* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
Download as a PDF
Survey
Document related concepts
Tissue engineering wikipedia , lookup
Endomembrane system wikipedia , lookup
Cytoplasmic streaming wikipedia , lookup
Extracellular matrix wikipedia , lookup
Cell encapsulation wikipedia , lookup
Cellular differentiation wikipedia , lookup
Programmed cell death wikipedia , lookup
Cell culture wikipedia , lookup
Organ-on-a-chip wikipedia , lookup
Cell growth wikipedia , lookup
Spindle checkpoint wikipedia , lookup
List of types of proteins wikipedia , lookup
Transcript
Sustained Microtubule Treadmilling in Arabidopsis Cortical Arrays Sidney L. Shaw1, Roheena Kamyar2 and David W. Ehrhardt2* 1Department of Biological Sciences Stanford University Stanford, CA 94305 USA 2Carnegie Institution of Washington Department of Plant Biology Stanford, CA 94305 USA *To whom correspondence should be addressed [email protected] 1 Abstract Plant cells create highly structured microtubule arrays at the cell cortex without a central organizing center to anchor the microtubule ends. In vivo imaging of individual microtubules in Arabidopsis plants revealed that new microtubules initiated at the cell cortex and exhibited dynamics at both ends. Polymerization-biased dynamic instability at one end and slow depolymerization at the other resulted in sustained microtubule migration across the cell cortex by a hybrid treadmilling mechanism. This motility caused widespread microtubule repositioning and contributed to changes in array organization through microtubule reorientation and bundling. 2 The microtubule cytoskeleton plays a key role in plant cell morphogenesis and multicellular development. Disruption of plant microtubule organization by drugs or through mutation causes defects ranging from changes in cell shape to a dramatic loss of organ form (1-5). The cortical microtubule array is proposed to inuence cell shape by guiding the deposition of new cell wall polymers (4,6,7). Aligning the cellulose microbrils in the cell wall restricts cell elongation, resulting in anisotropic cell wall expansion and the acquisition of specialized cell shapes (7). How the cortical microtubules are created and positioned to form organized arrays is not known. Plant cortical microtubule arrays are dynamic structures (8), continually reorganizing in response to environmental and developmental cues (2,9). In epidermal cells of the root or shoot axis, interphase microtubules show a progressive change in organization following cell division. Unlike interphase arrays in animal or fungal cells, the plant cortical array does not radiate from a central organizer. Microtubules rst appear at the cell cortex in a disordered arrangement and then form remarkable transverse helical arrays that change pitch as the cell expands (6,9,10). Several mechanisms have been proposed for the creation and dynamic organization of the cortical arrays (2,11-15), including de novo microtubule polymerization in a preferred orientation, transport of microtubules originating at the nuclear surface to dened cortical positions, lateral and axial sliding (translocation) of existing cortical microtubules into new positions, and the migration of polymers to new positions by the balanced addition and removal of subunits at the microtubule ends (treadmilling). To analyze the behavior of individual cortical microtubules and to ask which of these mechanisms contribute to cortical array organization, we created tubulin–green uorescent protein (GFP) fusions that permit imaging of individual microtubules in transgenic Arabidopsis plants. 3 Individual Microtubule Behaviors Time-lapsed confocal imaging of Arabidopsis epidermal cells expressing GFP-tubulin fusion proteins (n = 33 cells, 4.1min average duration at 3.85sec intervals) revealed discrete sites of apparent microtubule initiation at the cell cortex (Fig. 1A,B). Microtubules that depolymerized to visible completion rarely showed recovery (48 of 50 events with >2min of imaging after depolymerization), suggesting that most initiation sites represent de novo origins. Initiation sites were scattered throughout the observed area of the cell cortex and appeared both in association with existing microtubules and in regions with no other detectable microtubules. In several cases, multiple microtubules polymerized from the same site (Fig. 1A, Movies S1,S2). Limited examples of severing or breakage in elongated microtubules were recorded, though in 24 of 51 observed severing events at least one of the resulting microtubules depolymerized to extinction. In observations of 30 cells, we failed to observe a microtubule emerging from the cytosol to join the cortical array, while we observed 71 cortical initiation events in these same cells. Thus, the majority of new microtubules in mature interphase arrays were likely created at the cell cortex and did not come from interior organizing centers such as the nuclear surface. Consistent with this, AtSpc98, a proposed microtubule organizing center component, has been localized to the plasma membrane of Arabidopsis cells (16). New microtubules did not remain anchored to their site of initiation (Fig. 1A; Movies S1-S3). Initiating microtubules extended several micrometers before either shortening to extinction (7/43 initiation events) or moving away from the initiation site (36/43). Motility was unidirectional, with single microtubules moving most commonly in shallow arcs (Fig. 1B, C; Movies S1-S3), often changing trajectory several times on the cell cortex during the course of observation (2-8 min.). In cases where multiple microtubules initiated from the same location, they often departed at diverging angles (Fig. 1A,B; Movies S1-S3). Time-lapse images of digitally linearized microtubules further illustrated 4 the unidirectional motility and revealed markedly different dynamic properties for the leading and lagging ends (Fig. 1D-F, and below). After moving away from their sites of initiation, a subset of polymers shortened completely from the leading end, while the remainder migrated across the cell cortex and gradually elongated or became incorporated into microtubule bundles (Fig. 1D-F). To test whether the microtubule motility was due to sliding (translocation of the polymer) or treadmilling, the GFP-labeled microtubules were marked by photobleaching (n=12 cells, >50 single microtubules). While microtubules remained motile after photobleaching, the photobleached marks maintained xed positions with reference to the cell (Fig. 2A, Movie S4); in no case did we observe the movement of a photobleached mark on a single microtubule (Fig. 2B-D). Thus single microtubules were xed in place at the cell cortex and the apparent microtubule motility resulted from polymerization and depolymerization at the ends and not from translocation of the intact microtubule polymer. Motile microtubules were observed either to cross over other microtubules or to incorporate into bundles (Fig. 1C, Movies S1-S3). Bundling initiated when the leading end of a motile microtubule contacted another microtubule or bundle and changed trajectory to become co-aligned. Progressive changes in uorescence intensity along the encountered polymer suggest that the leading end continued to polymerize along the bundle after initial contact. Depolymerization from the lagging end completed the bundling process by consuming the unbundled portion of the microtubule. Photobleached marks made on microtubule bundles typically recovered rapidly but did not move (Fig. 2A,B,D; Movie S4). Thus bundled microtubules remained dynamic, with the observed dynamic behavior being caused by polymerization and depolymerization and not by the sliding of bundled microtubules. 5 To ask if lateral movements contributed to microtubule positioning, we drew a linear transect across the cell image to sample random discrete locations along multiple microtubules (Fig. 2E). The uorescence signal along this transect from a time-lapse series of images was projected as a kymograph to analyze the stability of microtubule position over time (Fig. 2F). Parallel, vertical lines in the kymograph indicated that the cortical microtubules showed almost no lateral translocation, despite rapid cytoplasmic streaming. This lateral stability was evident even when single microtubules displayed no overlap with other cortical microtubules, establishing that stabilization of microtubule position did not rely on inter-microtubule crosslinking. The stability of the uorescence signal in the focal plane over the duration of the experiment further indicated that microtubules did not show detectable movement on and off the cell cortex. Thus the majority of the microtubules in the observed cortical arrays were strongly associated with the cell cortex, as proposed previously from ultrastructural studies (17), and lateral translocation of both single and bundled microtubules was either rare or too slow to detect over a 6min observation interval. Exceptions to cortical association were found when a microtubule end moved rapidly out of focus and into the streaming cytoplasm (Fig. 2G, Movies S1-S3). The abrupt loss of cortical association occurred at the leading end of the motile microtubule (69 of 71 events, n = 33 cells) and was observed almost exclusively for single microtubules (70 of 71 events), not for microtubules in bundles. The detachment of a free end resulted in either re-association with the cortex (34 of 71 events), often reorienting the microtubule, or in complete depolymerization (37 of 71 events). Thus cortical attachment may be important for array organization as loss of attachment has signicant consequences for polymer stability and orientation. Also, bundling might protect microtubules from cortical detachment, possibly through intra-microtubule crosslinking by other proteins (18). 6 Polymerization Dynamics To investigate how polymerization dynamics contribute to cortical array behavior, we measured the dynamic properties of both the cortical array and the individual microtubule ends. Using uorescence redistribution after photobleaching (FRAP), we measured a recovery halftime of 58.95s (n = 27 cells, std = 14.7, sem = 2.83) from epidermal cells in the hypocotyl (Fig. 3AB, Movie S5). This recovery time is approximately 4 times faster than that measured in animal interphase arrays (19, supplemental materials). These data obtained with a GFP-tubulin fusion protein expressed in Arabidopsis conrm previous FRAP results from Tradescantia stamen hair cells injected with uorescent animal tubulin (8). To determine how individual microtubule polymerization dynamics contribute to cortical array turnover and behavior, we measured velocities of growth and shortening (Fig. 3C), and the transition frequencies between growth, shortening, and pause states for single microtubules where both ends were clearly visible (Table 1, n = 78 microtubules from 18 cells). The microtubule end leading the unidirectional motility displayed 5 fold more net polymerization-depolymerization per unit time (dynamicity) than the lagging end (Table 1), conrming that the two ends had distinct dynamic properties. The leading end showed persistent phases of both growth and shortening, the rate of shortening being faster on average (5.88+/-5.07µm/min) than the rate of growth (3.69+/-1.90µm/min). Catastrophe (0.043s-1) and rescue (0.082s-1) frequencies, however, favored time spent in growth (Table 1), resulting in a net gain in polymer at the leading end. Lagging end growth was slow (1.96+/-1.24µm/min) and rare (Table 1), possibly falling within the error of the measurement technique. The lagging end spent approximately the same amount of time shortening as did the leading end, but depolymerization occured more slowly (2.78+/-2.13µm/min) and commonly transitioned to a pause state (rescue = 0.128s-1, 7 Table 1). We measured an increase in total polymer for the microtubules sampled in this study (0.36µm/min per microtubule). This increase arose at least in part because microtubules that elongated and then associated into bundles could no longer be measured (Fig. 1C), but may also reect an actual bias in the dynamics of the population of sampled microtubules. Conclusions Microtubules were observed to migrate across the cortex of Arabidopsis epidermal cells using a hybrid treadmilling mechanism. Treadmilling motility was not caused by pronounced dynamic instability at both polymer ends (20), nor steady gain at one end and steady loss at the other (21). Rather, motility was the net result of slow, intermittent depolymerization at the lagging end, coupled with polymerization-biased dynamic instability at the leading end. Further, the discovery of dynamic instability as the dominant mode of dynamic behavior in the Arabidopsis interphase arrays suggests that dynamic instability is integral to the organization of both centriolar and acentriolar interphase arrays. Treadmilling events have been observed in animal cells and cytoplasts when microtubules escaped from the centrosome or suffered breakage events (22-27). These events are relatively rare and short lived, ending by rapid depolymerization from the minus end (23) or by depolymerization from the plus end when the minus end is stabilized (24,25). By contrast, treadmilling motility in Arabidopsis microtubules is neither rare nor short lived. The majority of microtubules we measured (50 of 78) showed strictly-dened treadmilling for 22.5% of the observation interval. The lagging end of the microtubule seldom remained stable over time (6/77 microtubules) and complete depolymerization of microtubules was only observed to occur from the leading end (n=50 of 50 from 10 cells), 8 even in cases of severing. The slow and intermittent depolymerization at the lagging end of plant interphase microtubules suggests that sustained treadmilling motility results from careful regulation of minus end. The extent of the treadmilling, the creation of microtubule bundles through treadmilling motility, and the absence of other observed mechanisms for polymer repositioning together suggest that treadmilling motility makes a signicant contribution to the organization of the cortical array. 9 References and Notes 1. C. Lloyd, J. Chan, Plant Cell 14, 2319 (2002). 2. R. J. Cyr, Ann Rev Cell Biol 10, 153 (1994). 3. A. Bichet, T. Desnos, S. Turner, O. Grandjean, H. Hofte, Plant J 25, 137 (2001). 4. A. T. Whittington et al., Nature 411, 610 (2001). 5. S. Thitamadee, K. Tuchihara, T. Hashimoto, Nature 417, 193 (2002). 6. T. I. Baskin, Protoplasma 215, 150 (2001). 7. P. B. Green, J Cell Biol 27, 343-63 (1965). 8. J. M. Hush, P. Wadsworth, D. A. Callaham, P. K. Hepler, J Cell Sci 107, 775 (1994). 9. C. L. Granger, R. J. Cyr, Protoplasma 216, 201 (2001). 10. M. Yuan, P. J. Shaw, R. M. Warn, C. W. Lloyd, PNAS 91, 6050 (1994). 11. G. O. Wasteneys, J Cell Sci 115, 1345 (Apr 1, 2002). 12. L. Clayton, C. M. Black, C. W. Lloyd, J Cell Biol 101, 319 (1985). 13. H. Shibaoka, in The cytoskeletal basis of plant growth and form C. W. Lloyd, Ed. (Academic Press, London, 1991) pp. 159. 14. S. M. Wick, Cell Biol Int Rep 9, 357 (1985). 15. B. E. Gunning, S. M. Wick, J Cell Sci Suppl 2, 157 (1985). 16. M. Erhardt et al., J Cell Sci 115, 2423 (2002). 17. A. R. Hardham, B. E. Gunning, J Cell Biol 77, 14 (1978). 18. J. Chan, C. G. Jensen, L. C. Jensen, M. Bush, C. W. Lloyd, PNAS 96, 14931 (1999). 19. W. M. Saxton et al., J Cell Biol 99, 2175 (1984). 20. T. Mitchison, M. Kirschner, Nature 312, 237 (1984). 21. C. M. Waterman-Storer, E. D. Salmon, Curr Biol 7, R369 (1997). 22. T. J. Keating, J. G. Peloquin, V. I. Rodionov, D. Momcilovic, G. G. Borisy, PNAS 10 94, 5078 (1997). 23. C. M. Waterman-Storer, E. D. Salmon, J Cell Biol 139, 417 (1997). 24. V. I. Rodionov, G. G. Borisy, Science 275, 215 (1997). 25. A. M. Yvon, P. Wadsworth, J Cell Sci 110, 2391 (1997). 26. I. A. Vorobjev, V. I. Rodionov, I. V. Maly, G. G. Borisy, J Cell Sci 112, 2277 (1999). 27. V. Rodionov, E. Nadezhdina, G. Borisy, PNAS 96, 115 (1999). 28. The authors wish to thank D. Allen for help in collecting and propagating seed stocks; T. Salmon, C. Somerville, W. Briggs, T. Stearns, J. Theriot, and S. Cutler for useful discussions and comments; and S. Long for her generous support. This work was supported by the Carnegie Institution of Washington (DWE, RK), the Howard Hughes Medical Institute and the Department of Energy (grant #DEFG03-90ER2001, SLS). 11 231.0 Time (sec) 173.5 115.5 57.8 0 A B C D E 12 F Figure 1. Microtubule initiation and unidirectional motility. (A) Time series (left to right) of two new microtubules (solid and open arrowheads) polymerizing from a site at the cell cortex (arrow), and diverging from this origin at different angles . (B) A newly polymerized microtubule (solid arrowhead) detaching from a cortical site of origin (arrow). After detachment, a second microtubule (open arrowhead) is initiated at the same location. (C) Motile microtubule (solid arrowhead) crossing one microtubule (open arrowhead) before encountering a second polymer and bundling (arrows). (D-F) Kymographs showing three single microtubules from the same cell, digitally linearized and moving from left to right. Single microtubules show dynamic instability at the leading end and primarily slow shortening at the lagging end. Note that the microtubule in (D) depolymerizes to extinction from the leading end at 170sec. Scale bar = 2.5µm, intervals between images = 7.6- 15.2 seconds (A,B), 3.8 seconds (D-F). 13 A B ] C ] D F G 14 Time E ] Figure 2. Microtubules associate with the cortex and move by treadmilling. (A) Marks generated by photobleaching GFP-tubulin labeled microtubules do not move, demonstrating that microtubule motility does not occur by translocation of existing polymer. Scale bar = 5µm. (B-D) Kymographs representing polymers from the entire time sequence in (A) show that bleach marks remain xed in place. (B) The bleached zone does not spread in microtubule bundles and the bleach border does not travel into the bleached zone. (C) Single microtubule migrating from left to right with xed bleach zone. (D) Bundle showing uorescence recovery via constituent microtubules. Vertical bar = 2min, horizontal bar = 2.5µm. (E) First image in a time-lapsed series (3.8 sec intervals) of a cortical microtubule array. (F) Kymograph of a linear transect across this cell (gray line in E) at each time interval. Straight lines in the kymograph indicate no lateral movement of microtubules during the 6.5min experiment. Vertical bar = 2.5min (F), horizontal bar = 5µm (E, F). (G) Detachment from the cell cortex was observed as rapid, lateral and out of focus movement of microtubule ends (arrows), resulting in re-attachment and re-orientation (arrowheads) or depolymerization. Total time interval for image series = 96 seconds, scale bar = 2.5µm. 15 A 1.1 1.0 Fraction of F0 0.9 0.8 0.7 0.6 T1/2 = 58.95s 0.5 0.4 350 300 250 200 150 100 50 B 0 0.3 Time (sec) Number of time intervals 400 Leading End 300 v = 3.69 v = 5.80 200 100 0 Lagging End 100 200 300 400 C v = 2.78 v = 1.96 (2024) 20 16 15 10 5 0 -5 -10 -15 -20 -25 -30 Velocity (mm/min) Figure 3. Microtubule dynamics measurements. (A) Cortical microtubule array in a hypocotyl cell immediately after laser-mediated photobleaching of a 10 µm circle. Scale bar = 10 µm. (B) Fluorescence redistribution after photobleaching. Two images were taken prior to photobleaching and the remainder at 8-12s intervals following bleaching. Fluorescence recovery measurements (mean +/- std, n = 27 cells) are corrected for photobleaching and normalized to the initial uorescence value before tting with an exponential function (solid line, (8)). The curve is t without the rst measurement following photobleaching (red symbols) to correct for bleaching of unincorporated tubulin dimer. (C) The growth and shortening velocities for both the leading and lagging ends of single microtubules recorded from 18 Arabidopsis epidermal cells. Measurements consist of 3064 velocities per end representing 3.5hrs total time (~3.85s intervals, 78 microtubules). The histogram is color-coded for growth (green), pause (blue), and shortening (red) velocities. Mean velocities for growth or shortening are depicted next to the histograms. 17 MT End Lagging Leading Transitions (events / min) Kg-g 0.21 6.75 Kg-s 0.52 0.97 Kg-p 0.28 0.47 Kp-g 0.26 0.51 Kp-p 6.52 0.12 Kp-s 1.30 0.24 Ks-p 1.09 0.23 Ks-g 0.59 0.87 Ks-s 1.40 2.03 Res. & Cat. (events / sec in phase) Rescue Catastrophe 0.128s-1 0.190s-1 0.082s-1 0.043s-1 % Time in Phase 8.4% 65.3% Growth Pause 66.3% 10.1% Shorten 25.3% 24.6% Dynamicity (mm / min) 0.83±0.74 4.10±1.41 Table 1. In vivo transition rates for single microtubules. K is the rate of transitions between dynamic states in events/min. g=growth, s=shorten, p=pause. 18 Supplemental and Online Materials Plant and animal microtubule dynamics. The halftime to uorescence recovery in the plant cell cortical array is approximately four times faster (this study, 8) than for animal interphase arrays (S1). Yet, our measurements of single microtubule dynamics showed that the total subunit turnover per unit time, the dynamicity, is approximately the same between the two systems (4.9±1.6 and 4.5 ± 2.8 (S1)). Further, we found that the growth and shortening velocities of plant microtubules are actually slower than those reported for animal cells by a factor of two (S1). An examination of polymerization patterns at both microtubule ends reveals a possible explanation for these apparently conicting measurements. While plant microtubule ends grow and shorten at about half the velocity of animal microtubules (S1), they have a similar dynamicity because they exhibit dynamic behavior far more often (90% in plants vs 35% in animals (S1)). In treadmilling, plant microtubules the lagging ends also contribute to dynamicity. Plant microtubules are slowly growing and shortening almost constantly, whereas activity in animal arrays is concentrated in fewer but faster bursts. In both plant and animal interphase microtubules, subunit addition occurs primarily at the leading ends. However, in treadmilling plant microtubules subunit loss occurs not at one end, but at both ends. Because the two systems have similar rates of subunit gain and loss, distributing loss over two ends results in a larger bias towards net subunit gain at the plant leading ends. In the FRAP experiment, the shape of the recovery curve is strongly inuenced by the initial period of recovery, which is solely due to polymerization. The large bias towards polymerization at the leading end of the plant microtubules will accelerate the FRAP recovery rate when compared to the non-treadmilling animal system. The minus end of the plant microtubules does not directly affect the uorescence recovery because the slow depolymerization only eliminates bleached polymer over the duration of 19 the experiment. In sum, the FRAP experiments highlight the bias toward polymerization at the leading end of the treadmilling plant microtubule and do not suggest a dramatic difference in the actual dimer ux rates or microtubule turnover between animal and plant interphase arrays. Origin of cortical microtubules. Nucleation and tethering of microtubules at a central organizing center in animal and yeast cells creates an astral interphase array with welldened polarity. By contrast, the cells of higher plants lack a discrete microtubule organizing organelle, such as the centrosome or spindle pole body, and do not contain cytoplasmic dynein, thought to be important for gathering and tethering the minus ends. Several lines of evidence show that nucleation of plant interphase microtubules occurs at the nuclear surface (S2). We present evidence in this work for additional nucleation of interphase microtubules at the plant cell cortex. Lengthwise attachment of new, intact microtubules from the cytosol or the introduction of new microtubules to the cortical array from trans-vacuolar strands was not observed in this study. We conclude that the majority of the new microtubules in the cortex are likely born at the cortex and not transferred from other sites such as the nuclear surface. While several sites of multiple microtubule initiations were found, in no case did the minus ends remain tethered together to form a polarized, astral array. Polymers either depolymerized to extinction or were released from their cortical initiation sites. Release could conceivably occur by dissolution of the initiation complex or through cleavage near the minus end by a katanin-like protein (S3,S4). Polymer gain at the lagging end. Release of the microtubule from the initiation site resulted in a free minus end that exhibited some capacity for dimer addition. The majority of lagging end growth events was within the measurement error for the experiment and in no case did we observe a persistence of growth events leading to elongation 20 of more than a micrometer. These observations suggest that polymerization is not strongly promoted at the lagging ends and that lagging end growth does not contribute signicantly to minus end dynamics. Polarity of the cortical array. The leading ends of adjacent microtubules often were oriented in opposite directions (Movie S3), showing that cortical microtubules are not organized in a uni-polar fashion but can have opposing polarity in the same array. Homogeneous recovery of uorescence in photobleaching experiments also revealed that the organization of the plant cortical array is not highly polarized (Movie S5). This lack of polarity is observed even in cells where the microtubules are dramatically co-aligned, showing a net transverse orientation relative to the long axis of the cell. The bi-directionality of polymers in the highly ordered cortical array suggests that net polarity in the array is not required for array organization or function. Further, tethering microtubules to the cell cortex requires a mechanism that can recognize microtubules in a variety of orientations. Methods and Materials Fusions between EYFP (Clontech) and Arabidopsis tubulin isoforms were created by amplication of AtTub3A and AtTub1A from a pooled Arabidopsis cDNA library using primer pairs homologous to the rst 22 base pairs of the tubulin open reading frame and to the rst 22 base pairs of the untranslated sequence immediately following the stop codon. Methyl-dCTP replaced dCTP during amplication to block cleavage of EcoR1 and HindII sites in the amplied products. Amplied sequences were cloned into the EcoR1 and HindIII sites proximal to the 35S promoter in the pEGAD plant expression vector (S5) using palindromic double-stranded linkers (S5). All constructs were veried by sequence analysis and introduced into Arabidopsis Col 0 by Agrobacterium-mediated 21 transformation (strain GV3101) (S5). T1 transgenic plants were characterized for GFP expression and the quality of microtubule labeling. Selected plants were allowed to self-pollinate to yield T2 seed for analysis. A concern when introducing a large molecule like GFP to mark a protein complex is that the presence of the label may interfere with normal cell function. In a recent study, Rusan et al analyzed a similar GFP-alpha tubulin fusion protein in animal tissue culture cells and found that the dynamic behavior of microtubules marked by expression of the GFP fusion protein did not differ signicantly from those marked by injecting dyeconjugates of tubulin (S1). Likewise, our FRAP analysis of cortical array dynamics in Arabidopsis plants expressing GFP-tubulin agrees remarkably well with measurements made in Tradescantia cells injected with dye-conjugated tubulin. Expression of the fusion proteins in Arabidopsis plants did not result in any obvious developmental or cellular abnormalities in the plants that were analyzed, suggesting that microtubule function in these plants is normal in most important respects. However, these transgenic plants did tend to grow slightly more slowly than wildtype individuals (unpublished observations) and also displayed a modest sensitivity to the microtubule destabilizing drug oryzalin. At 175 nM, a sub-threshold dose that has no measurable effect on wildtype plants, the transgenics show approximately a 15-20% decrease in root length as compared to wildtype. There is no evidence for cell swelling at this concentration (A. Paredez, personal communication). Arabidopsis seeds were refrigerated at 4°C for 2-3 days then germinated on MurashigeSkoog (MS) agar at 23°C under constant light. At 3-4 days, seedlings were transferred to large coverslips, mounted in MS media, and stabilized by an overlying coverslip held in place with silicon vacuum grease. Most confocal images were acquired with a BioRad 1024 confocal head mounted on a Nikon TMD 200 inverted microscope equipped with 22 a 60x 1.2 n.a. water immersion objective lens. Imaging was typically performed at 3% laser power with 2-5 second intervals between images for a total duration of 3-6 minutes. FRAP experiments were performed on a Zeiss 510 confocal microscope using a 60x, 1.2 n.a. multi-immersion objective. The position of free microtubule ends was recorded by hand after image scaling and contrast enhancement with the assistance of dedicated software routines developed in the MATLAB (v6.2) computing environment. Velocities and dynamicity were determined from the original position coordinates. Transition rates and percent time in phases were calculated from velocities. Kymographs and linearization of microtubules were created in MATLAB with dedicated routines. Cells for timelapse analysis were selected along the length of the hypocotyl, from petiole insertion to the root-shoot junction. Supplemental References S1. N. M. Rusan, C. J. Fagerstrom, A. M. Yvon, P. Wadsworth, Mol Bio Cell 12, 971 (2001). S2. A.-M. Lambert et al., in The cytoskeletal basis of plant growth and form C. W. Lloyd, Ed. (Academic Press, London, 1991) pp. 199. S3. D.H. Burk, B. Liu, W.H. Morrison, Z.H. Ye, Plant Cell 13, 807 (2001). S4. T. Bouquin, O. Mattsson, H. Naested, R. Foster, J. Mundy, J. Cell Sci. 116, 791 (2003). S5. S.R. Cutler, D.W. Ehrhardt, J.S. Griftts, C.R. Somerville, PNAS 97, 3718 (2000). Movies Movies S1,S2 and S3. Time-lapsed images of Arabidopsis hypocotyl cells expressing GFP-tubulin. Evidence for the initiation of microtubules at the cell cortex (O), the 23 formation of a microtubule bundle (B), and detachment of a microtubule from the cell cortex (D) are illustrated in each cell. Examples of nearby microtubules polymerizing in opposite directions, within array are shown in Movie S3 (AP). The images in each sequence were acquired every 3.8 seconds. Movie 1 consists of 60 frames and Movies 2 and 3 consist of 100 frames each. Movie S4. Photobleaching of a line across the cortical microtubule array reveals that both single microtubules and microtubule bundles move by treadmilling (33 images acquired at 8 second intervals). Photobleaching was accomplished using 4 laser scans at 100% laser power. Movie S5. Fluorescence recovery after photobleaching (FRAP) experiments were performed using 100% laser power for 4scans in ~10µm diameter circle. 20 images were acquired at 9 second intervals. The green circle denotes the position of the bleached area. Note the recovery of uorescence in the bleached region shows no obvious spatial bias. 24