of cir-cular segments along the microtubule axis and each individualprotofilament are still identical. Once the segments have been se-lected, they are aligned according to the microtubule supertwist(if there is one) and averaged. The results are shown for a microtu-bule complexed with monomeric Eg5 (Fig. 1B and C; a total of 218particles were averaged), a microtubule partially decorated withdimeric KarVik1 (Fig. 3C; 146 particles), Kar3Vik1 dimers bridgingbetween two adjacent, parallel microtubules (Fig. 4B; 44 particles)


and a microtubule fully decorated with Kar3Vik1 dimers (Figs. 5A–E and 6A; 99 particles).

Two factors are of utmost importance here: (i) the usable tilt-range and (ii) the number of different angular views integrated.Even with a relatively large group of particles as shown in Fig. 1Band C where 218 particles have been averaged, some missingwedge effects remain visible such as a smeared-out density wallof badly separated protofilaments along the z-axis, while the x-and y-axes show very clear separations. The same is visible inFigs. 3C, 5E, and 6A. This is an intrinsic effect of any single-tilttomogram. For microtubule reconstructions that focus on 3-D datasurrounding the lattice seams this means no rotational averagingaround the tubular axis can be done (Fig. 1B and C) and the onlyrotation that partially weakens the missing wedge effect comesfrom an axial supertwist (see Fig. 1E and G). That supertwist iscompletely absent in 13-protofilament microtubules (seeFig. 1D). The microtubule in Figs. 5 and 6, however, is a helical16-protofilament microtubule where rotational averaging due tothe helical symmetry can be applied in addition to averaging axialsegments (Figs. 5F and 6D, E). We have done such averaging not byimposing rotational symmetry on the original average, but by sep-arately aligning each segment of the microtubule in 16 differentorientations. The same rotational averaging could be done withany tomographic reconstruction of a microtubule that does notpossess a seam, but only where features along protofilaments areidentical such as fully decorated motor–microtubule complexes.As shown in Figs. 5 and 6, rotational averaging immediately in-creases the number of asymmetric units by the number of protofil-aments (i.e. 16 � 99 = 1584). The asymmetric unit is now reducedfrom an entire tubular slice to a single ab-tubulin–motor domaincomplex. Accordingly, the Fourier-shell correlation graphs reveala significantly increased signal to noise ratio, and an interpretableresolution (i.e. visible detail not obscured by noise) improvementfrom 3.8 nm to 3.2 nm (Fig. 6C and F).

3.4. Molecular details in cryo-electron tomograms

Here, we demonstrate that cryo-electron tomography and heli-cal 3-D approaches (or any other averaging-based methods) arehighly complementary. We can conclude that cryo-electrontomography is an excellent method for any kind of assembly thatlacks apparent symmetry and that precludes helical, single particle,or 2-D crystalline averaging, such as the examples shown here inFigs. 1, 3, and 4. Even without post-tomographic averaging, struc-tural details can be interpreted to about 4 nm resolution. Forexample, individual protofilaments are spaced 5 nm laterally and4 nm axially, features that are clearly visible in tomographic slices(see Fig. 3B), at least when not obscured by the missing wedge ef-fect. Other structural features such as lattice angles and missingdomains can be assessed reliably. However, molecular docking ofsmall molecules such as kinesin motor domains remains difficultat the current resolution limits attainable by cryo-electron tomog-raphy. Volume averaging, where applicable currently pushes theinterpretable resolution close to 3 nm detail and may reach beyond2 nm in the future. Here, features such as the two domains of theab-tubulin dimer and the binding geometry of motor domains tomicrotubules can be reliably visualized (see Fig. 5C; and after rota-tional averaging: Figs. 5F and 6D, E). However, the theoretical res-olution attainable by post-tomographic averaging ultimatelydepends on the initial resolution that can be preserved from theraw images by tomographic reconstruction, which is the limitingfactor in this case. Therefore, even for highly symmetric assembliessuch as the helical 16-protofilament motor–microtubule complexshown in Figs. 5 and 6, reaching better than 2 nm isotropic resolu-tion via a cryo-electron tomography approach may remain out ofreach for a while longer.

Acknowledgments

J.C. was supported by the National Institutes of Health (NIH)/University of Colorado Molecular Biophysics Training Program(NIH T32 GM-065103). D.M. was supported by Grant NIH/NCRRP41-000592. We would also like to thank our colleague John Heu-mann for helpful discussions and assistance with the volume aver-aging software.

References

Allingham, J.S., Sproul, L.R., Rayment, I., Gilbert, S.P., 2007. Vik1 modulatesmicrotubule-Kar3 interactions through a motor domain that lacks an activesite. Cell 128, 1161–1172.

Amos, L.A., Baker, T.S., 1979. The three-dimensional structure of tubulinprotofilaments. Nature 279, 607–612.

Amos, L.A., Klug, A., 1974. Arrangement of subunits in flagellar microtubules. J. CellSci. 14, 523–549.

Arnal, I., Metoz, F., DeBonis, S., Wade, R.H., 1996. Three-dimensional structure offunctional motor proteins on microtubules. Curr. Biol. 6, 1265–1270.

Barrett, J.G., Manning, B.D., Snyder, M., 2000. The Kar3p kinesin-related proteinforms a novel heterodimeric structure with its associated protein Cik1p. Mol.Biol. Cell 11, 2373–2385.

Bartesaghi, A., Sprechmann, P., Liu, J., Randall, G., Sapiro, G., Subramaniam, S., 2008.Classification and 3D averaging with missing wedge correction in biologicalelectron tomography. J. Struct. Biol. 162, 436–450.

Beuron, F., Hoenger, A., 2001. Structural analysis of the mirotubule-kinesin complexby cryo-electron microscopy. Methods Mol. Biol. 164, 235–254.

Blangy, A., Lane, H.A., d’Hérin, P., Harper, M., Kress, M., Nigg, E.A., 1995.Phosphorylation by p34cdc2 regulates spindle association of human Eg5, akinesin-related motor essential for bipolar spindle formation in vivo. Cell 83,1159–1169.

Bodey, A.J., Kikkawa, M., Moores, C.A., 2009. 9-Angström structure of a microtubule-bound mitotic motor. J. Mol. Biol. 388, 218–224.

Cochran, J.C., Sontag, C.A., Maliga, Z., Kapoor, T.M., Correia, J.J., Gilbert, S.P., 2004.Mechanistic analysis of the mitotic kinesin Eg5. J. Biol. Chem. 279, 38861–38870.

Cochran, J., Gatial, J.I., Kapoor, T., Gilbert, S., 2005. Monastrol inhibition of mitotickinesin Eg5. J. Biol. Chem. 280, 12658–12667.

Crevel, I.M., Alonso, M.C., Cross, R.A., 2004. Monastrol stabilises an attached low-friction mode of Eg5. Curr. Biol. 14, R411–R412.

De Marco, V., Burkhard, P., Le Bot, N., Vernos, I., Hoenger, A., 2001. Analysis ofheterodimer formation by Xklp3A/B, a newly cloned kinesin-II from Xenopuslaevis. EMBO J. 20, 3370–3379.

DeRosier, D.J., Moore, P.B., 1970. Reconstructions of three-dimensional images fromelectron micrographs of structures with helical symmetry. J. Mol. Biol. 52, 355–369.

Dubochet, J., Adrian, M., Chang, J.J., Homo, J.C., Lepault, J., McDowall, A.W., Schultz,P., 1988. Cryo-electron microscopy of vitrified specimens. Q. Rev. Biophys. 21,129–228.

Endres, N.F., Yoshioka, C., Milligan, R.A., Vale, R.D., 2006. A lever-arm rotation drivesmotility of the minus-end-directed kinesin Ncd. Nature 439, 875–878.

Förster, F., Hegerl, R., 2007. Structure determination in situ by averaging oftomograms. Methods Cell Biol. 79, 741–767.

Frank, J. (Ed.), 2006. Electron Tomography: Methods for Three-DimensionalVisualization of Structures in the Cell. Springer Science + Business Media, LCC,233 Spring Street, New York, NY 10013, USA.

Gardner, M.K., Haase, J., Mythreye, K., Molk, J.N., Anderson, M., Joglekar, A.P.,O’Toole, E.T., Winey, M., Salmon, E.D., Odde, D.J., Bloom, K., 2008. Themicrotubule-based motor Kar3 and plus end-binding protein Bim1 providestructural support for the anaphase spindle. J. Cell Biol. 180, 91–100.

Gulick, A.M., Song, H., Endow, S.A., Rayment, I., 1998. X-ray crystal structure of theyeast Kar3 motor domain complexed with Mg.ADP to 2.3 Å resolution.Biochemistry 37, 1769–1779.

Hirokawa, N., Noda, Y., Tanaka, Y., Niwa, S., 2009. Kinesin superfamily motorproteins and intracellular transport. Nat. Rev. Mol. Cell Biol. 10, 682–696.

Hirose, K., Cross, R.A., Amos, L.A., 1998. Nucleotide-dependent structural changes indimeric NCD molecules complexed to microtubules. J. Mol. Biol. 278, 389–400.

Hirose, K., Akimaru, E., Akiba, T., Endow, S.A., Amos, L.A., 2006. Large conformationalchanges in a kinesin motor catalyzed by interaction with microtubules. Mol.Cell 23, 913–923.

Hoenger, A., McIntosh, J.R., 2009. Probing the macromolecular organization of cellsby electron tomography. Curr. Opin. Cell Biol. 21, 89–96.

Hoenger, A., Sack, S., Thormählen, M., Marx, A., Muller, J., Gross, H., Mandelkow, E.,1998. Image reconstruction of microtubules decorated with monomeric anddimeric kinesins: comparison with X-ray structure and implications formotility. J. Cell Biol. 141, 419–430.

Hoenger, A., Thormählen, M., Diaz-Avalos, R., Doerhoefer, M., Goldie, K., Mueller, J.,Mandelkow, E., 2000. A new look at the microtubule binding patterns of dimerickinesins. J. Mol. Biol. 297, 1087–1103.

Kapitein, L.C., Peterman, E.J., Kwok, B.H., Kim, J.H., Kapoor, T.M., Schmidt, C.F., 2005.The bipolar mitotic kinesin Eg5 moves on both microtubules that it crosslinks.Nature 435, 114–118.


Kashina, A., Baskin, R.J., Cole, D., Wedaman, K., Saxton, W., Scholey, J., 1996. Abipolar kinesin. Nature 379, 270–272.

Kikkawa, M., Ishikawa, T., Nakata, T., Wakabayashi, T., Hirokawa, N., 1994. Directvisualization of the microtubule lattice seam both in vitro and in vivo. J. CellBiol. 127, 1965–1971.

Kikkawa, M., Okada, Y., Hirokawa, N., 2000. 15 Å resolution model of the monomerickinesin motor, KIF1A. Cell 100, 241–252.

Kozielski, F., Sack, S., Marx, A., Thormählen, M., Schönbrunn, E., Biou, V., Thompson,A., Mandelkow, E.M., Mandelkow, E., 1997. The crystal structure of dimerickinesin and implications for microtubule-dependent motility. Cell 91, 985–994.

Kremer, J.R., Mastronarde, D.N., McIntosh, J.R., 1996. Computer visualization ofthree-dimensional image data using IMOD. J. Struct. Biol. 116, 71–76.

Krzysiak, T.C., Wendt, T., Sproul, L.R., Tittmann, P., Gross, H., Gilbert, S.P., Hoenger,A., 2006. A structural model for monastrol inhibition of dimeric kinesin Eg5.EMBO J. 25, 2263–2273.

Kull, F.J., Sablin, E., Lau, P., Fletterick, R., Vale, R., 1996. Crystal structure of thekinesin motor domain reveals a structural similarity to myosin. Nature 380,550–554.

Li, H., DeRosier, D.J., Nicholson, W.V., Nogales, E., Downing, K.H., 2002. Microtubulestructure at 8 Å resolution. Structure 10, 1317–1328.

Lucic, V., Forster, F., Baumeister, W., 2005. Structural studies by electrontomography: from cells to molecules. Annu. Rev. Biochem. 74, 833–865.

Maliga, Z., Kapoor, T.M., Mitchison, T.J., 2002. Evidence that monastrol is anallosteric inhibitor of the mitotic kinesin Eg5. Chem. Biol. 9, 989–996.

Maliga, Z., Xing, J., Cheung, H., Juszczak, L.J., Friedman, J.M., Rosenfeld, S.S., 2006. Apathway of structural changes produced by monastrol binding to Eg5. J. Biol.Chem. 281, 7977–7982.

Mandelkow, E.M., Schultheiss, R., Rapp, R., Müller, M., Mandelkow, E., 1986. On thesurface lattice of microtubules: helix starts, protofilament number, seam andhandedness. J. Cell Biol. 102, 1067–1073.

Manning, B.D., Snyder, M., 2000. Drivers and passengers wanted! The role ofkinesin-associated proteins. Trends Cell Biol. 10, 281–289.

Manning, B.D., Barrett, J.G., Wallace, J.A., Granok, H., Snyder, M., 1999. Differentialregulation of the Kar3p kinesin-related protein by two associated proteins,Cik1p and Vik1p. J. Cell Biol. 144, 1219–1233.

Mastronarde, D. N., in preparation.Mastronarde, D.N., 2005. Automated electron microscope tomography using robust

prediction of specimen movements. J. Struct. Biol. 152, 36–51.Meluh, P.B., Rose, M.D., 1990. KAR3, a kinesin-related gene required for yeast

nuclear fusion. Cell 60, 1029–1041.Nicastro, D., Schwartz, C., Pierson, J., Gaudette, R., Porter, M.E., McIntosh, J.R., 2006.

The molecular architecture of axonemes revealed by cryoelectron tomography.Science 313, 944–948.

Nogales, E., Downing, K.H., Amos, L.A., Lowe, J., 1998. Tubulin and FtsZ form adistinct family of GTPases. Nat. Struct. Biol. 5, 451–458.

Norlén, L., Oktem, O., Skoglund, U., 2009. Molecular cryo-electron tomography ofvitreous tissue sections: current challenges. J. Microsc. 235, 293–307.

Pettersen, E.F., Goddard, T.D., Huang, C.C., Couch, G.S., Greenblatt, D.M., Meng, E.C.,Ferrin, T.E., 2004. UCSF Chimera—a visualization system for exploratoryresearch and analysis. J. Comput. Chem. 25, 1605–1612.

Sablin, E.P., Kull, F.J., Cooke, R., Vale, R.D., Fletterick, R.J., 1996. Crystal structure ofthe motor domain of the kinesin-related motor ncd. Nature 380, 555–559.

Sablin, E.P., Case, R.B., Dai, S.C., Hart, C.L., Ruby, A., Vale, R.D., Fletterick, R.J., 1998.Direction determination in the minus-end-directed kinesin motor ncd. Nature395, 813–816.

Sandblad, L., Busch, E.K., Tittmann, P., Gross, H., Brunner, D., Hoenger, A., 2006. TheSchizosaccharomyces pombe EB1 homolog Mal3p binds and stabilizes themicrotubule lattice seam. Cell 127, 1–10.

Scholey, J.M., 1996. Kinesin-II, a membrane traffic motor in axons, axonemes, andspindles. J. Cell Biol. 133, 1–4.

Sindelar, C.V., Downing, K.H., 2007. The beginning of kinesin’s force-generatingcycle visualized at 9-Å resolution. J. Cell Biol. 177, 377–385.

Skiniotis, Y., Surrey, T., Altmann, S., Gross, H., Song, Y.H., Mandelkow, E., Hoenger, A.,2003. Nucleotide-induced conformations in the neck region of dimeric kinesin.EMBO J. 22, 1518–1528.

Sosa, H., Dias, P.D., Hoenger, A., Whittaker, M., Wilson-Kubalek, E., Sablin, E.,Fletterick, R., Vale, R.D., Milligan, R.A., 1997. A model for the microtubule–Ncdmotor protein complex obtained by cryo-electron microscopy and imageanalysis. Cell 90, 217–224.

Taylor, K.A., Liu, J., Winkler, H., 2007. Localization and classification of repetitivestructures in electron tomograms of paracrystalline assemblies. In: Frank, J.(Ed.), Electron Tomography. Springer, New York, pp. 417–439.

Turner, J., Anderson, R., Guo, J., Beraud, C., Fletterick, R., Sakowicz, R., 2001. Crystalstructure of the mitotic spindle kinesin Eg5 reveals a novel conformation of theneck-linker. J. Biol. Chem. 276, 25496–25502.

Valentine, M.T., Fordyce, P.M., Krzysiak, T.C., Gilbert, S.P., Block, S.M., 2006.Individual dimers of the mitotic kinesin motor Eg5 step processively andsupport substantial loads in vitro. Nat. Cell Biol. 8, 470–476.

Volkmann, N., Hanein, D., 1999. Quantitative fitting of atomic models into observeddensities derived by electron microscopy. J. Struct. Biol. 125, 176–184.

Wade, R.H., Chrétien, D., 1993. Cryoelectron microscopy of microtubules. J. Struct.Biol. 110, 1–27.

Walz, J., Typke, D., Nitsch, M., Koster, A.J., Hegerl, R., Baumeister, W., 1997. Electrontomography of single ice-embedded macromolecules: three-dimensionalalignment and classification. J. Struct. Biol. 120, 387–395.

Wendt, T.G., Volkmann, N., Goldie, K.N., Müller, J., Mandelkow, E., Hoenger, A., 2002.Microtubule binding patterns of the reverse kinesin motor Ncd reveal a minus-end directed power stroke. EMBO J. 21, 5969–5978.

Wittmann, T., Hyman, A., Desai, A., 2001. The spindle: a dynamic assembly ofmicrotubules and motors. Nat. Cell Biol. 3, E28–E34.

Woehlke, G., Schliwa, M., 2007. Kinesin Kar3 and Vik1 go head to head. Cell 128,1033–1034.

Wriggers, W., Milligan, R.A., McCammon, J.A., 1999. Situs: a package for dockingcrystal structures into low-resolution maps from electron microscopy. J. Struct.Biol. 125, 185–195.

Xiong, Q., Morphew, M.K., Schwartz, C.L., Hoenger, A.H., Mastronarde, D.N., 2009.CTF determination and correction for low dose tomographic tilt series. J. Struct.Biol. 168, 378–387.

Yan, Y., Sardana, V., Xu, B., Homnick, C., Halczenko, W., Buser, C.A., Schaber, M.,Hartman, G.D., Huber, H.E., Kuo, L.C., 2004. Inhibition of a mitotic motorprotein: where, how, and conformational consequences. J. Mol. Biol. 335, 547–554.

Cryo-electron tomography of microtubule–kinesin motor complexesIntroductionMaterials and methodsExpression and purification of Eg5 and Kar3Vik1Eg5 monomerKar3Vik1 heterodimer

Microtubule polymerizationSample preparation for cryo-electron microscopyElectron microscopy data collectionTomogram reconstruction and sub-tomogram averaging

Results and discussionCryo-electron tomography versus helical 3-D reconstruction on kinesin–microtubule complexesCryo-EM on Kar3Vik1–microtubule complexes reveals a highly cooperative binding pattern and a one-head-down and one-head-up dimer-binding configurationThe power of tomography in combination with volume averagingMolecular details in cryo-electron tomograms

AcknowledgmentsReferences

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