Skip to main content
Proceedings of the National Academy of Sciences of the United States of America logoLink to Proceedings of the National Academy of Sciences of the United States of America
. 2010 Jun 21;107(27):12151–12156. doi: 10.1073/pnas.1004736107

Kinesin-1 heavy chain mediates microtubule sliding to drive changes in cell shape

Amber L Jolly 1, Hwajin Kim 1,1, Divya Srinivasan 1,2, Margot Lakonishok 1, Adam G Larson 1, Vladimir I Gelfand 1,3
PMCID: PMC2901438  PMID: 20566873

Abstract

Microtubules are typically observed to buckle and loop during interphase in cultured cells by an unknown mechanism. We show that lateral microtubule movement and looping is a result of microtubules sliding against one another in interphase Drosophila S2 cells. RNAi of the kinesin-1 heavy chain (KHC), but not dynein or the kinesin-1 light chain, eliminates these movements. KHC-dependent microtubule sliding powers the formation of cellular processes filled with parallel microtubule bundles. The growth of these cellular processes is independent of the actin cytoskeleton. We further observe cytoplasmic microtubule sliding in Xenopus and Ptk2 cells, and show that antibody inhibition of KHC in mammalian cells prevents sliding. We therefore propose that, in addition to its well established role in organelle transport, an important universal function of kinesin-1 is to mediate cytoplasmic microtubule–microtubule sliding. This provides the cell with a dedicated mechanism to transport long and short microtubule filaments and drive changes in cell shape.

Keywords: cytoskeleton, motor proteins, dynein, cell morphology, Drosophila


The microtubule network has traditionally been viewed as stationary tracks along which motor proteins such as conventional kinesin (kinesin-1) move cargo. Microtubules are well known to be mechanically stiff structures in vitro (1, 2), in line with their suggested role in providing structural integrity. However, microtubules are typically observed to buckle and loop in cultured cells (36). This motion is in apparent contrast to the need for microtubules in directing the precise delivery of cellular cargoes.

In the course of analyzing organelle movement along microtubules in cultured Drosophila S2 cells, we found that many microtubules in the cytoplasm undergo extensive buckling and looping (Fig. 1 A and C and Movie S1) (6). Our laboratory previously showed that microtubule buckling accounted for the observed cotransport of multiple peroxisomes in S2 cells. In this way, cargo can be transported not only along a stationary track, but by “piggybacking” along a moving microtubule (6). These findings support the idea that the microtubule network might be both pliable and portable.

Fig. 1.

Fig. 1.

Microtubule bending, looping, and sliding in cultured Drosophila S2 cells. (A) Differential interference contrast image and (B) initial frame of time-lapse taken of Drosophila S2 cell stably expressing mCherry tubulin under the metallothionein promoter induced for 48 h with 200 μM copper sulfate. (C) Time-lapse panels show a closer view of the microtubules delineated in box B. Two microtubules highlighted in green (arrows) display looping and bending. (D) Treatment of cells with 1 μM colcemid induces formation of short depolymerization-resistant microtubule fragments. (E) At 3.5 h after addition of colcemid these fragments (corresponding to boxed area in D) realign into bundles. (F) After 4 h, a majority of the cells have a few large bundles containing almost all fragments (Fig. S4). (Scale bars, 5 μm.)

There are several potential sources of force that might drive microtubule movement in interphase cells. Microtubules may be reacting to force generated from tubulin polymerization or depolymerization; additionally, the microtubules are subject to reactive forces generated by cargo moving along microtubules, and to indirect forces such as actin flow. Alternatively, the movement might be caused by a dedicated mechanism, and possibly be motor-driven, much like microtubule–microtubule sliding in the mitotic spindle. Microtubule sliding by the plus-end–directed kinesin motor Klp61F (7, 8), the minus-end–directed kinesin Ncd (9), and the minus-end–directed cytoplasmic dynein motor (9) are required for proper spindle separation during anaphase. However, Klp61F and Ncd activity is limited to mitosis (10, 11). Dynein activity is not limited to the mitotic phase (12, 13), making it a candidate for mediating both mitotic and interphase microtubule interactions. Microtubule bending and looping have been attributed to several forces from acto-myosin contractility (14, 15) to the activity of molecular motors (3, 5); these latter studies implicate the minus end directed motor dynein in the majority of the motility events based on the observation that the bending appears to be mostly in the anterograde direction.

Using a photoconvertible fluorescent tag fused to tubulin, we developed a method to quantify the lateral microtubule motion and normalize the motile fraction to the total microtubule population. This technique allowed us to identify conventional kinesin heavy chain (KHC) as the source of force powering the lateral microtubule motions, and this mechanism accounted for the observed buckling and looping. This finding builds upon previous work identifying an in vitro ability of KHC to slide microtubules (16), although kinesin-1 has no function in mitosis (17). In vitro evidence revealed the existence of an ATP-independent C-terminal microtubule-binding domain in the kinesin heavy chain (18, 19). Although overexpressed tail can bind to microtubules in cells (20), it was unclear whether full-length kinesin-1 uses this site and the in vivo relevance of the C-terminal microtubule binding site remained unknown. Our findings demonstrate a ubiquitous role for KHC-mediated microtubule sliding in transporting microtubules against one another during interphase, and in powering the formation of parallel microtubule bundles strong enough to deform cellular membranes.

Results

Lateral Microtubule Motion Is Independent of Microtubule Dynamics.

It is well established that microtubule dynamics can produce sufficient force to buckle a microtubule (21). To investigate whether the observed movement was caused by microtubule polymerization or depolymerization, we blocked microtubule dynamics by treating cells with paclitaxel. As long-term paclitaxel treatment can dramatically disrupt the microtubule network, we imaged cells between 5 and 30 min following the addition of 5 μM paclitaxel. This treatment did not cause microtubule reorganization, but was sufficient to block microtubule polymerization (Fig. S1). Strikingly, the lateral motion of microtubules was clearly observed when microtubule dynamics were suppressed, and was qualitatively similar to the motion observed in untreated cells (Movie S2).

Marking Microtubules with a Fiducial to Quantify Movement.

To elucidate the precise mechanism of microtubule movement, we developed a method to quantify the movement by applying a fiduciary mark to microtubules in Drosophila S2 cells. We created a fusion of Drosophila α-tubulin and an N-terminal fluorescent tag, photoconvertible protein Dendra2 (22), under the control of an inducible metallothionein promoter (pMT). Before photoconversion, the emission of Dendra2 has a characteristic peak at 505 nm, but following conversion with blue light the emission peak shifts to 575 nm. Using a line-scan confocal LSM510 microscope (Zeiss), and converting with a 405-nm diode laser, we limited the photoconversion to a small circular area approximately 5 μm in diameter between the cell nucleus and the periphery.

Performing the photoconversion in S2 cells allowed us to follow the movement of microtubule segments outside of the photoconverted area (Fig. S2). However, the fluorescence was rapidly lost from labeled segments as the photoconverted (red) tubulin was incorporated into newly forming microtubules elsewhere in the cytoplasm as a result of the microtubule dynamics. In paclitaxel-treated cells, the loss of fluorescence as a result of microtubule dynamics was prevented, allowing the observation of sliding over an extended period (Fig. 2 A and C and Movie S3). Our photoconversion experiments revealed that, rather than being the transport of small microtubule fragments along long filaments, entire long microtubule filaments undulate and buckle.

Fig. 2.

Fig. 2.

Microtubule sliding visualized and quantified using photoconversion. (A) S2 cell stably expressing pMT-Dendra2-α-tubulin before photoconversion shown in the green channel. The same cell in the red channels immediately after conversion (B) and 6 min 29 s later (C) demonstrates movement of the photoconverted segments beyond the converted area. (DF) Cell treated with 5 μM cyto D 1 h before imaging and imaged 1 h after plating. Other conditions are identical to AC, respectively. (G) Distribution of microtubule segment sliding velocities of 23 tracked segments from eight cells. Each point is equal to one velocity vector (i.e., the displacement between two consecutive frames). (H) Quantification of motile fraction of converted segments in 20 frames (15 s intervals) with or without cyto D treatment. Error bars indicate SEM. All cells were treated with 5 μM paclitaxel. (Scale bars, 5 μm.)

We determined the degree of microtubule movement relative to the total number of converted microtubule segments by calculating the motile fraction of the converted regions at the end of each 7-min time-lapse sequence (Materials and Methods). Photoconverting a circular region eliminated any angle bias in the analysis of microtubule movement. The mean motile fraction of converted microtubule segments was 0.36 (n = 10 cells), indicating that, on average, 36% of the total amount of fluorescent microtubule segments moved outside of the converted region. Tracking of the leading end of 23 microtubule segments from eight cells revealed that many segments spent most of the time not moving, but underwent sudden long-distance travel and were capable of moving during these bursts as fast as 13 μm/min (Fig. 2G). Other segments moved more constantly. The mean peak velocity of the 23 trajectories analyzed was 4.2 ± 0.5 μm/min (SEM).

Microtubule Movement Is Independent of Other Filament Networks.

Retrograde actin flow could be a potential source of pushing force driving microtubule movement in the cytoplasm. However, treatment of S2 cells with the actin-depolymerizing drug cytochalasin D (cyto D) did not inhibit microtubule motility (Fig. 2 DF). Untreated S2 cells and those treated with cyto D had the same mean motile fraction (Fig. 2H). S2 cells lack an intermediate filament network (23), and when treated with cyto D, the microtubule network is the only remaining intact cytoskeletal network. Therefore, our results show that the microtubules are not moving against any other cytoskeletal filaments.

Microtubule Movement Is Not a Reaction to Cargo Transport.

We used the RNAi method to identify motor proteins that might directly move microtubules or indirectly cause reactionary microtubule buckling by moving cargo. Successful protein knockdown was confirmed by Western blot (Fig. 3A). Cytoplasmic dynein is involved in the microtubule-based transport of the majority of cargoes studied, from nuclear envelope components (24) to RNA complexes (25). Kinesin-1 is also involved in a significant amount of microtubule based transport, and binds the majority of its cargoes through the light chain. If the lateral microtubule movement we observe were a reactionary force to the movement of cargo, RNAi of either dynein or the kinesin light chain (KLC) would inhibit the movement. In fact, knockdown of KLC had no effect on the amount of microtubule motility observed (Fig. 3A and Movie S4), although this treatment inhibited the movement of membrane organelles along microtubules (26). Similarly, Klp68D (a Drosophila kinesin II subunit), another motor involved in cargo transport, did not influence microtubule motility (Fig. 3A). RNAi of the dynein heavy chain led to a small but statistically significant (P = 0.0406) increase in the motile fraction (Fig. 3A and Movie S5). As it is well established that dynein knockdown stops bidirectional cargo transport (13, 25, 26), these results show that cargo transport contributes very little, if at all, to the movement of microtubules in the cytoplasm.

Fig. 3.

Fig. 3.

RNAi of conventional kinesin, but not other motors, results in the cessation of microtubule–microtubule sliding and prevents the formation of microtubule bundle–filled processes. (A) Motile fraction of converted microtubule segments in S2 cells subjected to RNAi-mediated depletion of motors and associated proteins. Time-lapse sequences taken every 15 s for 20 frames. *P < 0.05, **P < 0.00001. Student two-tailed t test was performed for independent samples assuming variance differs for each sample. Error bars indicate SEM. (Inset) representative Western blots. Dilutions of untreated (mock) cells provided to estimate degree of knockdown. (B) Replacement of endogenous KHC with exogenous KHC results in rescue of sliding. S2 cells stably expressing pMT-Dendra2-α-tubulin/pMT-KHC (full length) were induced for 1 wk and subjected to RNAi of endogenous KHC by targeting the UTR of the mRNA (3′UTR KHC) or RNAi of both endogenous and exogenous KHC by targeting the coding region (CDS KHC) for 96 h before microscopy. ‘WT’ cells are equivalent to mock cells in A. The motile fraction was determined after photoconversion. (Inset, Right) representative Western blot showing extent of KHC knockdown. Exogenous (pMT-KHC) kinesin is present after treatment with dsRNA against the 3′UTR. (C and D) Microtubule networks visualized in mCherry tubulin expressing S2 cells plated in 5 μM cyto D for 1 h. Control cells (C) form long processes containing microtubule bundles whereas KHC knockdown cells form irregularly shaped lamellas filled with straight and less bundled microtubules (D). (Scale bars, 10 μm.)

Together, these findings suggest that the movement we observe is explained by microtubules sliding against one another. To test this hypothesis, we treated stable mCherry tubulin–expressing S2 cells with the microtubule-depolymerizing drug colcemid to create short, linear microtubule fragments. We consistently observed the movement of apparently overlapping microtubule fragments slide against, rotate, and realign parallel to one another (Fig. 1 D and E and Movie S6). Long-term treatment of these cells with colcemid led to the formation of a few microtubule bundles including the entire set of original microtubule fragments (Fig. 1F). We therefore conclude that the microtubule movement we observe is the result of microtubule–microtubule sliding.

RNAi of Conventional KHC Eliminates Microtubule–Microtubule Sliding.

Recent studies suggest a role for molecular motors in driving microtubule buckling, and implicate the minus-end directed motor dynein in the majority of microtubule bending events (3, 5). Our results indicate dynein does not drive microtubule sliding, and suggest another motor may be responsible. In particular, the in vitro ability for kinesin-1 to slide microtubules against one another (16) and the existence of an ATP-independent C-terminal microtubule-binding domain in the kinesin heavy chain (18, 19) indicate that conventional kinesin may play a role in interphase microtubule sliding.

Accordingly, we knocked down the kinesin-1 heavy chain, which resulted in a dramatic inhibition of microtubule–microtubule sliding (Fig. 3A and Movie S7). Specificity of knockdown was verified using two independent dsRNA sequences against the 3′UTR region of KHC and the N-terminal coding region. In both cases, the motile fraction was dramatically reduced (P = 1.3 × 10−6) from 0.36 ± 0.03 in WT cells to 0.03 ± 0.02 in the 3′UTR KHC RNAi cells and 0.04 ± 0.02 in the KHC coding region RNAi cells (Fig. 3A). To verify specific knockdown, exogenous full-length KHC was stably expressed before RNAi of endogenous kinesin-1 using dsRNA against the 3′UTR region. The sliding phenotype was rescued to near-WT levels (0.29 ± 0.04) in these cells (Fig. 3B).

KHC Drives Process Formation in S2 Cells Treated with Cyto D.

We previously demonstrated that S2 cells plated on Con A–coated substrate in the presence of cyto D form long processes (27) (Fig. S3). Microtubules in the processes form parallel bundles with the plus ends at the process tips (27) (Fig. 3C). After 9 h, these processes are between 10 and 60 μm in length (n = 40). Time-lapse fluorescent imaging of S2 cells expressing mCherry tubulin plated in cyto D–containing media reveal the rapid formation of these processes, which grow at a maximal rate of 3.7 ± 0.6 μm/min (SD) just as the cells adhere to the surface (n = 10). Microtubule looping and entry into the growing processes in the boundary region apparently contributes to the growth of these processes (Movie S8).

Knockdown of KHC in S2 cells prevents the formation of these microtubule bundle–filled processes (Fig. S3). Microtubules do not form bundles or processes in KHC knockdown cells treated with cyto D. Instead, the cells remain spread; but, unlike an untreated WT cell, the microtubule network appears straight and absent of lateral motion (Fig. 3D). Furthermore, KHC RNAi prevents microtubule bundle formation in cells treated with colcemid (Fig. S4). Instead, the short microtubule fragments concentrate around the nucleus in an unorganized fashion. Kinesin-1 is known to be able to cross-link microtubules in vitro (16, 18) and localizes to microtubule bundles in cells (3). Together, these results indicate a role for KHC in sliding microtubule filaments against one another to drive the formation of the microtubule bundle–filled processes we observe in cyto D–treated cells. Moreover, the amount of force generated by kinesin-1 driven microtubule sliding is sufficient to deform the cell membrane and drive the formation of processes, suggesting that microtubule sliding provides more force than polymerization.

KHC-Driven Microtubule Sliding in Amphibian and Mammalian Cells.

Drosophila S2 cells have unique cytoskeletal networks. In addition to lacking intermediate filaments, they also lack interphase microtubule organizing centers (28). We wondered if the microtubule–microtubule sliding phenomenon we observed in S2 cells was a consequence of having free microtubules unhindered by “normal” cytoskeleton organizing structures and centrosome attachment of the minus ends. We demonstrate the occurrence of sliding in Xenopus fibroblasts by transiently transfecting with Dendra2-Xenopus α-tubulin and following the movement of photoconverted fragments outside of the converted region (Fig. 4 A and B and Movie S9). Additionally, time-lapse microscopy of GFP-tubulin–expressing rat kangaroo epithelial (Ptk2) cells treated with paclitaxel to inhibit microtubule dynamics reveals dramatic microtubule sliding and looping (Movie S10). Ptk2 cells were microinjected with a function-blocking SUK-4 antibody against the kinesin-1 heavy chain (29), along with a dextran tracer. SUK-4 injection caused all microtubule motion to cease, even in the absence of paclitaxel treatment (Movie S11). Six percent of injected cells displayed microtubule movement compared with uninjected or cells injected with control IgG (anti-myc antibody; Fig. 4C and Movie S12). We conclude that kinesin-1 drives sliding in mammalian cells in addition to Drosophila.

Fig. 4.

Fig. 4.

Microtubule sliding occurs in a variety of cell types, and is driven by KHC in Ptk2 cells. (A) Xenopus fibroblast transiently expressing Dendra2-α-tubulin immediately after photoconversion in a narrow bar; (B) the same cell approximately 5 min later. (Scale bars, 20 μm.) Arrows point to fluorescent microtubule segments that moved away from the converted region. (C) Percent of PtK2 cells injected with SUK-4 or a control antibody (mouse anti-myc) possessing moving microtubules as compared with uninjected cells (motility determined in a double-blind survey). (D) Model of kinesin-1–driven microtubule-microtubule sliding. (E) Alignment showing conservation of C-terminal microtubule binding domain (boxed region) identified in Drosophila KHC (19).

Discussion

There are several potential mechanisms by which kinesin might mediate microtubule sliding in interphase cells. Although it is theoretically possible that, like Eg5, kinesin-1 might form an oligomer such that the head domains walk along two antiparallel microtubules, sedimentation experiments show kinesin-1 is unable to form oligomers (30). Instead, we favor a model in which a single kinesin-1 molecule cross-links two microtubules through head-microtubule and tail-microtubule interactions using the highly conserved C-terminal microtubule binding region (Fig. 4 D and E). In this model, KHC more closely mimics the mechanism used by the mitotic Ncd motor. We attempted to test our model in S2 cells by creating kinesin mutants with truncated or scrambled C-terminal microtubule binding domains that we hypothesized would be unable to slide microtubules but still move cargo (and bind KLC). However, we could not express these mutants in S2 cells, presumably because they are degraded.

Bundling of cytoplasmic microtubules by conventional kinesin has been previously demonstrated by Straube et al. in the fungus Ustilago maydis (3), indicating that kinesin-1 can form cross-bridges between microtubules in living cells. However, the authors state that only 4% of all bending motility seemed to be supported by a plus end–directed kinesin, and in dynein mutants of U. maydis, a reduction in microtubule motility was observed (3). In our study, we find that kinesin-1 is the single major driving force responsible for microtubule sliding and bundle formation in several types of animal cells. The discrepancy may be a result of the technique used. Straube and colleagues concluded based on the directionality of the bending that a minus end–directed motor is responsible for most of the sliding events. However, microtubule orientation does not necessarily reveal the identity of the motor involved because it is equally probable for the microtubule interacting with the motor head to be translocated as it is for the microtubule interacting with the motor tail. In this study, we observe a modest but reproducible increase in sliding upon disruption of the dynein motor. Dynein therefore appears to inhibit the ability of kinesin-1 to slide microtubules, possibly by increasing the likelihood that kinesin-1 is participating in organelle transport.

The kinesin-1 motor is a kinesin that is able to transport cargo along microtubule tracks and transport the tracks themselves. We show that these jobs are distinct by knockdown of the cargo-binding light chain, which affects neither sliding nor the formation of cellular processes. In our model, the light chain is not involved in sliding, although it is known to be in a complex with the heavy chain. Interphase microtubule sliding also further differs from cargo transport in terms of speed. The approximately 10-fold slower rate of microtubule sliding compared with organelle transport can be attributed to the higher load imposed by a filament, as kinesin velocity decreases under increased load (31).

Microtubule–microtubule sliding is a fundamental process that drives anaphase during mitosis, and powers flagellar movement—in these examples, microtubules themselves transmit a substantial force. Our results suggest that the sliding of microtubules is even more ubiquitous, frequently occurring in interphase cells, and is explained by a dedicated mechanism. Furthermore, as the microtubule-binding domain at the C terminus of kinesin-1 is evolutionarily conserved, microtubule–microtubule sliding is likely used in many organisms and in different cell types. We propose that conventional kinesin-mediated microtubule sliding could be important in at least two biological processes. First, as demonstrated in S2 cells, kinesin-1–driven sliding of cytoplasmic microtubules might drive the biogenesis of similar processes in other cell types, for example neurite outgrowth in neuronal cells. The long-distance transport of short microtubule fragments in neurons has been suggested to contribute to axonal growth, as protein expression is limited in the far regions of the axon (for review, see ref. 32). Kinesin-1–mediated microtubule sliding might drive the transport of short microtubules toward the tips of these growing processes.

Secondly, kinesin-1–mediated microtubule sliding explains how the cell can control the simultaneous transport of multiple cargoes piggybacking along a moving microtubule (6). This second function may explain the irregular occurrence of microtubule sliding we observe in the dense microtubule lattice of an interphase animal cell. These two types of movement (sliding of microtubules and process formation) are likely driven by the same mechanism, but the generation of the cellular process has the additional requirement that a moving microtubule be positioned against the tip of the process.

This mechanism may explain other microtubule-based phenomena, such as early Drosophila development. Ooplasmic streaming during Drosophila oogenesis is required for the correct distribution of mRNA and protein, and has been shown to be dependent on microtubules and the KHC, but independent of the KLC (33). Our findings suggest a mechanism for potentially numerous microtubule-driven processes and highlight the importance of kinesin-1–mediated microtubule sliding in cell deformation and force generation.

Materials and Methods

Cloning.

Drosophila constructs pMT-mCherry-tubulin, pMT-Dendra-tubulin, and pMT-KHC were cloned under the control of the pMT using the pMT/V5-His A vector (Invitrogen). Dendra2 (Evrogen) or mCherry were introduced between KpnI and EcoRI, N-terminal of Drosophila α-tubulin (inserted between NotI and XbaI). Cloning of pMT-KHC (full-length) is described elsewhere (13).The pDendra2-human-α tubulin construct was made by replacing EGFP in pEGFP-Tub (Clontech) with Dendra-2 between NheI and BglII.

Cell Culture and Transfection.

Drosophila S2 cells were maintained in Schneider media supplemented with 10% FBS or in serum-free Insect-Xpress media (BioWhittaker). Xenopus cells were maintained in Leibovitz L-15 media supplemented with 10% FBS and were transfected with Lipofectamine 2000 (Invitrogen) in Opti-MEM media (Sigma) according to the manufacturer's protocol. Ptk2 cells were maintained in DMEM supplemented with 10% FBS.

Stable Cell Lines.

To create stable lines, S2 cells were transfected using Cellfectin (Invitrogen) with a 20:1 molar ratio of the plasmid(s) of interest and the selection plasmid pCoHygro (Invitrogen) according to the Drosophila Expression System protocol (Invitrogen). Transfection was verified by fluorescence microscopy, and 48 h after transfection, 300 μg/mL hygromycin was added for selection. Cells remained in selection for 4 to 5 wk. The stable Ptk2 GFP-tubulin line was a gift from Alexey Khodjakov (Wadsworth Center, Albany, NY) (34).

Antibodies for Immunoblotting.

Antibodies used were as follows: anti-DHC monoclonal antibody (35) provided by J. Scholey (University of California, Davis, CA); HD antibody against KHC head domain and pan-KLC antibody provided by A. Minin (Institute of Protein Research, Russian Academy of Sciences, Moscow, Russia); SUK-4 antibody against KHC obtained from the Developmental Studies Hybridoma Bank; and monoclonal actin antibody CLT9001 (Accurate Chemicals).

Microscopy.

Cells expressing the pMT-mCherry-tubulin construct were induced overnight with 200 μM copper sulfate before microscopy. Drosophila cells were plated on Concanavalin-A–treated coverslips for 1 h before microscopy. For photoconversions, confocal microscopy was performed on a Zeiss UV LSM510 META laser scanning microscope using a 100×, 1.46-NA Plan-Apochromat objective. S2 cells stably expressing pMT-Dendra-α-tubulin were induced with 200 μM copper sulfate for 1 wk before microscopy. Cells were treated with 5 μM paclitaxel and microscopy was performed between 5 and 30 min after paclitaxel addition. Bright cells were selected for photoconversions, and a green image of the entire microtubule network captured before conversion. Photoconversions were performed using a 405-nm diode laser, and converting in a circular region acquired with the bleaching module. Time-lapse images were taken in red every 23 s for 20 frames, with the first frame taken before conversion. A Nikon U-2000 inverted microscope equipped with a Perfect Focus system (Nikon) and Cascade II EMCCD (Roper Scientific) driven by Metamorph software was used for wide-field microscopy. A 100-W halogen light source was used for fluorescence excitation to minimize photobleaching and phototoxicity. Wide-field photoconversion of Dendra was performed using a 100-W mercury lamp and a DAPI filter, exposing for 8 s through a 75-μm slit inserted in the field diaphragm of the epiilluminator position and projected to a segment of approximately 7 μm in a specimen plane.

Calculation of Motile Fraction.

Confocal time-lapse movies of converted microtubules taken every 15 s for 20 frames (all movies were taken for a total of approximately 7.5 min as a result of acquisition lag time) were subjected to the following analysis to determine the motile fraction (MF), i.e., the (background corrected) fraction of fluorescence after conversion that moved outside of the initial converted region. All fluorescence measurements (integrated density) were limited to the final frame to eliminate the effects of photobleaching and/or variations in cell brightness.

graphic file with name pnas.1004736107eq1.jpg

where MF is the motile fraction, ai is initial converted object area, Ai is cell area in green before conversion, bf is background integrated density in final frame (i.e., in region “a” translocated to an unconverted region), if is integrated density in region “a” in final frame, and If is integrated density in region “A” in final frame.

Tracking.

Metamorph software was used to track the leading edge of converted microtubule segments or growing processes. Peak velocities are defined as the maximal velocities during a given object's trajectory.

Microinjections.

Cells were microinjected with protein-A purified monoclonal antibodies (SUK-4 against KHC or 9E11 anti-myc antibody used as a control). The final concentration of antibodies in the needle in microinjection buffer was 3.6 mg/mL for SUK-4 or 2.3 mg/mL for Myc. Injection buffer also contained 0.5 mg/mL Texas Red dextran. Cells were allowed to recover for at least 2 h after microinjection before microscopy.

Double-Stranded RNAi.

S2 cells were plated at a density of 5.4 × 105/mL in 1.5 mL in a 12-well plate with 20 μg dsRNA, and collected after 96 h. Efficiency of knockdown was determined by Western blotting for each experiment. Double-stranded RNA was transcribed in vitro with T7 polymerase, and purified using LiCl extraction (13). Primers used in PCR reactions to create T7 templates from cDNA were as follows (T7 promoter sequences (TAATACGACTCACTATAGGG) were added to the 5′ end of each primer): KHC (N-terminal CDS), forward, ATGTCCTCACACCAGAAGAAGC; reverse, GGTGAGGATGATGTTCTGAAGC; KHC (3′UTR), forward, ATCCAATCACCACCTGTCGC; reverse, TCTGCGACTTTTATTTAGGT; DHC, forward, AAACTCAACAGAATTAACGCCC; reverse, TTGGTACTTGTCACACCACT; Klp68D, forward, CATGATCAAAATCGAGATGTGC; reverse, AAGTTGACCCTCCAATTCTGC; and KLC, forward, GCATGTTTCGATTATGAACGGA; reverse, GTTGTGTCTGTCCTCGTTTTC.

Supplementary Material

Supporting Information

Acknowledgments

We thank A. Mikhailov for help with a spinning disk microscope, M. Mendez (Northwestern University) for the actin antibody, G. Albrecht-Buehler for experimental discussions, and A. S. Belmont and K. R. Yamamoto for reviewing the manuscript. This work was supported by the National Institutes of Health (NIH) Grant GM052111 (to V.I.G.). A.L.J. holds a fellowship from the National Science Foundation. A.G.L. was funded by NIH Grant GM072656 to S. E. Rice.

Footnotes

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1004736107/-/DCSupplemental.

References

  • 1.Gittes F, Mickey B, Nettleton J, Howard J. Flexural rigidity of microtubules and actin filaments measured from thermal fluctuations in shape. J Cell Biol. 1993;120:923–934. doi: 10.1083/jcb.120.4.923. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.VanBuren V, Cassimeris L, Odde DJ. Mechanochemical model of microtubule structure and self-assembly kinetics. Biophys J. 2005;89:2911–2926. doi: 10.1529/biophysj.105.060913. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Straube A, Hause G, Fink G, Steinberg G. Conventional kinesin mediates microtubule-microtubule interactions in vivo. Mol Biol Cell. 2006;17:907–916. doi: 10.1091/mbc.E05-06-0542. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Steinberg G, Wedlich-Söldner R, Brill M, Schulz I. Microtubules in the fungal pathogen Ustilago maydis are highly dynamic and determine cell polarity. J Cell Sci. 2001;114:609–622. doi: 10.1242/jcs.114.3.609. [DOI] [PubMed] [Google Scholar]
  • 5.Bicek AD, et al. Anterograde microtubule transport drives microtubule bending in LLC-PK1 epithelial cells. Mol Biol Cell. 2009;20:2943–2953. doi: 10.1091/mbc.E08-09-0909. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Kulic IM, et al. The role of microtubule movement in bidirectional organelle transport. Proc Natl Acad Sci USA. 2008;105:10011–10016. doi: 10.1073/pnas.0800031105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Scholey JM. Kinesin-5 in Drosophila embryo mitosis: Sliding filament or spindle matrix mechanism? Cell Motil Cytoskeleton. 2009;66:500–508. doi: 10.1002/cm.20349. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Sharp DJ, Yu KR, Sisson JC, Sullivan W, Scholey JM. Antagonistic microtubule-sliding motors position mitotic centrosomes in Drosophila early embryos. Nat Cell Biol. 1999;1:51–54. doi: 10.1038/9025. [DOI] [PubMed] [Google Scholar]
  • 9.Goshima G, Nédélec F, Vale RD. Mechanisms for focusing mitotic spindle poles by minus end-directed motor proteins. J Cell Biol. 2005;171:229–240. doi: 10.1083/jcb.200505107. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Heck MM, et al. The kinesin-like protein KLP61F is essential for mitosis in Drosophila. J Cell Biol. 1993;123:665–679. doi: 10.1083/jcb.123.3.665. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Goshima G, Vale RD. Cell cycle-dependent dynamics and regulation of mitotic kinesins in Drosophila S2 cells. Mol Biol Cell. 2005;16:3896–3907. doi: 10.1091/mbc.E05-02-0118. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Allan VJ, Vale RD. Cell cycle control of microtubule-based membrane transport and tubule formation in vitro. J Cell Biol. 1991;113:347–359. doi: 10.1083/jcb.113.2.347. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Ally S, Larson AG, Barlan K, Rice SE, Gelfand VI. Opposite-polarity motors activate one another to trigger cargo transport in live cells. J Cell Biol. 2009;187:1071–1082. doi: 10.1083/jcb.200908075. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Waterman-Storer CM, Salmon ED. Actomyosin-based retrograde flow of microtubules in the lamella of migrating epithelial cells influences microtubule dynamic instability and turnover and is associated with microtubule breakage and treadmilling. J Cell Biol. 1997;139:417–434. doi: 10.1083/jcb.139.2.417. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Gupton SL, Salmon WC, Waterman-Storer CM. Converging populations of f-actin promote breakage of associated microtubules to spatially regulate microtubule turnover in migrating cells. Curr Biol. 2002;12:1891–1899. doi: 10.1016/s0960-9822(02)01276-9. [DOI] [PubMed] [Google Scholar]
  • 16.Urrutia R, McNiven MA, Albanesi JP, Murphy DB, Kachar B. Purified kinesin promotes vesicle motility and induces active sliding between microtubules in vitro. Proc Natl Acad Sci USA. 1991;88:6701–6705. doi: 10.1073/pnas.88.15.6701. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Goshima G, Vale RD. The roles of microtubule-based motor proteins in mitosis: Comprehensive RNAi analysis in the Drosophila S2 cell line. J Cell Biol. 2003;162:1003–1016. doi: 10.1083/jcb.200303022. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Andrews SB, Gallant PE, Leapman RD, Schnapp BJ, Reese TS. Single kinesin molecules crossbridge microtubules in vitro. Proc Natl Acad Sci USA. 1993;90:6503–6507. doi: 10.1073/pnas.90.14.6503. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Hackney DD, Stock MF. Kinesin's IAK tail domain inhibits initial microtubule-stimulated ADP release. Nat Cell Biol. 2000;2:257–260. doi: 10.1038/35010525. [DOI] [PubMed] [Google Scholar]
  • 20.Navone F, et al. Cloning and expression of a human kinesin heavy chain gene: Interaction of the COOH-terminal domain with cytoplasmic microtubules in transfected CV-1 cells. J Cell Biol. 1992;117:1263–1275. doi: 10.1083/jcb.117.6.1263. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Dogterom M, Yurke B. Measurement of the force-velocity relation for growing microtubules. Science. 1997;278:856–860. doi: 10.1126/science.278.5339.856. [DOI] [PubMed] [Google Scholar]
  • 22.Gurskaya NG, et al. Engineering of a monomeric green-to-red photoactivatable fluorescent protein induced by blue light. Nat Biotechnol. 2006;24:461–465. doi: 10.1038/nbt1191. [DOI] [PubMed] [Google Scholar]
  • 23.Goldstein LS, Gunawardena S. Flying through the drosophila cytoskeletal genome. J Cell Biol. 2000;150:F63–F68. doi: 10.1083/jcb.150.2.f63. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Salina D, et al. Cytoplasmic dynein as a facilitator of nuclear envelope breakdown. Cell. 2002;108:97–107. doi: 10.1016/s0092-8674(01)00628-6. [DOI] [PubMed] [Google Scholar]
  • 25.Ling SC, Fahrner PS, Greenough WT, Gelfand VI. Transport of Drosophila fragile X mental retardation protein-containing ribonucleoprotein granules by kinesin-1 and cytoplasmic dynein. Proc Natl Acad Sci USA. 2004;101:17428–17433. doi: 10.1073/pnas.0408114101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Kim H, et al. Microtubule binding by dynactin is required for microtubule organization but not cargo transport. J Cell Biol. 2007;176:641–651. doi: 10.1083/jcb.200608128. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Kural C, et al. Kinesin and dynein move a peroxisome in vivo: A tug-of-war or coordinated movement? Science. 2005;308:1469–1472. doi: 10.1126/science.1108408. [DOI] [PubMed] [Google Scholar]
  • 28.Rogers GC, Rusan MN, Peifer M, Rogers SL. A multicomponent assembly pathway contributes to the formation of acentosomal microtubule arrays in interphase Drosophila cells. Mol Biol Cell. 2008;19:3163–3178. doi: 10.1091/mbc.E07-10-1069. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Ingold AL, Cohn SA, Scholey JM. Inhibition of kinesin-driven microtubule motility by monoclonal antibodies to kinesin heavy chains. J Cell Biol. 1988;107:2657–2667. doi: 10.1083/jcb.107.6.2657. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Kuznetsov SA, et al. The quaternary structure of bovine brain kinesin. EMBO J. 1988;7:353–356. doi: 10.1002/j.1460-2075.1988.tb02820.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Visscher K, Schnitzer MJ, Block SM. Single kinesin molecules studied with a molecular force clamp. Nature. 1999;400:184–189. doi: 10.1038/22146. [DOI] [PubMed] [Google Scholar]
  • 32.Baas PW, Vidya Nadar C, Myers KA. Axonal transport of microtubules: The long and short of it. Traffic. 2006;7:490–498. doi: 10.1111/j.1600-0854.2006.00392.x. [DOI] [PubMed] [Google Scholar]
  • 33.Palacios IM, St Johnston D. Kinesin light chain-independent function of the Kinesin heavy chain in cytoplasmic streaming and posterior localisation in the Drosophila oocyte. Development. 2002;129:5473–5485. doi: 10.1242/dev.00119. [DOI] [PubMed] [Google Scholar]
  • 34.Khodjakov A, Copenagle L, Gordon MB, Compton DA, Kapoor TM. Minus-end capture of preformed kinetochore fibers contributes to spindle morphogenesis. J Cell Biol. 2003;160:671–683. doi: 10.1083/jcb.200208143. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Sharp DJ, et al. Functional coordination of three mitotic motors in Drosophila embryos. Mol Biol Cell. 2000;11:241–253. doi: 10.1091/mbc.11.1.241. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supporting Information
Download video file (811.4KB, avi)
Download video file (1.4MB, avi)
Download video file (537.5KB, avi)
Download video file (419.2KB, avi)
Download video file (549.2KB, avi)
Download video file (1.6MB, avi)
Download video file (316.5KB, avi)
Download video file (1.6MB, avi)
Download video file (1.7MB, avi)
Download video file (1.3MB, avi)
Download video file (659KB, avi)
Download video file (408.5KB, avi)

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences

RESOURCES