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2 Department of Cell Biology, Harvard Medical School, Boston, MA 02115
Address correspondence to Tarun Kapoor, Laboratory of Chemistry and Cell Biology, Rockefeller University, 1230 York Ave., Box 202, New York, NY 10021. Tel.: (212) 327-8176. Fax: (212) 327-8177. E-mail: kapoor{at}mail.rockefeller.edu
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Abstract |
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Key Words: mitosis; Eg5; tubulin; speckle; kinesin
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Introduction |
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A nonmicrotubule "spindle matrix" was first hypothesized to provide an anchor for motor proteins that exert force on microtubules (for reviews see McIntosh et al., 1969; Pickett-Heaps et al., 1982, 1984). Support for the existence of a matrix came from several lines of experiment: extraction of microtubules from isolated spindles produces a "spindle remnant" that retains a spindle shape and contains kinesin and other nonmicrotubule spindle components (Mazia, 1961; Salmon and Segall, 1980; Leslie et al., 1987). Observations of polewards movement of chromosomes with severed kinetochore microtubule bundles (Forer et al., 1997) and the behavior of spindles in which motor proteins were disrupted (Gordon et al., 2001) can be interpreted as providing evidence for a matrix. Recently, the Drosophila protein Skeletor was suggested to be a component of a spindle matrix on the basis of its immunolocalization (Walker et al., 2000). This body of evidence is intriguing rather than compelling, and a matrix that permeates the whole spindle has not been directly observed or biochemically characterized. This is in contrast to the spindle pole, where biochemical and imaging evidence supports the idea that NuMA serves as a matrix component (Dionne et al., 1999; Merdes et al., 2000; Gordon et al., 2001) and the centrosome, where pericentrin and related proteins may serve a matrix function (Schnackenberg et al., 1998). Most recent authors have tried to account for spindle dynamics and mechanics in terms of models based purely on microtubules and motor proteins (Sharp et al., 2000). If the matrix hypothesis were correct, such models would require revision.
The mitotic kinesin Eg5 is a conserved spindle component with a key role in establishing bipolar organization of the spindle (Enos and Morris, 1990; Hagan and Yanagida, 1990; Hoyt et al., 1992; Sawin et al., 1992; Heck et al., 1993; Blangy et al., 1995). In vertebrate somatic cells and Xenopus extract spindles, Eg5 is present throughout the spindle, but enriched at the poles relative to microtubules (Sawin and Mitchison, 1995; Kapoor et al., 2000). This unexpected localization for a plus enddirected motor lead to the proposal that Eg5 might target to spindles in part by interacting with some unknown matrix component (Sawin et al., 1992). However, the observation that native Eg5 is a bipolar tetramer lead to an alternative proposal, that Eg5 targets to spindles by binding to two microtubules and cross bridging them, without interacting with other components (Sharp et al., 1999). We know that Eg5 must be phosphorylated on a cdc2 consensus site to target to spindles (Blangy et al., 1995; Sawin and Mitchison, 1995), and two-hybrid experiments suggested that it interacts with the dynactin complex (Blangy et al., 1997). However, the mechanism by which Eg5 targets to spindles, and exactly how it promotes bipolarity, are unresolved.
To probe Eg5 dynamics in Xenopus extract spindles, and thus gain insight into its targeting mechanism, we sought an imaging method that would provide some sense of its turnover behavior, but more important, would allow us to measure possible translocation of Eg5 along the spindle axis. Recently, fluorescent speckle microscopy has been described as a simple, nonperturbing method for obtaining high-resolution views of microtubule translocation in spindles (Waterman-Storer et al., 1998; Maddox et al., 2000). The speckled image results from stochastic variation in the number of fluorophores within minimal regions resolvable by light microscopy. Speckles serve as fiduciary marks, allowing measurement of microtubule movement and turnover (Waterman-Storer and Salmon, 1998). Fluorescent speckle microscopy provides a simple and reliable method for detecting slow, directed movements of immobilized fluorochromes, even in the presence of competing turnover or diffusion. It is more sensitive than either photobleaching or photoactivation for detecting movement in the face of turnover, and thus was our method of choice for parallel analysis of Eg5 and tubulin dynamics. Using this method, we found that although Eg5 can exchange between spindles, within the spindle it is relatively static, whereas microtubules flux poleward. We interpret our observations as revealing the existence of a static, nonmicrotubule mechanical scaffold that influences Eg5 dynamics and may play a central role in spindle organization.
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Results |
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We next used fluorescent speckle microscopy to examine the dynamics of Eg5 in spindles. The concentration of endogenous Eg5 in Xenopus egg extracts measured by Western blotting with expressed Eg5 as standard was 400 nM (not shown, calculated for monomer). Addition of 100150 nM of the labeled protein gave images with good speckle contrast (Fig. 2 c). Kymographs revealed streaks of Eg5 speckles normal to the spindle axis, indicating static populations of Eg5 (Fig. 2 d). These streaks of stationary Eg5 were observed throughout the spindle, and were typically 530 s in duration.
Comparison of the Eg5 and tubulin speckles in spindles revealed two major differences in their dynamic behavior (Fig. 2 e). First, most Eg5 speckles were static relative to the spindle poles. 70% of the speckles analyzed moved polewards at rates less the 0.25 µm/min. Most of the microtubules in the spindle flux polewards at rates >1.1 µm/min. Thus, populations of Eg5 are static while the microtubules move polewards. Second, the persistence of Eg5 speckles in spindles is less than that observed for tubulin speckles, as reflected by the lengths of the streaks in the kymographs, suggesting a faster turnover of Eg5 than tubulin.
Eg5's ATPase cycle influences its dynamic behavior in spindles
We next examined whether the dynamic behavior of Eg5 in the spindle depended on its ATPase activity. Adenylimidodiphosphate (AMPPNP),* a nonhydrolyzable ATP analogue, inhibits kinesins in a microtubule-bound state. 1.5 mM AMPPNP was added to spindles equilibrated with labeled Eg5 and immediately prepared for fluorescent speckle microscopy (Fig. 3
a). This concentration of AMPPNP is known to promote rigor binding of Eg5 to microtubules (unpublished data) and to block polewards flux and slow microtubule turnover in Xenopus extract spindles (Sawin and Mitchison, 1991, 1994; Waterman-Storer et al., 1998). Eg5 speckle kymographs in AMPPNP show vertical streaks of stationary Eg5 speckles that persist for over 200 s (Fig. 3 b). The same is true for tubulin speckle kymographs (Waterman-Storer et al., 1998). This is consistent with our Eg5 probe being a functional kinesin capable of coupling ATP hydrolysis to its microtubule association. This also suggests that the dynamic association of labeled Eg5 with microtubules we observed in normal spindles depends on ATPase activity of Eg5.
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Second, we assembled spindles in the presence of p50 dynamitin, which leads to the disruption of the dynactin complex and the formation of spindles with abnormally splayed poles and reduced microtubule density (Echeverri et al., 1996; Heald et al., 1997). p50 treatment does not alter microtubule polarity (Merdes et al., 2000) or block polewards flux in Xenopus extract spindles (Kapoor, T.M., A. Desai, P. Maddox, E.D. Salmon, T.J. Mitchison, personal communication). In p50-treated spindles, the distribution of endogenous Eg5 (Fig. 6 , a and b) detected by immunofluorescence and added labeled Eg5 (Fig. 6, c and d) was severely redistributed relative to microtubules (Fig. 7 c), becoming enriched in the center of the spindle and depleted at the poles. These data suggest a role for dynactin/dynein in positioning Eg5 in spindles. The similar distribution of the Eg5 probe and endogenous Eg5 in perturbed and unperturbed spindles suggests that the labeled recombinant Eg5 is a faithful reporter for the localization of endogenous Eg5.
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Discussion |
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We note a possible discrepancy between our results and a recent paper in which Eg5 dynamics were probed in microtubule asters induced by an allele of the GTPase, Ran (Wilde et al., 2001). In that paper, low concentrations of a fluorescently labeled antibody raised to the stalk domain of Eg5 were added to the extract to probe Eg5 dynamics. Speckles were observed, and a fraction of them moved towards the plus end of microtubules at 2.8 µm/min. In contrast, we observed essentially no movement of directly labeled Eg5 away from poles in regions near the poles of bipolar spindles. The difference may be due simply to differences in the behavior of Eg5 in bipolar spindles and Ran asters. It is not known if microtubules in Ran asters flux towards the center of the aster. It is also possible that the antibody technique may not provide a nonperturbing probe of Eg5 dynamics. Although the labeled antibody did not inhibit bipolar spindle formation at the low concentrations used for fluorescent speckle microscopy, antibodies to the same region of Eg5 are known to induce spindle collapse when added at higher concentrations (Sawin et al., 1992). It is possible that antistalk antibodies could induce dissociation of Eg5 tetramers from a binding site in spindles, and thus induce artificial plus enddirected motility of those Eg5 tetramers to which the antibody has bound. Analysis of antibody speckles in bipolar spindles and directly labeled Eg5 in Ran asters could help address these differences.
Our principle result is the observation of populations of Eg5 in spindles that are static during intervals over which microtubules move detectably poleward. We believe that this observation has important implications for the mechanism of Eg5 targeting to spindles, and by extension, for more general issues of spindle organization and Eg5 function. We considered four possible explanations for immobilization of Eg5 in spindles. First, Eg5 might be free in spindles, but exhibit limited diffusion due simply to high local viscosity or viscoelasticity. We think this unlikely for several reasons. Using the Einstein-Stokes equation, we estimate that spindle viscosity would have to be >70 Poise to keep individual Eg5 tetramers (Stokes radius of 13.5 nm) within regions resolvable by light microscopy (300 nm2) for 20 s (Alexander and Rieder, 1991). The maximum published estimate for spindle viscosity is 20-fold lower (Alexander and Rieder, 1991). Our observation that Eg5 can exhibit bulk exchange in and out of spindles over a distance of microns, on a time scale of seconds, argues against a purely viscoelastic mechanism for immobilization. Finally the density of microtubules in spindles with micron scale gaps between bundles is insufficient to act as a sieve for molecules the size of Eg5. We believe that the combined speckle and bulk turnover are better fit by a model in which Eg5 interacts transiently with some scaffold that causes it to be immobilized, but it diffuses freely when not interacting with this scaffold.
Second, individual Eg5 tetramers might walk, using their motor activity, towards the plus ends of microtubules at exactly the rate that the microtubules flux polewards. This could result in walking in place, as on a treadmill. This interpretation is complicated by the fact that microtubule orientation varies across the spindle (uniform polarity near the poles, mixed polarity near the center), and we found that Eg5 is equally immobile all over the spindle. If this interpretation were correct, Eg5 distribution in spindles would be very sensitive to inhibition of its motor function. Our monastrol data (Fig. 5) shows this is not the case, so we disfavor this interpretation.
Third, Eg5 might associate with a subset of microtubules that are not moving. Both photoactivation of fluorescence (Sawin and Mitchison, 1991) and speckle imaging (Fig. 2, a and e) suggest that the great majority of microtubules in Xenopus spindles are moving polewards. Both methods are biased towards observing flux in more stable microtubules, and it is possible that very recently polymerized microtubules, or a subset of unusually dynamic microtubules, are not fluxing. We did observe occasional tubulin speckles that moved poleward slowly (0.25 µm/min), and these tended to be clustered towards the pole, where recently polymerized microtubules may be enriched. The pole is also enriched in microtubule segments aligned in part along the z axis (perpendicular to the image plane), which would slow apparent movement rates along the x, y axis. Again, the observation that Eg5 was equally static all over the spindle argues against differential association with nonfluxing microtubules near poles. Since there is no evidence that Eg5 selectively associates with subsets of microtubules, and no definite evidence for a static subset, we disfavor this interpretation.
Although we cannot rule out the three preceding interpretations, we favor a fourth, that Eg5 physically interacts with some static, nonmicrotubule spindle component that we postulate corresponds to the hypothetical spindle matrix (Fig. 8 a). Our data provide few clues as to the identity of this hypothetical matrix. We do not think actin is involved, since spindle organization and dynamics in Xenopus extracts are not affected by cytochalasin D (which was present in all our experiments). We have not detected a high-affinity interaction between Eg5 and other spindle proteins, but two specific interactions are worth considering, Eg5dynactin and Eg5Eg5. Two-hybrid data revealed that Eg5 may physically interact with the dynein-targeting complex dynactin (Blangy et al., 1997), and we found that disruption of dynactin with p50 caused relocalization of Eg5. However, disruption of dynactin did not block targeting of Eg5 to spindles. Understanding the nature of the Eg5dynactin interaction requires more data. We currently suspect that dynactin influences Eg5 localization indirectly, for example through positioning an unknown matrix element (Fig. 8 a). Reversible polymerization of Eg5 tetramers in the spindle to form dynamic higher order complexes or filaments with reduced diffusion could also account for much of our data. Beyond dynactin and Eg5, we can only speculate as to the biochemical nature of a putative static spindle matrix. It might contain spindle-specific proteins such as NuMA (Dionne et al., 1999), TPX2 (Wittmann et al., 2000), or homologues of the Drosophila protein, Skeletor (Walker et al., 2000). It could also contain, or be made of, proteins that permeate the whole cytoplasm as well as the spindle or membrane systems. It could bind Eg5 directly, or perhaps immobilize it indirectly, by viscoelastic tethering (Seksek et al., 1997; Luby-Phelps, 2000; Papadopoulos et al., 2000). Identification of the matrix, if it exists, will require new biochemical experiments. Fluorescent speckle microscopy provides a tool to probe the dynamics of new spindle components, and thus determine if they interact primarily with fluxing microtubules or with a static matrix.
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Materials and methods |
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Spindle assembly in vitro
Spindles were assembled in cytostatic factorarrested Xenopus laevis egg extracts cycled through interphase to replicate the DNA and centrosomes as described (Desai et al., 1999). p50 dynamitin was added with the second addition of cytostatic factorarrested extract during spindle assembly.
Fluorescent speckle microscopy and analysis
The amount of labeled protein added to Xenopus egg extracts was optimized for each protein preparation. For microtubule fluorescent speckle microscopy, Texas redlabeled tubulin was used at concentrations ranging from 6 to 30 nM (0.9 fluorophores per tubulin dimer). For Eg5 fluorescent speckle microscopy, Texas redlabeled Eg5 was typically used at 100150 nM. Samples were imaged with a 100x objective (Plan Apo, Nikon; NA 1.4) and a camera (MicroMax cooled CCD, Princeton Instruments; bin 2 x 2). Time-lapse images (400-ms exposure) were collected at 1-s intervals using a digital imaging system controlled by Metamorph software (Universal Imaging Corp.). Kymographs were prepared from unprocessed images by selecting a seven pixel wide line extending from one spindle pole to the other. In brief, a line seven pixels wide (x-dimension) was drawn across the spindle (y-dimension). The software recorded the maximum intensity pixel across the x-dimension for each point along the line (y-dimension). The finished image consists of the above measurement performed for each timepoint, placed one after another by the software to display intensity differences over time. Slopes of streaks of correlated speckles were measured to determine rates of movement relative to the spindle poles. Five spindles from four independent experiments were analyzed to determine the rates of Eg5 and tubulin speckle movements. Each spindle was divided into two halves and rates for seven speckles uniformly distributed in each half were measured relative to the corresponding spindle pole. The velocities were binned into 0.5 µm/min increments to generate velocity distribution histograms.
Turnover of Eg5
500 nM Alexa 488labeled tubulin was added to spindles assembled in Xenopus egg extracts cycled through interphase. At the completion of assembly, Texas redlabeled Eg5 was added to the extracts (140 nM) and samples were removed at different time points, diluted, and fixed (7.5% formaldehyde). The samples were imaged using a Nikon 40x Plan Fluor objective (NA 0.75) and a Princeton Instruments MicroMax cooled CCD camera. The ratio of the average intensities of the Texas red signal and the Alexa fluor signal (both corrected for sample background) was determined for five spindles for each time point. The data for each experiment was normalized to the ratio at the 300 s time point. Averages of three independent experiments are shown with SD. For the "dilution" experiment, spindles in extract with labeled Eg5 incorporated to saturation and labeled tubulin were mixed with 5 vol of extracts with assembled spindles, labeled tubulin, and no labeled Eg5. After 5 min, samples were fixed and processed for imaging.
Linescans
Linescans were generated using functions in Metamorph software (Univeral Imaging Corp.). The average intensity over a 10 pixel wide region draw across the unprocessed image is calculated for each channel of fluorescence by the software. Regions selected for linescans extended through the center of the perturbed or control spindle images, parallel to the long axis of the structure. Overlays of pseudocolored graphs were prepared using Adobe Photoshop® 6.0.
Viscosity calculation
The Einstein-Stokes equation was used to determine the viscosity of the spindle with time (t) = 20 s, temperature (T) = 300 K, mean free path (X) = 300 nm, Stokes radius (a) = 13.5 nm, and k = Boltzman constant (viscosity = kTt/3aX2).
Online supplemental material
Data for immunodepletion and rescue by add-back of recombinant Eg5 is provided. Time-lapse videos corresponding to spindles shown in Fig. 2 and additional videos for Eg5 and microtubule fluorescent speckle microscopy are also provided. The videos are available at http://www.jcb.org/content/vol154/issue6.
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Footnotes |
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* Abbreviation used in this paper: AMPPNP, adenylimidodiphosphate.
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Acknowledgments |
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T.M. Kapoor is a Runyon-Winchell Fellow. This work was supported by grants from the National Institute of General Medical Sciences (39565) to Timothy J. Mitchison.
Submitted: 4 June 2001
Revised: 17 July 2001
Accepted: 13 August 2001
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References |
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Alexander, S.P., and C.L. Rieder. 1991. Chromosome motion during attachment to the vertebrate spindle: initial saltatory-like behavior of chromosomes and quantitative analysis of force production by nascent kinetochore fibers. J. Cell Biol. 113:805815.[Abstract]
Blangy, A., H.A. Lane, P. d'Herin, M. Harper, M. Kress, and E.A. Nigg. 1995. Phosphorylation by p34cdc2 regulates spindle association of human Eg5, a kinesin-related motor essential for bipolar spindle formation in vivo. Cell. 83:11591169.[Medline]
Blangy, A., L. Arnaud, and E.A. Nigg. 1997. Phosphorylation by p34cdc2 protein kinase regulates binding of the kinesin-related motor HsEg5 to the dynactin subunit p150. J. Biol. Chem. 272:1941819424.
Desai, A., A. Murray, T.J. Mitchison, and C.E. Walczak. 1999. The use of Xenopus egg extracts to study mitotic spindle assembly and function in vitro. Methods Cell Biol. 61:385412.[Medline]
Dionne, M.A., L. Howard, and D.A. Compton. 1999. NuMA is a component of an insoluble matrix at mitotic spindle poles. Cell Motil. Cytoskeleton. 42:189203.[Medline]
Echeverri, C.J., B.M. Paschal, K.T. Vaughan, and R.B. Vallee. 1996. Molecular characterization of the 50-kD subunit of dynactin reveals function for the complex in chromosome alignment and spindle organization during mitosis. J. Cell Biol. 132:617633.[Abstract]
Enos, A.P., and N.R. Morris. 1990. Mutation of a gene that encodes a kinesin-like protein blocks nuclear division in A. nidulans. Cell. 60:10191027.[Medline]
Forer, A., T. Spurck, and J. Pickett-Heaps. 1997. Ultraviolet microbeam irradiation of spindle fibres in crane-fly spermatocytes and newt epithelial cells: resolution of previously conflicting observations. Protoplasma. 197:230240.
Gheber, L., S.C. Kuo, and M.A. Hoyt. 1999. Motile properties of the kinesin-related Cin8p spindle motor extracted from Saccharomyces cerevisiae cells. J. Biol. Chem. 274:95649572.
Gordon, M., L. Howard, and D. Compton. 2001. Chromosome movement in mitosis requires microtubule anchorage at spindle poles. J. Cell Biol. 152:425434.
Hagan, I., and M. Yanagida. 1990. Novel potential mitotic motor protein encoded by the fission yeast cut7+ gene. Nature. 347:563566.[Medline]
Heald, R., R. Tournebize, A. Habermann, E. Karsenti, and A. Hyman. 1997. Spindle assembly in Xenopus egg extracts: respective roles of centrosomes and microtubule self-organization. J. Cell Biol. 138:615628.
Heck, M.M., A. Pereira, P. Pesavento, Y. Yannoni, A.C. Spradling, and L.S. Goldstein. 1993. The kinesin-like protein KLP61F is essential for mitosis in Drosophila. J. Cell Biol. 123:665679.[Abstract]
Hoyt, M.A., L. He, K.K. Loo, and W.S. Saunders. 1992. Two Saccharomyces cerevisiae kinesin-related gene products required for mitotic spindle assembly. J. Cell Biol. 118:109120.[Abstract]
Hyman, A., D. Drechsel, D. Kellogg, S. Salser, K. Sawin, P. Steffen, L. Wordeman, and T. Mitchison. 1991. Preparation of modified tubulins. Methods Enzymol. 196:478485.[Medline]
Inoue, S., and E.D. Salmon. 1995. Force generation by microtubule assembly/disassembly in mitosis and related movements. Mol. Biol. Cell. 6:16191640.[Medline]
Kapoor, T.M., T.U. Mayer, M.L. Coughlin, and T.J. Mitchison. 2000. Probing spindle assembly mechanisms with monastrol, a small molecule inhibitor of the mitotic kinesin, Eg5. J. Cell Biol. 150:975988.
Leslie, R.J., R.B. Hird, L. Wilson, J.R. McIntosh, and J.M. Scholey. 1987. Kinesin is associated with a nonmicrotubule component of sea urchin mitotic spindles. Proc. Natl. Acad. Sci. USA. 84:27712775.[Abstract]
Luby-Phelps, K. 2000. Cytoarchitecture and physical properties of cytoplasm: volume, viscosity, diffusion, intracellular surface area. Int. Rev. Cytol. 192:189221.[Medline]
Maddox, P.S., K.S. Bloom, and E.D. Salmon. 2000. The polarity and dynamics of microtubule assembly in the budding yeast Saccharomyces cerevisiae. Nat. Cell Biol. 2:3641.[Medline]
Mayer, T.U., T.M. Kapoor, S.J. Haggarty, R.W. King, S.L. Schreiber, and T.J. Mitchison. 1999. Small molecule inhibitor of mitotic spindle bipolarity identified in a phenotype-based screen. Science. 286:971974.
Mazia, D. 1961. Mitosis and the physiology of Cell Division. In The Cell. Biochemistry, Physiology and Morphology. J. Brachvhet and A.C. Mirsky, editors. Academic Press, NY. 61412.
McIntosh, J.R., P.K. Hepler, and D.G. Van Wie. 1969. Model for mitosis. Nature. 224:659663.
Merdes, A., R. Heald, K. Samejima, W.C. Earnshaw, and D.W. Cleveland. 2000. Formation of spindle poles by dynein/dynactin-dependent transport of NuMA. J. Cell Biol. 149:851862.
Mitchison, T.J. 1989. Polewards microtubule flux in the mitotic spindle: evidence from photoactivation of fluorescence. J. Cell Biol. 109:637652.[Abstract]
Mitchison, T., and M. Kirschner. 1984. Dynamic instability of microtubule growth. Nature. 312:237242.[Medline]
Mountain, V., C. Simerly, L. Howard, A. Ando, G. Schatten, and D.A. Compton. 1999. The kinesin-related protein, HSET, opposes the activity of Eg5 and cross-links microtubules in the mammalian mitotic spindle. J. Cell Biol. 147:351366.
Nicklas, R.B. 1988. The forces that move chromosomes in mitosis. Annu. Rev. Biophys. Biophys. Chem. 17:431449.[Medline]
Papadopoulos, S., K.D. Jurgens, and G. Gros. 2000. Protein diffusion in living skeletal muscle fibers: dependence on protein size, fiber type, and contraction. Biophys. J. 79:20842094.
Pickett-Heaps, J.D., D.H. Tippit, and K.R. Porter. 1982. Rethinking mitosis. Cell. 29:729744.[Medline]
Pickett-Heaps, J., T. Spurck, and D. Tippit. 1984. Chromosome motion and the spindle matrix. J. Cell Biol. 99:137s143s.
Salmon, E.D., and R.R. Segall. 1980. Calcium-labile mitotic spindles isolated from sea urchin eggs (Lytechinus variegatus). J. Cell Biol. 86:355365.[Abstract]
Sawin, K.E., and T.J. Mitchison. 1991. Poleward microtubule flux mitotic spindles assembled in vitro. J. Cell Biol. 112:941954.[Abstract]
Sawin, K.E., and T.J. Mitchison. 1994. Microtubule flux in mitosis is independent of chromosomes, centrosomes, and antiparallel microtubules. Mol. Biol. Cell. 5:217226.[Abstract]
Sawin, K.E., and T.J. Mitchison. 1995. Mutations in the kinesin-like protein Eg5 disrupting localization to the mitotic spindle. Proc. Natl. Acad. Sci. USA. 92:42894293.[Abstract]
Sawin, K.E., K. LeGuellec, M. Philippe, and T.J. Mitchison. 1992. Mitotic spindle organization by a plus-end-directed microtubule motor. Nature. 359:540543.[Medline]
Schnackenberg, B.J., A. Khodjakov, C.L. Rieder, and R.E. Palazzo. 1998. The disassembly and reassembly of functional centrosomes in vitro. Proc. Natl. Acad. Sci. USA. 95:92959300.
Seksek, O., J. Biwersi, and A.S. Verkman. 1997. Translational diffusion of macromolecule-sized solutes in cytoplasm and nucleus. J. Cell Biol. 138:131142.
Sharp, D.J., K.L. McDonald, H.M. Brown, H.J. Matthies, C. Walczak, R.D. Vale, T.J. Mitchison, and J.M. Scholey. 1999. The bipolar kinesin, KLP61F, cross-links microtubules within interpolar microtubule bundles of Drosophila embryonic mitotic spindles. J. Cell Biol. 144:125138.
Sharp, D.J., G.C. Rogers, and J.M. Scholey. 2000. Microtubule motors in mitosis. Nature. 407:4147.[Medline]
Walker, D.L., D. Wang, Y. Jin, U. Rath, Y. Wang, J. Johansen, and K.M. Johansen. 2000. Skeletor, a novel chromosomal protein that redistributes during mitosis provides evidence for the formation of a spindle matrix. J. Cell Biol. 151:14011412.
Waterman-Storer, C.M., and E.D. Salmon. 1998. How microtubules get fluorescent speckles. Biophys. J. 75:20592069.
Waterman-Storer, C.M., and E.D. Salmon. 1999. Fluorescent speckle microscopy of microtubules: how low can you go? FASEB J. 13:S225S230.[Medline]
Waterman-Storer, C.M., A. Desai, J.C. Bulinski, and E.D. Salmon. 1998. Fluorescent speckle microscopy, a method to visualize the dynamics of protein assemblies in living cells. Curr. Biol. 8:12271230.[Medline]
Wilde, A., S.B. Lizarraga, L. Zhang, C. Wiese, N.R. Gliksman, C.E. Walczak, and Y. Zheng. 2001. Ran stimulates spindle assembly by altering microtubule dynamics and the balance of motor activities. Nat. Cell Biol. 3:221227.[Medline]
Wittmann, T., and T. Hyman. 1999. Recombinant p50/dynamitin as a tool to examine the role of dynactin in intracellular processes. Methods Cell Biol. 61:137143.[Medline]
Wittmann, T., M. Wilm, E. Karsenti, and I. Vernos. 2000. TPX2, A novel Xenopus MAP involved in spindle pole organization. J. Cell Biol. 149:14051418.
Wittmann, T., A. Hyman, and A. Desai. 2001. The spindle: a dynamic assembly of microtubules and motors. Nat. Cell Biol. 3:E28E34.[Medline]