To study Kar3 motors at the single molecule level, we developed a protocol to express and purify full-length kinesin-14 heterodimers from Saccharomyces cerevisiae using affinity tagged Cik1 and Kar3 fused COOH-terminally to a HaloTag that served as covalent attachment site for the fluorescent dye tetramethylrhodamine (TMR). Purification and labeling yielded a homogenous preparation containing a heterodimer of Cik1 and TMR-labeled Kar3 (Figure 1B). During size exclusion chromatography, Cik1–Kar3 motors eluted as a single major peak with a Stokes radius of ∼9.1 nm, well separated from the void volume of the column (Figure 1C,D). Sucrose-gradient centrifugation revealed the presence of a single major species with an apparent sedimentation coefficient of ∼5.6S (Figure 1E). Combining these hydrodynamic values yielded a native molecular weight of 214 kDa, close to the calculated molecular weight of a Halo-tagged Cik1–Kar3 heterodimer of 190 kDa. We further characterized the oligomeric state of full-length Cik1–Kar3 motors by performing low-angle Pt/C rotary shadowing electron microscopy on peak fractions from the gel filtration experiments. This analysis revealed individual well-defined, highly elongated molecules that were characterized by globular domains separated by a 61 ± 8 nm (mean ± SD, n = 100) long coiled-coil (Figure 1F). Typically, two closely spaced globular domains likely corresponding to catalytic and non-catalytic head domains were discernible at one end, while a single globular domain decorated the other (Figure 1G). The highly elongated shape of the Cik1–Kar3 molecules can explain their early elution from the gel filtration column. Overall, the hydrodynamic analysis and the direct visualization of motor molecules by EM support the presence of heterodimeric Cik1–Kar3 molecules.

We next observed the behavior of single motor molecules on surface-immobilized microtubules in vitro using time-lapse multi-color TIRF microscopy. Unexpectedly, and contrary to the classification of kinesin-14s as non-processive motors, Kar3 molecules displayed efficient ATP-dependent movement over several micrometers and accumulated at microtubule minus ends (Figure 2A, Video 1). Automated tracking of motors revealed a Gaussian velocity distribution of Cik1–Kar3 with a mean speed of 77 ± 23 nm/s (mean ± SD; Figure 2B) in range with reported microtubule gliding velocities for truncated Cik1–Kar3 molecules (Chu et al., 2005; Allingham et al., 2007). The motor is therefore approximately 10-fold slower than conventional Kinesin-1, but similar in speed to yeast cytoplasmic Dynein, the major minus-end directed-motor in eukaryotes (Reck-Peterson et al., 2006). The run-length histogram followed an exponential distribution and revealed that individual Cik1–Kar3 motors advanced processively for an average of 5.2 μm before detaching from the microtubule track (Figure 2C). The motile parameters were highly sensitive to the ionic strength of the imaging buffer: at the same ATP concentration higher salt concentration increased the mean squared displacement of the motors (Figure 2D), but decreased the on-rate, run length and minus-end dwell time (Figure 2E). We additionally noticed that Cik1–Kar3 oscillated back-and-forth when entering microtubule overlap zones. Because Cik1–Kar3 is exclusively moving towards the minus-end on single MT filaments, we concluded that reversing motors encountered an antiparallel-oriented MT bundle. Individual motors were able to switch the track microtubule multiple times leading to a prolonged association with antiparallel bundles (Figure 2F, Video 2).

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The processive movement of Kar3 may either be a property of individual heterodimers or alternatively require the formation of motor ensembles combining multiple catalytic domains that coordinate stepping. Photobleaching experiments in the absence of nucleotide, in which the motor is persistently bound to the microtubule, revealed that 43 out of 50 molecules lost fluorescence in a single step consistent with the presence of a single Cik1–Kar3 heterodimer (Figure 2—figure supplement 1A–D). Mixing experiments combining TMR- with Alexa488 labeled Cik1–Kar3 molecules prior to imaging showed that the majority of moving complexes displayed exclusively either red or green fluorescence (Figure 2—figure supplement 1E, Video 3). To discriminate between processive and non-processive motility modes by an alternative approach, we varied the motor concentration in a standard microtubule gliding assay. Titration of Cik1–Kar3 over a 50-fold concentration range revealed no decrease in velocity for microtubule gliding, a characteristic feature of processive motility (Hancock and Howard, 1998) (Figure 2—figure supplement 1F). Contrary to previous reports, we did not observe substantial depolymerization of taxol-stabilized microtubules in the presence of Cik1–Kar3. On dynamic microtubules motors moved towards the minus-ends, overall microtubule dynamics appeared unchanged in the presence of Cik1–Kar3 and catastrophes did not coincide with plus-end localization of the motor (Figure 2—figure supplement 1G).

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Quantification of TMR brightness of moving motors allowed us to compare the motile properties of Cik1–Kar3 heterodimers vs larger teams that consisted of two or more heterodimers (Figure 2—figure supplement 2A–G). We found that Cik1–Kar3 velocity was largely independent of motor team size (Figure 2—figure supplement 2H,I). The run length, however, increased with larger team size while the variance of the velocity decreased (Figure 2—figure supplement 2J). These motile behaviors of Cik1–Kar3 complexes of different sizes can be quantitatively explained by a biophysical model in which individual motors influence each other through mechanical coupling with spring-like properties (Figure 2—figure supplement 3 and Supplementary file 1). Importantly, a key feature of the biophysical model is the ability of an individual Cik1–Kar3 heterodimer to move processively.

We next sought to establish the molecular requirements for Kar3 motility: processive movement could be a property intrinsic to the head domains or require secondary microtubule interaction sites in the motor tails (Gudimchuk et al., 2013; Su et al., 2013). To distinguish between these possibilities, truncation constructs were designed, systematically eliminating parts of the tail, coiled-coil, or globular domains of either the motor or the partner protein (Figure 3A). Truncation of the amino-terminal tail domain, either in Kar3 (up to aa 174) or in Cik1 (up to aa 250) allowed robust processive movement in the single molecule assay and also had little effect on multi-motor gliding velocity. This distinguishes Cik1–Kar3 from the homodimeric Drosophila kinesin-14, Ncd, which has been shown to be capable of a weakly processive motion at very low ionic strength depending on its tail region that acts as an electrostatic tether to microtubules (Furuta and Toyoshima, 2008). Further truncations of the coiled-coil interfered with heterodimer formation (not shown). In contrast, carboxyterminal truncations that either completely eliminated the predicted globular Cik1 motor homology domain (Cik1^1–360) or removed a portion of the carboxyterminus (Cik1^1–521) had severe effects and prevented directional movement. The specific nature of the defect was most apparent for the shorter truncation Cik1^1–521–Kar3, which was able to bind microtubules under our standard conditions, but instead of smooth translocation it displayed erratic forward and backward displacements that did not lead to directional movement as revealed by kymographs (Figure 3B, Videos 4 and 5). The defect imposed by the Cik1^1–521 mutation was also readily apparent in multi-motor gliding assays, where microtubules frequently switched their direction of movement and displayed little net transport (Figure 3C). The pronounced defect of the Cik1^1–521 mutant points to an essential role for the non-catalytic head in the motility mechanism. To corroborate this point, we also co-overexpressed Flag-tagged Kar3 with Kar3-Halo and purified fluorescently labeled Kar3 homodimers that can form in the absence of Cik1 (Chu et al., 2005). We failed to observe processive movement of Kar3-Kar3 combining two catalytic domains or of Kar3-Kar3^rigor complexes combining catalytically active and inactive Kar3 heads (Figure 3A). These results point to a specific role of the non-catalytic Cik1 head that cannot be simply replaced by a second catalytically active or inactive Kar3 head.

Figure 3

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To analyze the motile cycle in more detail, we imaged single Cik1–Kar3 motors in different nucleotide states by TIRF microscopy with high temporal resolution. In the absence of nucleotide (+Apyrase) and in the presence of the non-hydrolyzable ATP analog AMP-PNP Kar3 motors bound with high affinity to the microtubule but displayed no movement (Figure 4A). Binding under these conditions was sensitive to ionic strength with on-rate and lifetime of motor–microtubule interactions decreasing as the salt concentration was raised from 30 mM to 100 mM KCl (not shown). In the ADP state, motors displayed diffusive microtubule interactions (D = 0.061 ± 0.003 μm²/s) with short residence times (τ = 0.6 ± 0.1 s) (Figure 4B,C).

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What is the contribution of the non-catalytic domain to the translocation mechanism? To gain insights into this question, we quantitatively compared the microtubule-binding properties of single wild-type Cik1–Kar3 molecules with a mutant that lacks the globular Cik1 motor homology domain (Cik1^1–360–Kar3) (Figure 4D). In contrast to wild-type motors, the Cik1^1–360–Kar3 mutant displayed short-lived diffusive interactions with microtubules (Figure 4D). MSD data obtained in ATP for wild-type and mutant were fitted to the equation <x²> = a·tⁿ. n = 1 suggests standard diffusion and in this case, a = 2D, where D is the one-dimensional diffusion coefficient. For the mutant, an optimal fit was obtained with n = 1.01 ± 0.04, indicating unconstrained diffusion with no directional component. By contrast, n = 1.96 ± 0.03 was obtained for the wild-type motor, consistent with directional processive movement (Figure 4E). Further MSD analysis indicated that the mutant is about 20-fold more diffusive than the wild-type motor in the no nucleotide state (D = 0.01 ± 0.0006 μm²/s vs 0.00045 ± 0.00011 μm²/s) and about 30-fold more diffusive in the AMPPNP state (D = 0.0036 ± 0.00011 μm²/s vs 0.00011 ± 0.00001 μm²/s) (Figure 4F). Further analysis indicated that the Cik1^1–360 mutation also increased diffusion in the ADP state, but the difference was less pronounced in comparison to the other states (Figure 4G,H). We additionally constructed and purified a monomeric Kar3 head encompassing residues 353–729 with an N-terminal Halo-Tag. Under standard imaging conditions in the presence of ATP only very short-lived microtubule interactions without directional movement could be observed (Figure 4I). Taken together, these results indicate only a heterodimer containing a non-catalytic Cik1 head is able to move processively and a key contribution of the non-catalytic domain is to promote tight, non-diffusive binding of the motor in no nucleotide and AMPPNP states.

Having established that the non-catalytic partner is critical for achieving mechanically processive Kar3, we sought to gain further insights into the underlying mechanism by studying the alternative Vik1–Kar3 motor. The non-catalytic proteins Cik1 and Vik1 are paralogs that display about 45% sequence similarity between each other. We purified Vik1–Kar3 complexes from yeast extracts and imaged their interaction with microtubules under the same conditions as previously employed for Cik1–Kar3 (Figure 5A). Interestingly, Vik1–Kar3 also displayed highly processive movement, but with faster velocity compared to Cik1–Kar3. Under standard assay conditions, Vik1–Kar3 complexes moved at 234 ± 29 nm/s towards microtubule minus-ends where they strongly accumulated (Figure 5B, Video 6). To ask how these differences might be determined by the non-catalytic partner, we constructed a chimeric protein in which the globular motor homology domain and the neck of Vik1 (aa 351–647) were fused to the tail domain of Cik1 (aa 1–353) (Figure 5C). The Vik1–Cik1 chimera formed a stable heterodimer with Kar3-HALO and supported movement with 188 nm/s to the minus-end, significantly faster than Cik1–Kar3 and approaching the velocity of Vik1–Kar3 (Figure 5D,E). Thus, the globular non-catalytic domain is a key determinant for setting the velocity of the motor. The engineered chimeric protein was able to functionally replace Cik1 in cells as demonstrated by its ability to rescue the phenotypes of a Cik1 deletion and to support growth at wild type level under all tested conditions (Figure 5F).

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In addition to directly controlling the motile characteristics of individual Kar3 motors, the non-catalytic partners may also provide functional diversification by allowing differential interactions with regulatory proteins or cargos. To identify such interactors, we performed affinity purifications of Cik1–FLAG at different cell cycle stages and analyzed associated proteins by mass spectrometry. In addition to Kar3, we reproducibly mapped two plus-end tracking proteins Bim1 and Bik1, the EB1 and CLIP-170 homologues of budding yeast, as well as the microtubule-binding Ndc80 kinetochore complex (Figure 6A). By performing pull-down assays with recombinant proteins, we established that Cik1–Kar3, but not Vik1–Kar3, directly interacted with Bim1 (Figure 6B). The binding interface involved the C-terminal EB homology domain of Bim1 and the aminoterminal tail domain of Cik1 (Figure 6—figure supplement 1A–D). A physical interaction with Bim1 may account for the previously observed microtubule plus-end localization of Cik1–Kar3 in vivo (Sproul et al., 2005; Gardner et al., 2008). Live cell imaging showed that the localization of Kar3 to distinct foci along the yeast spindle, as well as to the tips of shmoo-tip-directed microtubules in alpha-factor-arrested cells (Figure 6—figure supplement 1E), was abolished in a bim1Δ strain, pheno-copying a cik1 deletion (Manning et al., 1999) (Figure 6C,D,E). Consistent with our biochemical experiments, spindle localization was maintained in the Cik1^1–360 and Cik1^1–521 mutants. Thus, the Cik1 tail domain specifies the differential localization of Kar3 in vivo by allowing a direct interaction with Bim1.

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The insights into the localization determinants of the Kar3 motor allowed us to evaluate the key roles of Kar3 in the cell: yeast strains expressing Cik1^1–360 or Cik1^1–521 from the endogenous chromosomal locus displayed slow growth at 25°C and were inviable at 37°C, similar to a cik1 deletion (Figure 6F). By contrast neither a bim1 deletion, which eliminates spindle localization, nor a vik1 deletion, removing spindle pole localization, displayed a pronounced growth phenotype (data not shown). Together with our finding that the Vik1^head–Cik1^tail chimera fully supports viability, this implies that the key function of Kar3 in vegetatively growing yeast cells requires motility—supported by any of the two non-catalytic heads—and the Cik1 tail domain, but does not involve spindle localization via Bim1. Based on previously characterized mutants and on our finding that the Ndc80 complex co-purifies with Cik1, this supports a key role for Kar3 at the kinetochore. Consistent with this notion, we found that 80% of Cik1^1–521 cells arrested as large budded cells upon shift to the restrictive temperature, indicative of mitotic checkpoint activation. Analysis of chromosome segregation at the semi-permissive temperature of 25°C by fluorescent labeling of chromosome V showed that 50% of large budded Cik1 mutant cells attempted nuclear division without proper bi-orientation of sister chromatids (Figure 6G). We conclude that the key contribution of the non-conventional Cik1 motility mechanism for cell proliferation lies in the promotion of sister chromatid bi-orientation in mitosis.

To ask whether Cik–Kar3 can act as a transport motor, we first performed bulk in vitro recruitment assays in the absence of nucleotide using purified motor proteins and recombinant, fluorescently labeled Ndc80 complex as the candidate kinetochore binding partner revealed in the mass spectrometry experiments. In the absence of motor, the Ndc80 complex only weakly associated with taxol-stabilized microtubules under standard conditions (Figure 7A,B). Vik1–Kar3 motors strongly decorated microtubules in the absence of ATP, but had no effect on the Ndc80 complex. By contrast, combining Cik1–Kar3 and the Ndc80 complex resulted in decoration of microtubules with both molecules. Upon lowering protein concentrations, Cik1–Kar3 and the Ndc80 complex co-localized to distinct spots on the microtubule where their intensities were correlated (Figure 7C). Inclusion of ATP initiated co-transport of the Ndc80 complex and Cik1–Kar3 towards the minus end (Figure 7D, Video 7). While transporting the Ndc80 complex, the average speed of the motor was 76 ± 15 nm/s (mean ± SD), indicating that cargo binding did not significantly impede movement. The value is also in range with the reported velocity for kinetochore transport in vivo (Tanaka et al., 2005). Microtubule binding by the Ndc80 complex was not required for efficient transport, as shown by effective recruitment and translocation of the microtubule-binding defective calponin-homology domain mutant K122E K204E (Lampert et al., 2013) (Figure 7—figure supplement 1). This result is in agreement with the observation that Kar3 localizes to unattached kinetochores prior to microtubule binding. We conclude that the Ndc80 complex can be transported by the Kar3 motor in vitro and that in addition to promoting processive motility a key function of Cik1 may lie in the differential binding of this complex.

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Our study provides direct evidence for the processivity of Kinesin-14 motors by analyzing full-length yeast Kar3 motors on the single-molecule level. Contrary to other kinesin-14s, we demonstrate that individual Cik1–Kar3 motors can move processively towards the minus-end and show that the non-catalytic Cik1 head domain is functionally required for this activity. We further demonstrate that the combination of Kar3 with two different non-catalytic partners generates motors with different motile properties and also allows the differential binding of partner proteins such as Bim1 and the Ndc80 complex (Figure 8A).

Figure 8

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The demonstration of Cik1–Kar3 processivity presented here provides evidence that effective translocation along microtubules does not strictly require the conventional hand-over-hand walking mechanism that has been established for conventional Kinesin-1. So far this point has been most clearly demonstrated for the Dynein motor, as recent studies have shown that cytoplasmic dynein is capable of processive movement in vitro despite inactivating mutations in one of the two force-generating AAA ring domains (DeWitt et al., 2012; Qiu et al., 2012), or even as a combination of one active head with a second passive microtubule-binding domain (Cleary et al., 2014).

Also several kinesin motors have been reported to move in violation of the hand-over-hand model: monomeric constructs of the kinesin-3 KIF1A have been shown to move along microtubules in vitro using a biased diffusion mechanism (Okada and Hirokawa, 1999), but the in vivo function of these motors probably involves dimeric molecules for effective cargo transport (Tomishige et al., 2002; Endres et al., 2006). We could not detect evidence for processive movement of monomeric Kar3, making it unlikely that it works via a KIF1A-type mechanism. Among the kinesin-14 family, Ncd can exhibit processive motility but requires at least two homodimers coupled together (Furuta et al., 2013). A number of studies have shown that inactivating mutations in one of the two heads of conventional kinesin can still allow residual processivity (Subramanian and Gelles, 2007; Thoresen and Gelles, 2008; Kaseda et al., 2003). We note, however, that in these examples processive movement of the mutant heterodimeric kinesins was compromised compared to wild-type homodimers. By contrast, Kar3 does not simply tolerate the loss of a second active head but instead it has evolved a specific requirement for the non-catalytic domain in order to move processively. In other words, Kar3 can only walk with a combination of an active and an inactive ‘leg’.

A precise elucidation of the stepping mechanism of Kar3 motors will require further biophysical experiments. Based on the results obtained in this and in previous studies, we propose a working model that describes the non-catalytic head as a ‘foothold’ for Kar3. Based on recent structural data Kar3 and its partner non-catalytic head may be located on adjacent protofilaments (Rank et al., 2012), where the non-catalytic domain could contribute to hold the motor complex in place and prevent diffusion. During its ATP cycle, the motor goes through rounds of tightly and weakly bound states. When ADP is bound, the motor is in its weakly bound state which allows for one-dimensional diffusion along the microtubule lattice. On its own, movement in this state would lack directionality and an additional mechanism providing minus-end directed bias is required. We speculate that the presence of Cik1 in combination with the kinesin-14 typical stalk rotation upon ATP binding by Kar3 (Endres et al., 2006; Gonzalez et al., 2013) might provide such a mechanism either by shifting the overall position of the molecule or by allowing subsequent directed binding/unbinding (Figure 7C). Because of the combination of one foot on the track and a diffusive movement to the next binding site, we refer to this possible mechanism as the ‘skateboard’ model. This model can explain a number of our experimental observations: the effect of raising the ionic strength is that initially motor movement occurs with faster velocity, probably by increasing the diffusion term. At some point however, stable binding of the non-catalytic domain is prevented and directional displacement is disrupted, similar to the situation in which the motor lacks the motor homology domain. In addition, diffusive movement in the ADP state and the observed large variation in movement rates are compatible with the model. The biochemical properties of the non-catalytic head binding to the microtubule will have profound effects on the motile characteristics as evidenced by the different properties of Cik1–Kar3 and Vik1–Kar3 motors.

One of the open questions regarding the mechanism of processivity is how the motor homology domain can provide ‘footing’ in no nucleotide and AMPPNP states yet allow effective diffusion in the ADP state. In agreement with previous work, this suggests that similar to conventional kinesins, a form of head-to-head communication must occur in Kar3 motors such that the catalytic head influences the microtubule affinity of the motor homology domain (Allingham et al., 2007; Chen et al., 2011; Duan et al., 2012; Rank et al., 2012). Such coordination may involve intramolecular strain communicated by mechanical elements between the heads as has been reported for conventional kinesin (Yildiz et al., 2008).

Based on our experiments and the previously reported rates of microtubule shortening in the presence of Cik1–Kar3, we consider it unlikely that Kar3 functions as a microtubule depolymerase in vivo. We note that none of the Kar3 functions poses a strict requirement for such an activity and indeed recent experiments analyzing Kar3 function during karyogamy support the view that its primary function is transport, not shortening (Gibeaux et al., 2013).

What are the implications of Kar3's processivity? The ability to move processively may be used in the cell to enrich Kar3 motors at minus-ends. In addition, Kar3 could work more effectively in small teams if it can undergo multiple catalytic cycles before releasing from the microtubule. This may be especially important during kinetochore transport, where Ndc80 complexes can provide only a limited number of binding sites for the motor. Kar3 heterodimers have probably evolved from conventional Kinesin14-homodimers that performed Ncd-like roles in spindle organization. We speculate that with the exclusion of dynein from the budding yeast nucleus additional functional requirements for minus-end directed motility in spindle assembly and kinetochore function arose that could be fulfilled with the ‘invention’ of the Cik1 and Vik1 non-catalytic domains. While commonly perceived as conferring a loss of functionality, the pseudo-motor domains have the ability to convert a kinesin-14 into a processive motor and additionally create functional diversity through allowing differential interactions with partners. Together this allows Kar3 to function as a processsive sliding and transport motor that substitutes for key roles of dynein in the yeast nucleus. Other examples for heterodimeric motors exist in the Kinesin-2 family with different subunits conferring distinct activities (Brunnbauer et al., 2010). Kar3 motors may therefore constitute an extreme example for such diversification strategies that could be more widely used in other motor families. More generally, we note that pseudoenzymes have emerging roles in other cellular contexts, for example in the form pseudo-GTPases during human kinetochore assembly (Basilico et al., 2014), or as pseudo-kinases and pseudo-phosphatases in cell signaling (Boudeau et al., 2006; Tonks, 2009). This may suggest that the use of catalytically inactive enzyme derivatives could be a more widespread strategy employed by the cell. The establishment of an in vitro assay for Ndc80 transport serves as a starting point for a detailed analysis of the function and regulation of kinetochore motility. Furthermore, the unique design principle of the Kar3 motor, which allows the same catalytic domain to be paired with different non-catalytic heads, generates functional diversity that may also be exploited in nanotechnological applications.

The protein coding sequences of S. cerevisae (S288c) full-length Kar3, Cik1, and Vik1 were amplified by PCR and cloned into the overexpression pESC-TRP (Agilent Technologies, Santa Clara, CA) vector. The non-catalytic motor subunit was tagged C-terminally with 1xFLAG and the motor itself was fused to a HaloTag (DHA, Promega) at the 5′ end of the coding sequence, separated by a 13 amino acid linker. Mutated and truncated versions of Cik1 and Kar3 were generated by site-directed mutagenesis PCR (Phusion, Thermo Scientific). To determine the TMR-labeling efficiency, Halo-Kar3 was fused with enhanced green fluorescent protein (eGFP), creating Halo-eGFP-Kar3. All motor proteins were overexpressed in budding yeast using the pESC vectors (Agilent Technologies) following the manufacturer's instructions. In brief, yeast cells harboring the respective pESC plasmid were induced with 2% Galactose at OD₆₀₀ = 1.0 for 7–9 hr, harvested by centrifugation, washed, and frozen as droplets in liquid nitrogen. Lysis was conducted in liquid nitrogen using a freezer mill (Biospec Inc.). The cell powder was resuspended in lysis buffer (25 mM Hepes [7.4], 300 mM NaCl, 1 mM MgCl₂, 5% glycerole, 0.1 mM EDTA, 0.5 mM EGTA, 0.1% Tween-20, 0.01 mM ATP, 0.1 mM PMSF supplemented with PhosSTOP Phosphatase Inhibitor Cocktail [Roche]). The lysed cells were centrifuged twice, first at 43.146×g for 20 min and afterwards at 125.749×g for 1 hr. The resulting supernatant was incubated with M2 affinity agarose (Sigma–Aldrich) for 1 hr, gently rotating at 4°C. The agarose resin was washed five times with lysis buffer (adjusted to 150 mM NaCl, 0.09% Tween-20, omitting the PhosSTOP reagent). Elution of the kinesin heterodimeric constructs from M2 agarose was achieved by applying one resin volume of 3xFLAG peptide at final 2 mg/ml in lysis buffer (adjusted to 250 mM NaCl, 1 mM DTT, 0.09% Tween-20, omitting ATP and PhosSTOP). If needed, the elution was loaded onto a cation exchange chromatography (MonoS 5/50 GL, GE Healthcare) in running buffer (10 mM Hepes [pH 7.2], 150 mM NaCl, 1 mM MgCl₂, 1 mM DTT, 1 mM EGTA) to remove the FLAG peptide. Afterwards a linear salt gradient eluted a single peak, pure motor fraction at 250 mM salt. Elution fractions were supplemented with glycerol (final concentration: 5%), snap-frozen in liquid nitrogen and stored at −80°C. The protein concentration was measured using the DC Assay kit (Bio-Rad). All proteins were pre-cleared by centrifugation using a 0.1-μm spin filter (Millipore) to remove aggregates before each experiment.

Labeling of the NH₂-terminus of Kar3 with the HaloTag TMR ligand or HaloTag Alexa488 ligand (Promega, Madison, WI) was performed during the above-described purification before eluting the kinesin heterodimer from the M2 affinity agarose: the proteins were incubated with 10 μM HaloTag ligand for 3 hr. Extensive washing removed unbound TMR ligand and the kinesin was eluted as described before. In order to assess the labeling efficiency the Halo-eGFP-Kar3 construct was expressed, purified, and TMR-labeled. Observing the motor in our multi-color single-molecule imaging setup in the absence or presence of ATP revealed that >90% of eGFP-labeled kinesins also had a TMR-ligand covalently bound to the HaloTag.

To obtain a stoichiometric heterodimer of Cik1 and Kar3, the motor was subjected to analytical SEC. The purified motor was loaded onto a Superose 6 PC 3.2/30 column (GE Healthcare), and 100 μl fractions were collected and separated by SDS-PAGE. Proteins were stained with Coomassie Brilliant blue R250.

Expression and purification of the full-length and mutant Ndc80 complex (Ndc80p-6xHis/Nuf2p-EGFP and Spc24p/6xHis/Spc25p) was performed as described previously (Lampert et al., 2013; Lampert et al., 2010). Bim1 was expressed and purified as described previously (Zimniak et al., 2009).

Peak fractions of motor from the gel filtration experiments were diluted to a final concentration of 60 μg/ml in spraying buffer, containing 100 mM ammonium acatate and 30% (vol/vol) glycerol, pH adjusted to 7.4. After dilution, the samples were sprayed onto freshly cleaved mica chips and immediately transferred into a Bal-Tec MED020 high vacuum evaporator equipped with electron guns. After drying in the vacuum, the rotating samples were coated with 0.6 nm Platinum at an elevation angle between 5° and 6°, followed by 9.5 nm Carbon at 90°. The produced replicas were floated off from the mica chips and picked up on 400 mesh Cu/Pd grids. The grids were inspected in an FEI Morgagni 268D TEM operated at 80 kV. Electron micrographs were acquired using an 11 megapixel Morada CCD camera from Olympus-SIS. Images were examined and analyzed using ImageJ.

Single-step purified Cik1–Halo–Kar3 was loaded on top of a 4.4 ml 5–25% linear sucrose gradient and spun at 50,000 rpm for 16 hr using an Sorvall TH-660 rotor and a Sorvall Discovery 90SE centrifuge. Fractions (270 μl) were collected and analyzed by SDS-PAGE and Coomassie Brilliant Blue R250 staining. The sedimentation value for Cik1-Halo-Kar3 was defined by comaring the mobilities of the motor with linear plots of mobility standards.

The single molecule motor assays were conceptually designed as described previously (Korten et al., 2011; Lampert et al., 2010; Bieling et al., 2010). Biotin-PEG-SVA- and mPEG-SVA-functionalized coverslips (Laysan Bio) were prepared as described (Lampert et al., 2010; Jain et al., 2012). Coverslips were assembled onto passivated glass slides using double-sided tape creating a flow chamber. First, a solution of 1 mg/ml avidin DN (Vector Laboratories) was applied to the chamber for 30 min and exchanged for 1% pluronic F-127 (Sigma–Aldrich) in BRB80 (Zimniak et al., 2009; Nitzsche et al., 2010). Porcine-derived HiLyte-647-labeled, biotinylated and taxol-stabilized microtubules (MTs) were immobilized (labeled and biotinylated tubulin source: Cytoskeleton Inc., unlabeled tubulin was purified from pig brains as described previously [Ashford and Hyman, 2006]) and excess of MTs was washed out with BRB80 buffer supplemented with 5 μM taxol, 0.5% (vol/vol) β-mercaptoethanol, 4.5 μg/ml glucose, 200 μg/ml glucose-oxidase, and 35 μg/ml catalase. Single molecule imaging was performed by introducing the TMR- or Alexa488-labeled kinesins at low nanomolar range in assay buffer (BRB80, 0.33 mg/ml casein, 16.6 μM taxol, 0.13% [vol/vol] methylcellulose, 0.5% [vol/vol] β-mercaptoethanol, 4.5 μg/ml glucose, 200 μg/ml glucose-oxidase and 35 μg/ml catalase, 0.06% [vol/vol] Tween-20, the indicated amount of KCl and the respective amount of nucleotide) into the flow cell. Time-lapse videos were recorded at 28°C in 3 s intervals between frames (if not annotated differently) using a TIRF microscopy setup described previously (Lampert et al., 2010). Multi-color imaging was achieved by the use of an external filterwheel (Ludl Electronic Products Ltd.). Each channel (excitation: 488 nm, 561 nm, 639 nm) was exposed for 100 ms at every time-interval, recorded by a Cascade II EMCCD camera and projected to two-dimensional images (software: Metamorph [Molecular Devices], ImageJ). For photobleaching analysis, the oxygen-scavenger mix was omitted (β-mercaptoethanol, glucose, glucose-oxidase, catalase), and images were recorded at maximum laser power. Videos are represented as kymographs (time-space plot) or as example single frame (software: MetaMorph [Molecular Devices]).

High temporal resolution recordings (as presented in Figure 2) have been obtained using custom-made TIRF microscope based on Olympus IX-71 body and Coherent CUBE lasers in a temperature stabilized room (21 ± 0.1°C). Images were acquired using a Andor iXon3 897 EMCCD camera and subsequently analyzed using custom-made software written in MATLAB (MathWorks, Inc).

The microtubule gliding assay was designed as described before (Nitzsche et al., 2010). For this assay, hydrophobic coverslips were prepared according to the following scheme: sonication in acetone for 15 min was followed by sonication for 15 min in ethanol. Coverslips were incubated 1 hr in boiling Piranha solution and rinsed with a lot of water afterwards. Then rinsed with 0.1 M KOH, MilliQ and dried with nitrogen and immersed in 5% dichlorodimethylsilane in heptane for 1 hr at room temperature. Coverslips were rinsed again with MilliQ and sonicated for 5 min, sonicated in chloroform for 5 min and air-dried.

Motor proteins were attached to the hydrophobic coverslips via application of 0.2–20 μg/ml anti-Halo antibody (Promega) to the flow chamber. After the excess of antibody was washed out with BRB80 supplemented with 0.5 mg/ml casein, motors were introduced to the chamber and incubated for 5 min. Finally a microtubule-containing solution supplemented with 5 mM ATP and oxygen scavenger mix was perfused into the chamber. Microtubules used for this experiment were assembled from porcine tubulin mixed together with HiLyte-647-labeled tubulin (Cytoskeleton Inc.) in the presence of 10 μM taxol. Time-lapse videos were recorded at 28°C in 3-s intervals between frames using a TIRF microsopy setup described previously (Lampert et al., 2010). MetaMorph (Molecular Devices) software was used to compile images into videos and create kymographs out of individual moving microtubules. Velocity of gliding microtubules was calculated based upon the slope of the kymographs.

The tracking of the proteins was performed automatically using the Definiens Software Suite. Prior to the analysis, the image data were processed performing a shading correction and smoothing. This was done by applying an 11 × 11 pixel (1463 × 1463 nm) kernel median filter and dividing the original raw image by the filtered image data. The resulting image was smoothed, using a 3 × 3 (399 × 399 nm) kernel Gaussian filter. The processed image data were searched and segmented for fluorescent signal. The signal area was searched for local intensity maxima within a search range of 532 nm. Circular objects with a radius of 332 nm were created on the found maxima and used for tracking the identified proteins.

The tracking of the movement of the TMR-, Alexa-488-, or GFP-labeled proteins was done through linking the proteins frame by frame by direct overlap, using the best fitting overlap.

Cik1 protein sequences of Saccharomyces cerevisiae (accession number NP_013925.1), Saccharomyces kudriavzevii (EJT42048.1), Saccharomyces arboricola (EJS44163.1) and Vik1 sequences from S. cerevisiae (NP_015070.1), S. kudriavzevii (EJT43871.1), S. arboricola (EJS41496.1) were retrieved from the National Center for Biotechnology Information (NCBI). The Saccharomyces bayanus protein sequences of Cik1 (WashU_Sbay_Contig651.30) and Vik1 (WashU_Sbay_Contig637.18) were retrieved from the Saccharomyces Genome Database (SGD). The sequences were aligned with MAFFT (version 6, L-INS-I method) and further processed with Jalview and colored according to ClustalW.

Varying amounts (0.1–1 μM) of recombinant bait protein (GST, GST-Bim1^FL, GST-Bim1^0–185, GST-Bim1^185–344) were immobilized on 30 μl glutathione sepharose (GE Healthcare) in 0.5 ml binding buffer (25 mM Hepes pH 7.2, 250 mM NaCl, 1 mM MgCl₂, 1 mM EGTA, 0.5 mM DTT, 0.05% NP-40). The binding partner was added at a constant concentration between 0.5 and 1 μM and incubation lasted for 1 hr at 4°C. Afterwards beads were washed three times with 0.5 ml binding buffer and analyzed by SDS-PAGE and Coomassie Brilliant blue R250 staining.

All modifications were performed in the S288C background (Supplementary file 2). Genetic modifications were introduced by using standard procedures.

For the spot assay, the desired strains were grown overnight in YPD medium. The following day cells were diluted to OD₆₀₀ = 0.6 which was the starting point of a 1:4 dilution series and spotted on YPD or 100 mM hydroxyurea (HU) plates. Plates were incubated at the indicated temperatures to 2–3 days.

Imaging strains were grown in synthetic medium containing 2% glucose and imaged on concanavalin A-coated culture dishes (Matek) at ambient temperature. Eight z stacks with planes 0.3 μm apart were acquired at 30 s intervals on an Axiovert 200M microscope (Carl Zeiss) using an UPlanSApo 100× NA 1.40 oil immersion objective lens (Olympus) and a Coolsnap HQ CCD camera (Photometrics). Images were projected to two-dimensional images (SoftWoRx software) and further analyzed by MetaMorph (Molecular Devices). The linescans showing the fluorescence intensity for Kar3-3xGFP on spindles were plotted using ImageJ.

Desired strains were grown to OD₆₀₀ = 1.2 in YPD, centrifuged, drop frozen in liquid nitrogen, and lysed by freezer mill treatment. 5 g of yeast powder was dissolved in 10 ml buffer A (25 mM Hepes pH 8.0, 2 mM MgCl₂, 0.5 mM EGTA pH 8.0, 0.1 mM EDTA, 0.1% NP-40, 15% glycerole, 150 mM KCl, 0.01 mM ATP, 0.1 mM PMSF, 1× protease inhibitor cocktail set IV [Calbiochem]). The lysed cells were centrifuged twice, first at 43.146×g for 20 min and afterwards at 125.749×g for 1 hr. The resulting supernatant was incubated for 2–3 hr with 100 μl Dynabeads (Life Technologies) that were coupled to 50 μl anti-FLAG M2 antibody (Sigma Aldrich). Beads were washed three times with buffer A and four times with buffer B (25 mM Hepes pH 8.0, 150 mM KCl). Elution was achieved by using 1 beads volume 2 mg/ml 3xFLAG peptide in buffer B. The elution fractions were analyzed by SDS-PAGE and silver stain. For mass spectrometry analysis, an on-bead digest replaced the elution procedure: 500 ng LysC protease was added per 50 μl dynabeads in ammonium bicarbonate buffer and incubated at 37°C overnight. The supernatant was filtered and applied to mass spectrometry analysis, which was performed on three independent preparations.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

The authors thank all members of the Westermann lab for discussions and Jan-Michael Peters and David Keays for critical reading of the manuscript. We thank Karin Aumayr, Gabriele Stengl, and Pawel Paserbiek for help with microscopy and image analysis and Martin Colombini for fabrication of custom mechanical components. We thank Marlene Brandstaetter and Alexander Schleiffer for excellent technical support. This work received funding from the European Research Council under the European Community's Seventh Framework Programme (SW FP7/2007-2013)/ERC grant agreement number [203499], from the Austrian Science Fund FWF (SW, SFB F34-B03) and the Austrian Research Promotion Agency (FFG). MIM acknowledges the VIPS Program of the Austrian Federal Ministry of Science and Research and the City of Vienna. The research leading to these results has received funding from the Vienna Science and Technology Fund (WWTF) project VRG10-11, Human Frontiers Science Program Project RGP0041/2012, Research Platform Quantum Phenomena and Nanoscale Biological Systems (QuNaBioS). The IMP is funded by Boehringer Ingelheim.

© 2015, Mieck et al.

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