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1
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
2
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 migra-
tion 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.
3
The microtubule cytoskeleton plays a key role in plant cell morphogenesis and multicel-
lular 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 organiza-
tion, we created tubulin–green uorescent protein (GFP) fusions that permit imaging of
individual microtubules in transgenic Arabidopsis plants.
4
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 dis-
crete 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
5
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 photobleach-
ing, 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 incorpo-
rate 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.
6
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 microtu-
bules (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 corti-
cal detachment, possibly through intra-microtubule crosslinking by other proteins (18).
7
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,
8
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),
9
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 organi-
zation of the cortical array.
10
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
11
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 #DE-
FG03-90ER2001, SLS).
12
B
C
023
1.0
115.
557
.817
3.5
Tim
e (s
ec)
D FE
A
13
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).
14
E
F
A ]
]
]
Time
D
C
B
G
15
Figure 2. Microtubules associate with the cortex and move by treadmilling. (A) Marks
generated by photobleaching GFP-tubulin labeled microtubules do not move, demonstrat-
ing 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.
16
Time (sec)
T1/2 = 58.95s 0 50 100
150
200
250
300
350
0.30.40.50.60.70.80.91.01.1
Frac
tion
of F
0N
umbe
r of t
ime
inte
rval
s Leading End
Lagging End
-30-25-20-15-10-5 0 5 10 15 20
Velocity (mm/min)
100200300400
100200300400
0
v = 3.69v = 5.80
v = 1.96v = 2.78
A
B
C (2024)
17
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 fol-
lowing 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.
18
Transitions (events / min)Lagging Leading
% Time in Phase
Kg-gKg-sKg-pKp-gKp-pKp-sKs-pKs-gKs-s
GrowthPause
Shorten
0.210.520.280.266.521.301.090.591.40
8.4%66.3%
0.83±0.74 4.10±1.41
6.750.970.470.510.120.240.230.872.03
65.3%10.1%24.6%
CatastropheRescue 0.128s-1
0.190s-1 0.043s-10.082s-1
Res. & Cat. (events / sec in phase)
Dynamicity (mm / min)
25.3%
MT End
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.
19
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
20
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 well-
dened 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 dissolu-
tion 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 major-
ity 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
21
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
22
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 dye-
conjugates 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 cel-
lular 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 Murashige-
Skoog (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
23
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
24
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 per-
formed 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.
